EDITORS

SARAH A. CORBET, SCD

PROF. RICHARD WEST, ScD, FRS, FGS

DAVID STREETER, MBE, FIBIOL

JIM FLEGG, OBE, FIHORT

PROF. JONATHAN SILVERTOWN

The aim of this series is to interest the general

reader in the wildlife of Britain by recapturing

the enquiring spirit of the old naturalists.

The editors believe that the natural pride of

the British public in the native flora and fauna,

to which must be added concern for their

conservation, is best fostered by maintaining

a high standard of accuracy combined with

clarity of exposition in presenting the results

of modern scientific research.

Contents

Editors’ Preface

Author’s Foreword and Acknowledgements

1 The Nature of Galls

Part One: Virescences and Solid Galls

2 Virescences and Witches’ Brooms

3 Solid Swellings

Part Two: Open Galls

4 Erinea, Folds and Rolls

5 Pits, Blisters and Pouches

6 Big Buds, Cigars, Pineapples and Rosettes

Part Three: Closed Chambered Galls

7 Flower Head Galls

8 Galls in Stems and Roots

9 Galls in Buds, Leaves and Fruits

Part Four: Galls in Context

10 Interactions and Adaptations

11 Evolution of Galls

12 Galls and Geography

13 Galls and People

14 Galls in History

Endnotes

Glossary

Appendices

A: Gall Inducers

B: Host Plants

C: Inquilines and Enemies (parasitoids and predators) of Gall Causers

D: Other Species

References

Searchable Terms

Species Index

Subject Index

Copyright

About the Publisher

Editors’ Preface

GALLS WERE MENTIONED in an earlier volume in this series, but since then great advances have been made in their study, and now they certainly deserve a whole volume of their own. They have long been of peripheral interest to botanists and zoologists, but of central interest to a group of enthusiasts with minds flexible enough to appreciate the multiplicity of organisms of many different groups that cause them, usurp them, parasitize the gall causers, inhabit them or eat them. The recent publication of a new user-friendly key to British galls has done much to facilitate identification (Redfern, Shirley & Bloxham, 2002; a revision is ‘in press’, 2010). Now, by bringing together so much ecological, evolutionary and historical information, we hope that this book will make more biologists aware of the compelling fascination of these remarkable structures. While the focus is on British species, the book also deals with galls from other regions and considers worldwide biogeographic patterns. Galls have already contributed much to the elucidation of ecological and evolutionary questions; and in making the natural history of galls accessible to a wider audience this book will encourage naturalists to undertake further studies of galls and their associated communities, and to consider how these can help to improve our understanding of fundamental aspects of biology. We also expect the book to act as a useful reference source for all aspects of the study of galls for many years to come.

Author’s Foreword and Acknowledgements

NO PREVIOUS VOLUME IN THE New Naturalist Series has dealt with plant galls as a subject in their own right, warranting a book to themselves. However, the classic Insect Natural History by A.D. Imms, the eighth in the series and published in 1947, devoted a whole chapter to galls, a generous allocation considering the huge size of his subject. Imms noted that galls are caused by mites, nematodes and fungi as well as by insects. He concentrated on insect galls, describing their history as well as their natural history, and touched on topics about which little was known, such as communities and interrelationships between species within galls, and possible processes that cause gall development. How an organism induces a plant to produce a gall is still obscure, although knowledge of the possible processes involved has increased dramatically over the last 60 years. This new book aims to update subjects introduced by Imms and to describe new topics based on more recent information, and to bring together information scattered in a wide variety of scientific journals. Compared to insect and mite galls, bacterial and fungal galls are relatively neglected in the literature, although there is useful information in other New Naturalist volumes, particularly in Ingram & Robertson’s Plant Disease (1999) and Fungi (2005) by Spooner & Roberts.

Plant Galls is not an identification guide. Cecidologists in Britain and western Europe are well served by keys and guides published in the twentieth century and continuing into the twenty-first. This book has two approaches. First, it describes the variety of galls of different types and caused by different organisms, largely arranged from the simpler galls to the more complex (Chapters 2–9). These chapters are arranged in three groups, each with a short introduction: first, the less organised witches’ brooms, other strange growths and solid galls (Chapters 2 and 3) caused by a wide range of organisms including fungi; second, open galls (Chapters 4, 5 and 6) with permanent openings to the outside and caused by insects and mites; and third, more organised closed galls (Chapters 7, 8 and 9), most of them caused by insects. The second half of the book (Chapters 10–14) discusses the ecology of galls more generally; it considers communities of organisms within galls, the evolution and distribution of galls, and human and historical perspectives. In many chapters, some information is included in boxes. These include detail that adds to or helps to explain material in the main text but can be skipped without detracting from the main story of the chapter.

One of Imms’ main aims in 1947 was to encourage amateur naturalists to investigate galls in their own locality and to encourage studies that would add to existing scientific knowledge. This also is the aim of the present volume–interested amateurs can add to the pool of knowledge with little more equipment than a microscope, information that research biologists in university departments and museums have little time or resources to investigate themselves. I hope that professional biologists also might find interesting, perhaps new, information in this book. Extensive references are included for each chapter to allow readers to take their studies further; particularly useful may be the compilations of papers marked by an asterisk (*) in the General references, backmatter; these papers are not quoted individually.

The subject of plant galls is wide ranging, one that requires knowledge of plants as well as of the various galling organisms. Several specialists have provided me with information or have read one or more chapters and have suggested improvements: Dick Askew (ex Manchester University), Jonathan Briggs, Arthur Cain (ex Liverpool University; now deceased), Keith Harris (ex the Commonwealth Institute of Entomology), Wim Nijveldt (ex Institute of Phytopathological Research, Wageningen), Tom Preece (ex Leeds University), Peter Price (Northern Arizona University), Karsten Schönrogge (Centre for Ecology and Hydrology, Wallingford), Brian Spooner (Royal Botanic Gardens, Kew), Graham Stone (Edinburgh University) and David Wool (Tel Aviv University). I have learnt a tremendous amount as a result of their help and thank them all. Any errors that remain are, of course, entirely my own responsibility. Most members of the British Plant Gall Society are amateur cecidologists whom I hope will be interested in this book. The current Chairman of the BPGS is Tom Higginbottom, who read a draft of every chapter and made sure that unfamiliar subjects were described in an understandable way. I am very grateful to him for all of his advice. Several people have generously allowed me to use their slides and other images of galls: Robert Cameron (Sheffield University), Michael Chinery, György Csóka (Hungarian Forest Research Institute, Mátrafüred), Keith Harris, Peter Harris (Agriculture Canada, Lethbridge, Alberta), Tom Higginbottom, Conrad Labandeira (Smithsonian National Museum, Washington), Graham Stone and Robin Williams. Jacky Hodgson, librarian in charge of Sheffield University’s Special Collections, provided me with images of text and figures in Gerard’s Herball and Hooke’s Micrographia. A number of colour images and line drawings have been reproduced from Redfern, Shirley and Bloxham’s British Plant Galls (2002, 2010) with permission from the publisher, the Field Studies Council, and Michael Bloxham, the artist. The photographer, and publisher if from a published source, are acknowledged in the caption to each image and figure. I am particularly grateful to Michael Bloxham who drew all of the many line drawings in this volume, accurately and with close attention to detail, devoting a lot of time and expertise to the task. Also to my son Alexander Cameron, who compiled Fig. 244. Special thanks too are due to my husband Robert Cameron for reading everything and for making suggestions for improving the text. Finally, I thank the publishers of the New Naturalist Series and Sally Corbet (series editor, HarperCollins) for checking each chapter and suggesting improvements, and for their forbearance and patience in allowing me several times to extend the deadline for completing the manuscript. The book has taken considerably longer to write than I had anticipated at the outset.

CHAPTER 1

The Nature of Galls

MOST NATURALISTS HAVE COME across oak apples, robin’s pincushions, marble galls and witches’ brooms. These are just the most familiar examples of the strange growths that are plant galls: beautiful, often bizarre and colourful, and amazingly diverse in structure and in the organisms which cause them. They have been known since ancient times; both the ancient Greeks and the Chinese used them in herbal medicine, and in later times they had a variety of commercial uses. Ideas about their nature were generally vague (and wrong) until the great scientist Marcello Malpighi (1628–94) used microscopic studies to reveal that they were caused by other organisms using the plant as food, and modifying its structure to suit their own requirements. Even today, while subject to many detailed studies, the precise mechanisms by which the gall causer induces its plant host to form a gall are not known.

Now we do know that galls can be caused by many kinds of organism: bacteria, fungi, nematode worms, mites and insects. Galls caused by insects in particular have attracted attention as a very convenient source of information on herbivore/plant relationships and on the interaction between herbivores and their enemies. The more complex galls provide accessible microcosms of ecological communities and their interactions. With climate change and human activities, galling organisms have spread beyond their original natural ranges. Hence, galls provide endless fascination for a small, but growing, group of biologists and natural historians. This book aims to introduce the world of galls and the various organisms that cause them, to describe their variety and their history, and to illustrate how humans have interacted with them since the time of the ancient Greeks. I hope it will encourage more studies of galls and their inhabitants, by amateur cecidologists as well as professionals. There is a lot to discover.

WHAT IS A GALL?

The word ‘gall’ is derived from the Latin galla, an excrescence produced on trees, particularly on oaks. Galla became the French galle, a gall-nut or oak apple, and the verb galer means to gall, annoy, fret or tease; also to break the skin by chafing. Similar words in other European languages have the same meaning: gealla Anglo-Saxon, galla Old Saxon, gal Dutch and Galle German (see Redfern et al., 2008). ‘Cecidium’ and ‘cecidology’ derive from the Greek or , an older root than the Latin. It means something gushing forth, ooze or outgrowth and links with the production of poison and bile. This meaning extends to include the dye and ink extracted from the galls and thence to the galls themselves (Arthur Cain, pers. com.). Thus, there is a connection between galls (to the Greeks and Romans, mainly growths on oak trees) and gall, or bile, as the product of the gall bladder. The extension of the term to other growths on other plants is a later development, and raises the question: how do we define a gall?

In modern terms, galls are growths on plants formed of plant tissue but caused by other organisms. They vary in size and structure from small bumps, pustules, pimples and hairy patches to large swellings and bizarre growths, often colourful and with elaborate appendages, and nothing like any structure typically produced by the plant. All galls are the result of intimate contact between a herbivore and its host plant, the plant being induced to provide the galler with food or nutrients and with shelter from an adverse environment including, probably, protection from enemies. The host is usually a plant but may be a fungus (a mushroom or toadstool) or a lichen, or a seaweed or other alga. In order to form a gall, the affected plant cells must be young and omnipotent, able to develop into any type of tissue required by the gall causer. Such cells are meristematic, found in the growing points of plants in buds, and in the cambium of leaf veins, stems and roots. It is in these tissues that most galls originate. The galler is dependent on its host and is specific to it or to a few closely related plant species. The relationship is nearly always a parasitic one: all benefits pass to the gall causer, enabling it to grow and reproduce. In only three instances does the gall benefit the host as well as the galler; these are examples of what is known as mutualism, a symbiosis from which both parties benefit. In figs and fig wasps, the fig wasps pollinate the fig flowers as well as causing galls in some of them, so providing an essential service for the fig trees. They are the only insects that can do this. The fig wasps get food and shelter and cannot live anywhere else. In bacterial nodules on the roots of legumes, alders and a few other plants, the galling bacteria convert nitrogen gas in the atmosphere into a form that the plant can use to synthesise proteins for growth, and the bacteria get nutrients and shelter. And in mycorrhizal growths caused by fungi on plant roots, the plant supplies the fungus with products of photosynthesis while the fungus enables the plant to absorb water and nutrients more efficiently from the soil.

The key feature here is that the structure of the plant is modified by the galler in such a way as to benefit the causer, and not simply as a response to wounding. As we will see later, this is not always a simple matter to determine. Hence, galls are produced by one organism under the direction of another totally unrelated organism, both of them necessary for the unique growth that is the gall. Lichens have sometimes been considered analogous to galls; they are structures that depend on the association of two unrelated organisms (a fungus and an alga or blue-green bacterium). The two species are intimate co-inhabitants and often depend on each other for survival, and they live together in a new structure that is the lichen. The gall, in contrast, is formed by one partner under the direction of the other. This does not happen to the same extent in lichens. The fungus makes up most of the lichen and houses the alga or cyanobacterium, although sometimes the appearance of the lichen is influenced by the species of green cells that it contains. Usually in lichens, too, both partners benefit from the association whereas in nearly all galls, the gall causer is a parasite of the plant and all benefits flow only in one direction, to the parasite.

Several types of organisms cause galls. Insects cause the majority and induce the greatest variety of structures: gall midges and a few other families of flies (Diptera), gall wasps and sawflies and a few chalcid wasps (Hymenoptera), gall-causing aphids, adelgids, phylloxerans and some psylloids and scale insects (all Hemiptera), and a few beetles (Coleoptera) and moths (Lepidoptera). In some parts of the world, thrips galls (Thysanoptera) are common. Apart from insects, eriophyoid mites (Acari: Eriophyoidea) cause galls of several types, and a rotifer and some copepods cause galls on filamentous algae. After insects and mites, fungi are the commonest gall causers, with some smuts and rusts causing the most elaborate fungal galls. A few bacteria, phytoplasmas and viruses cause growths, some of them larger than any swelling caused by an insect. In addition, some plants gall their fellow plants: mistletoes and dodders in Britain and dwarf mistletoes in other parts of the world, and some of these can grow larger than their hosts. All organs of the plant may be galled by these various agents, the roots, stems, flowers, fruits and seeds, though most galls develop on leaves or in buds.

It is evident that galling has arisen independently many times in all of these groups (Chapter 11). Usually, animal, fungal or bacterial galls can easily be distinguished. Most animal galls contain chambers or cavities enclosing juveniles and/or adults, often large enough to be recognised with the naked eye or with a hand lens. Fungal galls are more or less solid but with air spaces between cells and contain hyphae that can be seen only with high magnification, but their swellings on leaves and stems covered with powdery spores when the galls are mature are characteristic. Bacterial galls are usually recognised in the field by negative attributes: they are solid, often woody, without chambers and without any inducing organism recognisable with a hand lens. But they can be confused with growths caused by physiological and developmental abnormalities of the host plant, e.g. fasciation caused by ‘errors’ in development or due to environmental factors, such as unfavourable climate or soils. Some insects and mites (and a few microorganisms) cause fasciation-type symptoms, ‘virescences’ known as phyllanthies or phyllodies, and these are considered to be galls; the gallers can usually be found between the tightly overlapping leaves. Mechanical damage and wounding, and genetic or physiological abnormalities, can cause swellings or brooming too. These are not galls, of course, but they sometimes look like galls and can be difficult to distinguish from them. Wound tissue, caused by grazing animals (caterpillars, rabbits, cattle) or by coppicing or lopping of tree trunks and branches can sometimes cause gall-like swellings too. Growths caused by gall wasps and crown gall bacteria are compared with genetic tumours in Box 1.1; gall wasp galls can be safely distinguished in the field but crown gall and genetic tumours require expert investigation to be separated with certainty. Fortunately, galls caused by gall wasps and other animals and by bacteria are usually much more common in the countryside than genetic tumours.

The plants that are galled vary across the world, dictated largely by distribution of the plants. About 98 per cent of known gallers affect flowering plants (angiosperms), with most (90 per cent) on dicotyledons (Meyer, 1987). Galling often affects the commonest and largest plant species more than others. In Europe and North America, about 50 per cent of galls occur on oaks and beeches (Fagaceae), 20 per cent on the daisy family (Asteraceae) and 15 per cent on roses, brambles and cherries etc. (Rosaceae). In South America, Africa and India, galls on legumes (Fabaceae) and acacias (Mimosaceae) predominate. In Australia, more than 50 per cent occur on eucalypts (Myrtaceae). The type of gall causer varies too, with some families commoner in temperate climates and others in tropical and subtropical areas. For example, cynipid wasps are commonest in the oak forests of warm temperate Europe and North America; galling sawflies on willows are commoner in northern temperate zones than further south; scale insects dominate on the eucalypts of Australia; and thrips and psylloid galls are common in India. Gall midge galls seem to occur everywhere although, as in all of the groups, different species occur in different parts of the world. Most gall-causing insects and mites infest dicotyledons rather than other plants, but galling fungi partly redress the balance by being regular gallers of cereals and other monocots.

BOX 1.1. Comparisons between a cynipid gall (an Andricus species on oak), crown gall (caused by Agrobacterium tumefaciens) and genetic tumours (See Bayer, in Williams, 1994)

Gall wasps (Cynipidae) cause new structures on plant organs, structures with an organised and restricted growth, usually with definite layers of plant tissues inside. All insect and other animal galls require continual stimulation from the animal to grow and develop. Bacterial crown galls and genetic tumours are generally unorganised and are unrestricted, able to continue to grow even if the initiating stimulus (the bacteria in the case of the crown gall) is no longer present. Despite this similarity, bacterial-caused growths on plants are regarded as galls because they are induced by a ‘foreign’ organism while tumours caused by genetic abnormalities of the host are not. Similarities and differences between the three types of growth on host plants are summarised here.

Induction

Cynipid gall: stimulated by oviposition by the female and by feeding of the first instar larva.

Crown gall: due to transfer of plasmid DNA from the bacterium.

Genetic tumour: genetic imbalance in host cells.

Proliferation of cells

Cynipid gall: requires continual stimulation by feeding larva.

Crown gall: automatic after initial stimulus.

Genetic tumour: automatic after initial stimulus.

Tissues affected

Cynipid gall: meristematic and young parenchyma cells.

Crown gall: meristematic cells and intercellular spaces.

Genetic tumour: meristematic cells.

Tissue growth and histology (structure of cells)

Cynipid gall: organised into distinct layers with different functions.

Crown gall: amorphous, mostly unorganised.

Genetic tumour: mostly unorganised.

Location of growth

Cynipid gall: on all host organs but most on leaves and buds.

Crown gall: on all host organs especially where wounded; often on shoots and roots.

Genetic tumour: on organs where growth is under stress.

Artificial propagation

Cynipid gall: not possible.

Crown gall: easily induced by grafting.

Genetic tumour: easily induced by grafting.

TYPES OF GALLS

Galls vary from single enlarged cells (on algae) to complex multicellular growths predictable in shape and size (on conifers and flowering plants). They vary in structure from simple pits, blisters, swellings, and leaf folds and rolls that remain open, to complex closed and highly organised structures that involve several types of tissues. Küster, in 1911, introduced a system of classification for galls that was followed during most of the twentieth century. It is cumbersome, though, and has largely been replaced by simpler and more familiar terminology. Both systems are described in Box 1.2. Categories of galls overlap considerably. Erinea may be found inside marginal rolls, deep pits can be confused with small pouches, and open pouch galls can become sealed and closed. So, categories tend to merge and overlap, and are not completely distinct.

BOX 1.2. Classification systems for galls

Küster’s classification (1911)

(Summarised in Dreger-Jauffret & Shorthouse, in Shorthouse & Rohfritsch, 1992). Küster based his classification of galls on their structure, dividing them into two major groups:

Organoid galls

Organs that are modified or multiplied but are still recognisable; examples are witches’ brooms and virescences.

Histioid galls

Growths that involve enlargement and proliferation of cells that produce a more or less new organ. Histioid galls are divided into two further categories:

(a) Kataplasmas (or cataplasmas): growths that vary in size and shape or are irregular, with tissues that are scarcely differentiated; examples are tumours and cankers.

(b) Prosoplasmas: more complex and organised growths with layers of specialised tissues, forming a structure new to the plant.

Many non-arthropod galls tend to be kataplasmic or organoid while most arthropod galls are prosoplasmic. There is overlap between all of the categories, with a large variety of transitional forms perhaps not fully appreciated in Küster’s time.

Classification based on gall shape

(Modified from Dreger-Jauffret & Shorthouse, in Shorthouse & Rohfritsch, 1992). The classification is based on the appearance of the galls; most of them belong to the histioid prosoplasmic category of Küster. This classification neglects galls of bacteria and fungi, many of which belong to Küster’s histioid kataplasmic group.

Filz galls: patches or tufts of hairs, usually on leaf blades; all are caused by eriophyoid mites and are more often known as erinea.

Pit galls and blister galls: doming of one side of a leaf blade with a depression (or pit) beneath; the lower surface may be covered by epidermis, forming a blister; caused by gall midges, psylloids and scale insects.

Pouch galls: a hollow invagination of the leaf blade forming a pouch on one side of the leaf with an opening remaining on the other side; caused by eriophyoids, cecidomyiids, aphids and psylloids.

Roll and fold galls: roll or fold of leaf margin or of most of the blade; caused by eriophyoids, cecidomyiids, aphids, psylloids, sawflies and thrips.

Covering galls: the gall causer at first feeds on the surface of a leaf or stem and becomes completely enclosed by plant tissues growing up around it; caused by aphids, scale insects and cecidomyiids.

Mark galls: the egg is deposited inside plant tissue or the newly hatched larva burrows inside, embedded in plant tissue from the start; caused by cynipids, cecidomyiids and tephritids. Mark is a German word; in this context, its most useful meaning is ‘bone marrow or pith’ mark galls of cecidomyiids and tephritids are often deep inside the pith of stems.

Covering and mark galls include the most complex galls and can only be distinguished if their mode of development is known. The end result is often similar and this reduces the usefulness of the terms. Nowadays, these terms are rarely used, replaced by words that describe the appearance of the galls, e.g. ball, marble, spindle or spangle galls; or hedgehog or spiny galls.

Bud and rosette galls: these range in complexity from enlargements of buds (big buds) to clusters of modified leaves forming rosettes, artichokes and pineapples; caused by eriophyoids, cecidomyiids and adelgids.

It is worth saying here that gall formation can also merge with other herbivorous lifestyles. Many families of insects have galling and non-galling species within them; a good example involves the sawflies. Most sawfly larvae are specific to particular host plants and most are free-living herbivorous caterpillars that graze leaves. Some cause the leaf to fold or roll, which gives a degree of protection to the larva feeding inside. This is regarded as a gall, a simple one, if the roll or fold is thickened, even if the larva feeds outside the roll from time to time. The Phyllocolpa sawflies rolling and folding willow leaves come into this category. Other genera of sawflies (Pontania, Eupontania and Euura) cause more elaborate bean-and pea-shaped swellings or swollen buds that are unambiguous galls.

There are other insects that cause distorted crumpled leaves that are not usually categorised as galls. The unthickened crumpled young leaves common in spring on blackthorn and other Prunus species and caused by aphids are examples, as is the leaf distortion caused by the cuckoo-spit froghopper Philaenus spumarius, which has a wide variety of effects on a large range of host plants. There are other species that cause slight distortions that are regarded as galls by some authorities but not others. The effects on the plant of these species are not known well enough to determine whether they are true galls or not. Most leaf mines and stem borings are not regarded as galls, the miners and borers simply tunnelling in the thickness of the leaf or stem without causing swellings. But a few cause thickened tissue that the larvae feed on, and these are regarded as simple galls (e.g. some leaf-mining moths of the family Nepticulidae). Conversely, the mine of the holly leaf miner Phytomyza ilicis is not regarded as a gall even though tissues in the mine are distinctly thickened. But the larva inside the mine does not feed on this extra tissue; it is wound callus only.

Most plant parasitic fungi, moulds and mildews and so forth, do not cause galls. But some do; some rusts and smuts cause swollen cushions on leaves and stems and some of their growths are elaborate (e.g. the galls of Gymnosporangium and Exobasidium species).

This ambiguity (when is a gall not a gall?) can cause problems of identification in some cases, but it also shows us the variety of ways in which herbivores adapt to feeding on plants and protecting themselves from enemies and adverse climate. Furthermore, variation in the complexity of galls caused by a single group, for example the sawflies, gives us clues to the way in which gall-forming has evolved.

The first part of this book deals with galls and their causers in a sequence from the structurally simple (Chapters 2 and 3) to the most complex (Chapters 7, 8 and 9). Chapters 2 and 3 describe galls with no constant shape or structure, on a plant part that is more or less still recognisable though often much distorted. These ‘indeterminate’ galls include witches’ brooms and other strange growths (Virescences, Chapter 2) and solid lumps of various shapes and sizes (Solid Galls, Chapter 3). A few of these galls are caused by insects and mites but most of the gallers are fungi, bacteria and viruses, and the galls are simpler than most insect and mite galls. The best-known virus gall is due to the wound tumour virus, and crown gall and root nodules are the most familiar of the bacterial galls. Unlike galls caused by most fungi, nematodes, mites and insects, the wound tumour virus and crown gall develop on a variety of plants; they are not host-specific. Fungi cause swellings on leaves, stems, roots, flowers and fruits; some species are not nowadays classified as fungi (but are still studied by mycologists). Some gall-causing fungi cause diseases, like clubroot of brassicas, potato wart disease, ergotism and black stem rust of wheat; these were devastating until their lifecycles were understood and they were brought under control (their lifecycles are described in Chapter 3 and the economic effects of some of them in Chapter 13).

Galls caused by animals, insects, mites and nematodes, are very varied and have a more complex structure than fungal galls but do not damage their host plants to any great extent. Their galls are described in Chapters 4–9 in a series of ever-increasing complexity. ‘Open’ galls have permanent apertures through which gallers can push their way out (though, also, enemies can enter). These include erinea (hairy patches inhabited by mites), leaf folds and rolls (Chapter 4), pits, blisters and pouches (Chapter 5) and, the most elaborate, cigars, pineapples, rosettes and artichokes (Chapter 6). ‘Closed’ galls are sealed and include the structurally most complex galls. Organisms inside these must chew their way out, or push their way through a specially prepared weak spot, or wait until the gall rots on the ground before they can escape. Those in flower heads of thistles and knapweeds and in fig fruits are described in Chapter 7, galls in stems and roots in Chapter 8, and galls in leaf buds and on leaves make up Chapter 9. The structure of the galls is described and all of the chapters include examples of the lifecycles of typical gall causers, of insects and mites and of a few gall-causing nematodes.

HOW ARE GALLS FORMED?

One topic common to all parts of this book is not dealt with directly elsewhere. Understanding how galls develop is a major problem that has exercised cecidologists for years. The gall causer redirects normal growth and development of the plant, the details probably varying in different gall-causing groups. In bacterial galls the processes are fairly well understood (Box 2.2 and Chapter 3) but not so in animal galls; in these, several ‘morphogens’ have been suggested but none have been shown conclusively to cause galls. The gall causer manipulates the plant to ensure an adequate, reliable food supply and also to protect itself from adverse environmental conditions including unfavourable climate and predators. How this happens is the subject of active research today, though not much progress has been made over the last 25 years. But as knowledge of plant development increases and as techniques improve, the molecules involved in gall formation and the genes that trigger their production perhaps will be identified and lead to a greater understanding of the processes involved. Even the processes involved in normal development are incompletely understood; this is the developing field of ‘Evo-Devo’, the way in which genes switch on and off to direct development.

We do know a little. These processes differ in animal-caused galls and those caused by fungi. Galling fungi cause plant cells to increase in size and perhaps also in number, but there is no development of the elaborate structure and differentiation into layers of specialised tissues as there is in many insect galls. Gall wasps and gall midges cause the most elaborate galls, and galls of other insects and some mites and nematodes also develop distinct tissues. The structure of galls as they grow is known for many groups of insects and for some mites and nematodes and, in nearly all of them, a live feeding larva is necessary for complete development. The anatomy and morphology of the galls of different groups, described in Chapters 2–9, shows similarities between groups, although it is more complex in some galls than in others. Most animal galls develop patches or a layer of nutritive cells, rich in soluble compounds (glucose, sucrose, amino acids, etc.) that the larva needs, and rich in enzymes needed to convert storage compounds like starch into sugars to ensure a continuous food supply. The nutritive tissue and the actively feeding larva act as a ‘nutrient sink’, drawing nutrients from elsewhere in the plant through vascular elements that are more abundant than in the ungalled plant. Most galls develop in parts of the plant that are nutrient sinks anyway, such as the meristems of leaf and flower buds, the cambium of young stems and roots, and developing fruits and seeds. The galler enhances this natural transport system of plants, turning it to its own advantage and making sure that the sink effect lasts until it is full-grown. The gall also protects the gall causer, and often a layer of woody cells develops outside the nutritive zone, protecting it and the larva within. Gall growth, therefore, coincides with and enhances an active growth period of the host plant, when it is young and growing rapidly, or when trees flush a new crop of leaves in spring, or when flowers and fruits develop.

Cells of meristems are undifferentiated and omnipotent (or totipotent); they can potentially develop into any type of plant tissue. Sometimes the gall causer induces older cells to de-differentiate, to lose characteristics they have already developed and to revert to a more juvenile state. They then can develop new tissues that benefit the galler. Galls develop through three main phases: initiation, growth and maturation. The initiation phase is specific to a particular gall causer and affects a small group of cells, or perhaps just one cell, and this develops into all or most of the cells in the future gall. Usually, only the newly hatched larva has the ability to initiate a gall although, in some species, feeding or oviposition by the parent female starts the process. This phase is the most critical and must be precisely synchronised with growth and development of the plant. Usually there is only a narrow ‘window’ during which the plant can be manipulated by the galler; lack of synchrony is perhaps the main reason that the number of galls in a population varies so much from year to year.

The second phase is growth and development when the gall increases dramatically in size, and depends on actively feeding larvae. Cells grow and multiply forming nutritive tissue around the larva, vascular elements arise and grow to join existing veins, and a protective woody layer may develop. Information on the nutritive tissue and possible processes causing gall development is given in Box 1.3. The gall grows to its full size and shape, perhaps adorned with hairs or spines or other ornamentation on the outside, and its colour may change as it matures. It then enters the third or maturation phase and the larvae become fully fed. By this stage, the nutritive cells have all been eaten or are collapsed and no longer supplied by nutrients from the vascular bundles, and the nutritive tissue no longer acts as a nutrient sink. The larva may now fill all of the space inside the woody shell. Full-grown larvae may overwinter in the gall, pupate and emerge as adults next spring, or larvae may escape from the gall to pupate in the soil.

We also know that even if the process of gall induction is well synchronised with growth of the plant, a gall may not develop successfully; the plant can ‘fight back’. Some plant individuals are more resistant to gall formation than others and larvae attacking them die within a few days as nutritive tissue fails to develop. Either the plant does not react to the stimulus from the larva, or the plant produces a toxin that kills the larva. Differences in resistant and susceptible plant individuals help to explain why some plants of a species are regularly galled while close neighbours are not. From a plant perspective gall causers are a disease and, as with hosts for all diseases and parasites, evolution promotes resistance. The unique feature of at least some gallers is the extent to which they can not only overcome such resistance, but oblige the plant to modify its development to suit them.

BOX 1.3. Development of galls: the possible processes involved

It is generally accepted that the galling organism, insect, mite, nematode, fungus or bacterium, causes the gall to develop even though it is made entirely of plant tissue. The key process in development of most insect galls, and probably of galls of mites and nematodes as well (though not of bacteria or fungi), is the formation of nutritive cells. The structure of nutritive tissue is characteristic but the processes that cause it to develop are obscure.

Eggs of gall insects are either inserted into plant tissue, or are laid on the outside of the plant and the larvae crawl into the place that will develop into a gall. As soon as the larva hatches or reaches the appropriate spot and starts to feed, nutritive cells develop around its head. These cells are distinctive. The nucleus and nucleolus enlarge, ribosomes and other organelles multiply, chloroplasts become modified, sugars, lipids, amino acids and proteins become more concentrated, and the cytoplasm becomes dense as the cell vacuole fragments or disappears. The cells become brick-shaped and tightly packed as intercellular spaces disappear, and the cells multiply to form a nutritive patch around the head of the larva or a complete layer of tissue enclosing it. Outside this layer, in cynipid galls at least, a layer of storage nutritive tissue develops. This is rich in starch, which is converted to sugars as required by the growing larva. Vascular strands develop outside the nutritive layer to grow and link with the vascular system of the host organ, the bud, leaf, stem or root and, as the larva feeds, nutrients flow from nearby ungalled parts of the plant to the nutritive tissue, forming a nutrient sink that lasts as long as the larva is actively feeding. Similar nutritive cells appear in many insect galls although, in cynipid galls, the cells are larger than in most other galls. In galls of some sucking insects, e.g. some aphids, psylloids and scale insects, nutritive tissue does not develop; the long proboscis is inserted deep into the plant and taps the vascular bundles directly. But the more elaborate galls of these groups, e.g. of adelgids, do develop nutritive tissues. Sawfly galls, and those of moths, beetles and some flies, also do not develop a characteristic nutritive tissue. As the young sawfly caterpillar feeds it wounds nearby cells and this stimulates more cells to grow so ensuring its food supply, but the cells do not develop the characteristics of typical nutritive cells.

In bacterial galls, such as the root nodules caused by Rhizobium on legumes, formation of the nodule is initiated by ‘nodulating factors’ (Nod factors). Some Nod factors are oligosaccharides that are involved generally in cell proliferation and growth and development. In Rhizobium, one Nod factor is NodC (Schönrogge et al., in Csóka et al., 1998), an enzyme that causes the plant to develop an infection thread through which the rhizobia enter the plant (described further in Chapter 3), and these stimulate development of the nodule. It seems that NodC isolates a group of young cells in the root and prevents them from developing normally–the first step in production of the gall. Substances similar to NodC have been found in a wide range of organisms (in mice, a toad, yeasts) and are involved in development in all of them. They also occur in the cynipid galls of Diplolepis spinosa (on a rose, Rosa rugosa) and of the oak gall wasp Andricus quercuscalicis although it is not known what role they may have. In Rhizobium galls, the Nod factors produce a whole cascade of developmental effects before a nodule develops. Development of the endosperm of seeds involves a similar cascade of processes. Endosperm is tissue in the seed that supplies nutrients to the germinating seedling before it develops its first green leaves, and the structure of endosperm cells is similar to nutritive tissue; its development may involve a similar series of processes as in galls.

The formation of a gall may not involve any novel developmental pathways in the plant. The gall causer may tap into existing metabolic processes, modifying them and perhaps extending their period of influence. The normal development of a plant, germination of seeds, growth of the seedling, flushing of leaves in spring and growth of flowers and fruits, are all stimulated and controlled by plant hormones, auxins, cytokinins and other substances that direct growth and cause young undifferentiated cells of meristems to develop. As a result, the plant develops its organs at the right time and in the correct place. The galler modifies these processes perhaps by producing plant hormones itself, or by accumulating, modifying and redirecting hormones produced by the plant. The galler might initially acquire hormones or their precursor molecules from the plant and then inject them back into the plant. The auxin IAA (indole acetic acid) is a common hormone involved in normal plant development and has been found in young larvae of Cynips quercusfoliae and Eurosta solidaginis. C. quercusfoliae causes the cherry gall on oak leaves and E. solidaginis causes stem galls in goldenrod Solidago. Tryptophan, a precursor of IAA, is present in the chestnut gall wasp Dryocosmus kuriphilus. Cytokinins are also involved in normal plant development and have been found in galls and in larvae of some cynipids, tephritids and chalcids.

Gall formation is not understood in any animal gall. It is a complex process and probably involves many substances, plant hormones and perhaps RNA molecules, viruses or virus-like particles or phenolic compounds (tannins and other molecules that are involved in general plant defence processes and are highly concentrated in the outer layers of many cynipid galls). And details of the galling process may be different in different groups of gall causers. Harper et al. (in Stone et al., in press) summarise current knowledge on the possible processes involved in different groups of insects and in gall mites.

WHY GALL?

Gall-causing, at least by animals, would seem to have considerable advantages over other types of herbivory. Once inside the plant, an adequate food and water supply is assured, the animal need not move and, like leaf miners and stem borers, gallers are protected from extremes of climate and from many enemies outside. But there are disadvantages too. Gall causers need to be small, able to fit inside a growth on a leaf or in a stem, and the galling species tend to be amongst the smallest within a family of insects. Smallness limits the number of eggs a female can produce and so limits the rate of population growth; and the relatively modest numbers dispersing to form new galls in each generation are at risk from predators and accidents. Gall causers must also be specific to their host plants, to one species or to a few closely related species. This is necessary if galling is to succeed. The galler must manipulate the plant at precisely the correct stage of its development and if it is suitably synchronised with one host species it is unlikely to be well synchronised with another. The galler must be able to withstand or overcome the defences of the plant too, and a mechanism that works against one host species may not be successful against another. A gall species cannot immediately turn to an alternative food source if a catastrophe wipes out its host.

Gall-causing is a successful strategy but, it seems, only for a minority of herbivorous organisms. A recent estimate for the number of gall-causing insect and mite species in the world is 133,000 (Espirito-Santo & Fernandes, 2007), 6 or 7 per cent of a total world fauna of perhaps 2 million insects and mites. Both figures, the gallers and the total world fauna, are rough estimates only, little more than enlightened guesswork. This is the best we can do until more areas of the world have been adequately surveyed; the tropics especially are poorly worked and it is here that species richness of insects and mites is likely to be the highest. Gall-causing is probably as common amongst tropical insects and mites as elsewhere, and so a realistic world total must be considerably higher than 133,000. Even less is known about the abundance of gall-causing nematodes, fungi, bacteria and so on, although the proportion of gallers amongst these groups is probably considerably less than it is for insects and mites. Clearly, though, galling is only one of several herbivore strategies, and not the most common. Nevertheless, it has evolved many times in different organisms. In some it is an ancient lifestyle. The oldest fossil of a gall found so far is about 300 million years old in the leaf stalk of a tree-fern and was caused by a chewing insect, the larva of a sawfly perhaps, or a beetle. In aphids, which suck juices from plants, galling occurs in the more ancient families and subfamilies that appeared perhaps 200 million years ago. Although no fossil aphid galls of this age have been found, fossils of this sort of age of fronds of tree-ferns with sucking punctures are not unusual. Other fossil galls of tree-ferns perhaps 290 million years old may be caused by fungi. But these very ancient galls are rare; it is likely that galls did not become common until 150 million years later when flowering plants and insects were diversifying rapidly. The most complex galls, for example of gall wasps and gall midges, represent a more recent lifestyle and appear in the more recent members of these groups; fossil galls of gall wasps are not known before the Late Tertiary, perhaps 20 million years ago, well after the first gall wasps appeared. Although gall-causing is a specialised way of life it is not a dead end. Gallers are adaptable, it seems, and galling species can evolve different lifestyles, sometimes even losing their galling ability (see Chapter 11 for more detail).

These issues and others are dealt with in the second part of this book. Chapter 10 explores the interactions between plants, gallers and their enemies and lodgers (inquilines) and other ‘hangers on’. In some cases these are amazingly precisely adapted, and in others there are complex communities centred round the gall-causing species. The evolution of galls and galling, including the results of recent studies with new molecular techniques, is considered in Chapter 11. A consequence of the repeated evolution of galling as a way of life is that there are pronounced geographical patterns in the kinds of animal that have evolved galling, and also in the plants they affect. These patterns are examined in Chapter 12. Chapter 13 looks at the use humans have made of galls, and at some of the other impacts galls have had on human activities through their effects on crops and other plants. The final Chapter (14) traces the history of cecidology from the first recorded writings to the present day.

PART ONE

Virescences and Solid Galls

Chapters 2 and 3 form a pair whose galls are distinct from those described later (in Chapters 4–9) largely for negative reasons: they do not have a constant size or shape, they do not contain chambers, and they often do not contain any organisms inside–at least, nothing that can be seen with a hand lens. These galls do not have a distinct structure and most do not have any specialised tissues, and they involve parts of plants that are still recognisable despite being considerably modified and distorted.

‘Virescences’ are the subject of Chapter 2. They appear as oddly shaped parts of plants, where shoots, roots, leaves or flower parts are distorted, multiplied or bunched together. On trees and shrubs, these are ‘witches’ brooms’ on non-woody hosts, ‘chloranthy’, ‘phyllanthy’ and ‘phyllody’ are common descriptive terms. Other oddities are included as virescences, for example, the single leaf or berry that has a strange shape amongst the normal ones. In Chapter 3, solid swellings, without regular chambers and cavities, are described. These vary hugely in size, from single enlarged cells invisible to the naked eye to growths a metre or more across. For both virescences and solid galls, the effect on the host plant may be slight and trivial or considerable, and may even be serious enough to kill the host.

The galls in Chapters 2 and 3 are caused by a wide range of organisms: viruses, phytoplasmas, bacteria, parasitic plants, fungi, nematodes, rotifers, mites and insects but, unlike Chapters 4–9, it is the non-animal groups that are the major players here. Many are microscopic in size, although the galls they produce may be huge. Others are large, sometimes nearly as big as the hosts they parasitise. Fungi are measured in micrometres (µm) or millimetres (mm; 1 mm = 1,000 µm), bacteria are ten times smaller, phytoplasmas ten times smaller than bacteria, and viruses 100 or 1,000 or more times smaller than this. Parasitic plants, measured in centimetres and metres, go to the other extreme and include the largest of all gall causers. Galling is the exception rather than the rule in all of these groups; most of their relatives that depend on a living host are straightforward parasites. Gall-causing bacteria and parasitic plants tend not to be host-specific and can survive on a wide variety of host plants. Fungi, mites and insects are usually restricted to one host species or to a few in the same genus. Little is known about the other groups and their host preferences.

CHAPTER 2

Virescences and Witches’ Brooms

MOST VIRESCENCES ARE BUSHY growths of distorted and multiplied parts of plants, either greened and leafy or woody and twiggy. Those on herbaceous plants are variously known as chloranthies, phyllanthies or phyllodies, while on trees and shrubs they are witches’ brooms. Other distortions, also included as virescences, are less common: odd-shaped leaves or fruits or bunches of roots in strange places. In all, the plant part is recognisable even though it no longer performs its usual function efficiently.

Virescence is a cumbersome and unfamiliar word, derived from the Latin virescens = becoming green. It should not be confused with virus or virosus, also Latin words, which mean stinking or poisonous, and have been adopted into the English language as viruses–even though some viruses cause virescences! Greening of petals, stamens and ovaries that normally are not green are included as virescences, though the more specific term chloranthy is often used (chloranthus is Latinised Greek meaning ‘green-flowered’). These plant parts are multiplied too, and often are smaller than normal, bunching together to form the virescence. If leaves, normally already green, are involved, the terms phyllanthy or phyllody are often used (phyll- or -phyllus, as a prefix or a suffix, is Latinised Greek, meaning ‘relating to leaves’), but the three terms are often used more-or-less interchangeably. Included as virescences are witches’ brooms: untidy, twiggy bunches of stems, leaves or roots that form bushy growths on trees and shrubs, and can grow very large. Like the smaller virescences on herbaceous plants, these include odd-shaped leaves, flowers or roots that are often reduced in size. The French verb virer seems appropriate and has a broader meaning than the Latin virescens: besides meaning ‘to turn colour’, it means ‘to twist, to turn about’. The individual stems and leaves of virescences are often twisted and distorted. Other odd-shaped growths of recognisable plant parts are included as virescences too. ‘Fasciations’ are easy to confuse with virescences and may be indistinguishable from them; they involve multiplied stems or flower heads that remain fused together, and are probably due to a genetic or physiological error–but these are not caused by another organism and so are not classed as galls.

Virescences are caused by microorganisms, viruses, phytoplasmas, bacteria and fungi, by parasitic plants, and by a few mites and insects. Background information on the biology of the non-animal galling groups, which cause the majority of galls in this chapter and in Chapter 3, is given below and in Boxes 2.1–2.5 (equivalent information for mites and insects is given in Chapters 4 and 7). Examples of virescences caused by each group follow and make up the bulk of the chapter.

VIRUSES

The general biology and classification of plant viruses is given in Box 2.1. Many viruses cause serious diseases of plants, particularly of cereal crops, and some are responsible for major economic losses. Most are known by the symptoms they cause, and their names are usually abbreviated to letters, e.g. TMV, tobacco mosaic virus; MSV, maize streak virus. Many virus diseases are described as ‘yellows’ or ‘mosaics’, or cause dwarfing or stunting of plants, or leaf-rolling and necrosis (blackened patches of dead cells), but do not produce galls, even under the plant pathologist’s definition: an overgrowth with local proliferation of cells without tissue differentiation (see What is a Gall?, Chapter 1, for the definition of a gall used in this book). The best-known virus gall is caused by the wound tumour virus (WTV), which causes crown gall-like growths on a wide range of unrelated host plants (see Chapter 3). Others cause thickening of shoots or leaf veins, e.g. cocoa swollen shoot virus (CSSV), which is spread by scale insects and has caused serious crop losses in West Africa, and lettuce big vein virus (LBVV), spread by the parasitic fungus Olpidium. No virus is known for certain to cause witches’ brooms or other virescences, although some are implicated, e.g. the grossly distorted catkins of willows (see below). Proving without doubt that a distortion is caused by a virus is difficult, and requires the following three steps:

Virus particles must be shown to occur in cells of the host.

The virus must be isolated as a pure extract.

It must be inoculated into a healthy host plant and symptoms characteristic of the gall must appear.

BOX 2.1. Biology of plant viruses (Information from Lucas, 1998)

All viruses are obligate parasites, completely dependent on their host and capable of surviving outside a host cell only for a very short time. Parasitism by viruses is unlike that of any other organism in that the virus is incorporated into the metabolism of the host cell, programming it to synthesise more virus. Virus structure is simple–a single or double strand of nucleic acid (either deoxyribonucleic acid DNA or ribonucleic acid RNA) wrapped in a protective protein coat, the capsid (Fig. 1). Only the nucleic acid enters the host cell; the capsid is discarded outside and plays no part in the functioning of the parasite in the cell. Thus, although they have their own genetic material, they require the biochemical resources of cells of another organism to manufacture new nucleic acids, proteins and other essentials of life. Five classes of plant viruses have been described (Table 2.1), based on the type of nucleic acid present, on whether it is single- or double-stranded, and whether the order of nucleotides on the strand is the same as or complementary to the messenger RNA of the host cell (mRNA, a molecule essential in the synthesis of proteins in cells of all organisms; if the strand is the same, it can act directly as mRNA and cut out a step in protein synthesis, and therefore speed up replication of more virus particles).

FIG 1. A particle of tobacco mosaic virus (TMV) showing helical arrangement of the coat protein units around a coiled strand of RNA (after Lucas, 1998).

TABLE 2.1. Classification of plant viruses based on the type of nucleic acid present (DNA or RNA), whether it is double-(ds) or single-(ss) stranded, and whether the strand is the same as (+) or complementary to (–) the host’s messenger RNA. The related viroids are included (Simplified after Lucas, 1998)

The best-known gall-causing virus, wound tumour virus (WTV), belongs to Class III with double-stranded RNA. Because sometimes its RNA structure is the same as mRNA of its host’s cell (+) and sometimes complementary (–) to it, it exists as several strains. Most plant viruses, including tobacco mosaic virus with a single strand of RNA, the first to be discovered, belong to Class IV. Viroids differ from viruses in the size of their RNA molecule (about ten times smaller than that of a virus) and their lack of a protein coat. They have been found only in plant cells (no animal viroids have been discovered) and may be of recent origin. None is known to cause a gall.

Most plant virus particles live in the phloem of plants. They can move from cell to cell via plasmodesmata (strands of cytoplasm linking cells via tunnels through their cell walls) or be transported rapidly around the plant in the sieve tubes along with the products of photosynthesis. They tend not to occur in those parts of plants without phloem, such as the meristems of shoots and roots (so, virus-free plants can be propagated from meristems). Because they cannot survive outside the host cell and do not produce spores or any other kind of propagule that could be spread by wind or water, most plant viruses require vectors to enable them to spread from one host to the next. Common vectors are aphids, leaf hoppers and other plant bugs (Hemiptera) which suck sap from the phloem sieve tubes. The bugs take in virus during one meal and inject it into another plant during the next and, in some types, the virus multiplies in the insect as well as in the plant host. The insect then is infective for life and may even transmit the virus to its offspring via the eggs. A few viruses are spread in the soil by nematodes or parasitic fungi, plants becoming infected via their roots. Table 2.2 summarises some characteristics of virus vectors.

The parasitic flowering plant dodder (Cuscuta) is capable of transmitting viruses under experimental conditions, but is not known to do so in the wild. Other routes, exploited by some viruses of fruit and vegetable crops, are via pollen or seeds. These are efficient methods of spread for the virus but, fortunately for the host plant, seem to be rare and none of these is a gall causer. Viruses can also be spread to new host individuals by humans, entering wounds caused by grafting, and transported in plant debris carried on boots and tools.

Viruses are the ultimate parasites, manipulating the functioning of the host cell so that it manufactures more virus. An important property of all organisms, including viruses, is their ability to change, to evolve new varieties able to cope with varying conditions. Sexual reproduction is the main mechanism in animals, plants and fungi, and cytoplasmic factors (plasmids) in bacteria (see Box 2.2). Neither of these mechanisms is available to viruses; they do not contain plasmids and do not indulge in sex. Some variation is caused by mutations in their nucleic acids, occurring at much the same rates as in other organisms (mutations are usually rare). Their main source of variation seems to be RNA recombination, occurring when pieces of RNA split off and recombine in a variety of configurations.

TABLE 2.2. Some characteristics of virus vectors (simplified after Lucas, 1998)

Viruses do occur in the cells of these distorted willow catkins but, because the vital third step has not been carried out, it is not possible to be certain that they are the cause of the gall.

BACTERIA AND PHYTOPLASMAS

Background information on these organisms is given in Box 2.2. Both groups are considered together because they are both prokaryotes (see the Box), although they are not closely related. Most bacteria are not damaging to plants. Many species are beneficial, free-living in soil and water, and they perform many roles essential for ecosystems to function (concerned with decomposition and the recycling of nutrients, for example). Others are parasitic in plants (in animals, too) but may be benign or beneficial to their hosts, and relatively few are pathogenic. Most of the pathogenic species cause diseases variously described as wilts, blights, rots, leaf spots and scabs, which would not be classed as galls–few bacteria, and phytoplasmas too, cause true galls. Phytoplasmas are smaller than bacteria and are less well known. Probably all are parasitic and, like viruses, probably cannot survive outside their host cells.

BOX 2.2. Biology of bacteria and phytoplasmas (Information from Maramorosch & Raychaudhuri, 1981; Lucas, 1998)

Bacteria and phytoplasmas are ‘prokaryotes’, their cells lacking true nuclei and other membrane-bound organelles. Most plant pathogenic species are single cells, smaller than cells of ‘eukaryotes’ and with a simple structure (all other organisms apart from viruses are eukaryotes). The earliest forms of life on Earth were prokaryotes. They evolved perhaps 3500 million years ago, and dominated life for 1,500 or 2,000 million years before eukaryotes and multicellular life appeared. Most existing prokaryotes are true bacteria, with a rigid cell wall and usually with flagella. Phytoplasmas (also known as mycoplasmas or mycoplasma-like organisms, MLOs) are smaller, about half the size of the smallest bacteria, and without a cell wall or flagella. Bacteria and phytoplasmas can be cultured in the laboratory on artificial media outside cells (unlike viruses) and many true bacteria survive free-living in the wild; phytoplasmas, though, probably cannot survive and replicate outside host cells in natural conditions.

The genetic material (the DNA) of a prokaryote cell is coiled into a long chromosome and, generally, there are also one or more rings of DNA called ‘plasmids’, lying loose in the cytoplasm. True bacteria that are pathogenic invade the host by secreting enzymes that degrade the middle lamella (the layer that binds cells together), enabling them to spread between the cells. The bacteria absorb nutrients released by this digestion, allowing them to grow and multiply. Some species gain access to the xylem vessels, which allows them to spread rapidly through the plant and sometimes to cause death. Phytoplasmas behave more like viruses and, for this reason, they were thought to be similar and were classified with them until 1967. They cause symptoms similar to virus diseases and, like viruses, infect the phloem of vascular tissue (see Box 2.1), and are spread from host to host by sucking insects and by grafting. They infect the sieve tubes and can block them or cause necrosis (scarring), and adjacent parenchyma cells may enlarge and divide abnormally, so causing gall-like growths. This can disrupt the transport of nutrients around the plant leading to death of twigs and branches, and even of whole trees.

FIG 2. Processes involved in induction of crown gall tumours by Agrobacterium tumefaciens (modified after Melchers & Hooykaas 1987, in Lucas, 1998). For explanation and interpretation, see text.

The best-known bacterial galls are caused by crown gall and root nodule bacteria. Unlike other true bacteria, these species do not degrade the middle lamella between cells or destroy host tissues, and nor do their daughter cells separate as they multiply. Instead, they congregate into colonies that can be very large, causing substantial growths. These bacteria grow biotrophically, i.e. without killing the host’s cells, and are dependent on a healthy host in order to complete their development.

In many bacteria, the genes that enable the pathogen to enter the host cell are located on a plasmid. The process of invasion has been studied intensively in the crown gall bacterium, Agrobacterium tumefaciens, and is summarised in Fig. 2. Crown gall bacteria contain large plasmids which are tumour-inducing (Ti plasmids). The bacterium attaches itself to the wall of a cell of the host individual. This damages the cell, causing it to produce wound chemicals (e.g. the phenol, acetosyringone), which activate a set of genes on the plasmid in the vir (virulence) region. The vir genes cause a segment of the plasmid to be cut out (the T region) and to be inserted into the DNA in the nucleus of the host cell. The plant cell then starts producing messenger RNA (mRNA; these are molecules essential in the production of proteins in all cells) containing the new information from the bacterium, and this leads to the synthesis of two groups of chemicals:

(i) The hormones, auxins and cytokinins, which stimulate cell division and cause tumour formation; these are the same hormones that regulate normal growth of the plant.

(ii) Opines, unusual amino acids, which cannot be used by the plant but which serve as a nutrient source for the bacterium.

The pathogen, therefore, is switched on by signals from the plant itself to act as a ‘genetic engineer’, inducing the host cell to produce a tumour and to provide nutrients which only this bacterium can use. This mechanism has led to gene manipulation in plants by scientists, bacterial plasmids being used to carry genes from the nucleus of one plant to another, so introducing desirable characteristics into the new host, such as resistance to viruses or to insects and herbicides.

Reproduction in bacteria is normally simple and asexual, the parent cell splitting into two daughter cells that separate and live independently. The crown gall and root nodule species are unusual in that the daughter cells congregate together. Phytoplasmas, like viruses, cannot survive outside their host cells and so need vectors to transmit them to new hosts. Bacteria, though, can survive outside the host as resistant spores, and these can be dispersed widely by wind and water and so be spread to new hosts.

The best-known galls caused by bacteria are crown gall, caused by Agrobacterium tumefaciens, and Rhizobium root nodules (both are described further in Chapter 3). Crown gall is notorious and is one of the best known of all plant diseases; the study of its bacterium has led to advances in knowledge in many branches of biology (e.g. biochemistry, genetics, physiology, evolution and ecology). Related to Agrobacterium is Rhizobium, the cause of the root nodule galls common on peas, beans and other legumes (Fabaceae). This gall is one of the few in which both host and gall causer benefit from the association. The usual gall interaction is one-sided, the gall causer acting as a parasite and gaining most or all of the advantage, with the host suffering some degree of disadvantage. A mutually beneficial association also exists between the actinomycete bacteria Frankia (not related to Rhizobium) and their host plants, e.g. alder Alnus glutinosa. Another well-known example of benefit on both sides is the association of fig wasps and figs (see Chapter 7, Table 7.1.). Most bacterial galls are swellings and are included in Chapter 3. The few that cause virescences are described below–phytoplasma witches’ brooms and phyllodies are probably more widespread than those of bacteria.

FUNGI

Information on the general biology and classification of fungi is included in Box 2.3. Most fungal galls are solid, non-chambered swellings; these are described in Chapter 3. They belong to a wide range of families, some not nowadays regarded as true fungi (‘fungi’ is used here colloquially, to include all groups traditionally studied by mycologists, see Table 2.3 in Box 2.3; true fungi are included in the kingdom Fungi, with a capital initial letter). A few species, not closely related, cause virescences and witches’ brooms, and are included below. Most galling species are host-specific, restricted to a single plant species or genus. Their growths tend to be due to simple enlargement of cells of the host rather than to an increase in cell number, and also to an increase in the size and number of air spaces in the plant. A few galls are more elaborate but none approaches the complexity of insect galls (e.g. those of cynipids and cecidomyiids), with their distinct layers of highly differentiated tissues.

BOX 2.3. Biology of fungi (Information from Kirk et al., 2001; Alexopoulos et al., 1996; Ingram & Robertson, 1999)

‘Fungi’, as described in this book, include organisms traditionally studied by mycologists, i.e. true fungi (belonging to the kingdom Fungi) and also fungus-like organisms now classified as Protozoa or Chromista. True fungi are organisms composed of branching thread-like tubes called ‘hyphae’ that form a more or less diffuse mass called a ‘mycelium’. The hyphae are usually microscopic in size, with a diameter of about 1–15 µm (occasionally thicker; 1,000 µm = 1 mm), and a few centimetres to several metres long. Their cell walls are composed of chitin and glucans, as in insects and other arthropods, sometimes with septa dividing them into separate cells, and are without chlorophyll. Some species exist as rounded cells and do not form hyphae, and sometimes are also without cell walls; many of these have cell walls composed of cellulose, as in plants, and are not true fungi. Table 2.3 shows a simplified classification.

The main characteristics of selected groups, and examples of gall causers, are summarised below:

Myxomycota–slime moulds; no hyphae; rounded cells without a cell wall called plasmodia, with many nuclei; no galls.

Plasmodiophoromycota–grow as a plasmodium with many nuclei, no cell wall; zoospores with 2 flagella; gall causers are clubroot (Plasmodiophora brassicae), powdery scab (Spongospora subterranea).

Oomycota: Albuginales, Peronosporales–hyphae with cellulose cell walls, no septa, contain many nuclei (diploid); zoospores with 2 flagella; include potato blight (Phytophthora infestans), not a gall, downy mildews (Peronosporaceae), most of which are not gall causers, and white blister (e.g. Albugo candida), which is gall-causing.

Chytridiomycota: Chytridiales–no hyphae; rounded cells with several nuclei and chitin cell walls; zoospores with 1 flagellum; potato wart disease (Synchytrium endobioticum) is gall-causing.

Zygomycota–hyphae with chitin walls, no septa, contain many nuclei (haploid); spores without flagella; no galls.

Ascomycota–hyphae with chitin and glucan cell walls, septa with large pores allowing nuclei (haploid) to move freely; spores without flagella; includes the powdery mildews (Erysiphaceae), ergots (Claviceps purpurea) and Taphrina species, many of which are gall causers.

TABLE 2.3. A simplified classification of fungi, i.e. the organisms traditionally studied by mycologists; only orders with plant pathogens are identified (from Kirk et al., 2001; Ingram & Robertson, 1999; Brian Spooner, pers. com.)

Basidiomycota–hyphae with chitin and glucan cell walls, septa with pores too small to allow free movement of nuclei; cells dikaryotic (each with a pair of haploid nuclei); spores of several types, without flagella; includes Basidiomycetes (with large fruit bodies; none is gall-causing), and rusts, smuts and Exobasidium species, some of which do cause galls. The mycorrhizas are also gall-causing.

‘Deuteromycetes’–no gall causers.

Fungal hyphae are surrounded by a membrane (the plasmalemma) and most have a rigid cell wall, except at the extreme tip where the wall is thin and flexible and allows the hypha to grow. The plasmalemma controls the uptake of nutrients and water and the secretion of chemicals, e.g. enzymes that degrade plant cells and enable the fungus to invade plant tissues efficiently. Hyphae are not waterproof and cannot survive where the environment is too dry–living inside plants avoids this problem. Many plant pathogenic fungi (i.e. those causing diseases), including gall causers, are biotrophs, feeding on living plant cells without killing them. They debilitate the host, reducing the efficiency of photosynthesis and often causing premature senescence. Their hyphae grow between and sometimes into cells, absorbing nutrients from intercellular spaces and from the cytoplasm of plant cells. Most fungal biotrophs produce specialised absorption surfaces called ‘haustoria’, absorption organs, which are inserted into cells. These penetrate the plant’s cell wall and invaginate the cell membrane without piercing it; the space between the haustorium and the cell membrane develops layers derived from both host and parasite, forming an elaborate surface for exchange of materials that move in both directions–the fungus absorbs nutrients from the host cell, and the cell receives substances from the fungus that modify its metabolism, to the benefit of the fungus. Although the Taphrinaceae and the smuts do not produce true haustoria, their hyphae penetrate host cells and absorb nutrients from them in a similar way.

Fungi disperse to new sites via spores. The spores of the more primitive groups are zoospores, with flagella, and require water in order to swim to new hosts. Ascomycetes and basidiomycetes have non-swimming spores, without flagella, and are less dependent on water. They produce a variety of spores that are known by different names in different groups, e.g. conidia and ascospores in ascomycetes; urediniospores, teliospores and basidiospores are examples of basidiomycete spores. The ascospores are protected inside a special chamber, the ascus, which in most families (but not the Taphrinaceae) is enclosed inside a fruit body, the ascocarp. Basidiospores are produced by special hyphae (the basidia) that, in mushrooms and toadstools, occur in the familiar fruitbodies, the basidiocarps. Rusts, smuts and exobasidiums do not produce basidiocarps; instead, the hyphae form cushions or blisters in the plant, which rupture the epidermis when mature, allowing the spores to escape. The spores may be shot into the air and carried away by the wind, or are dispersed in raindrops or by insects. Ascocarps and basidiocarps, and the cushions and blisters of rusts and smuts, can withstand desiccation, though humid conditions are necessary for germination of the spores. The variety of spores and lifecycles of rusts are particularly complex and are described further in Box 2.4; smuts are less complex (see Box 2.5).

To survive, biotrophic fungi need to be well adapted to their hosts and well synchronised with their hosts’ lifecycles, and so tend to be specific to a particular plant species or genus. Some species are so specialised that particular strains can survive only on particular cultivars of the host. They produce large numbers of spores, which ensures that some, at least, survive to germinate. Their lifecycles are often complex, usually with sexual and asexual phases, and with each involving different spore-producing structures. Lifecycles for selected gall-causing species are described in the text, and are best known in species that cause diseases of major economic importance. Although gall-causing fungi are highly specialised parasites, most host individuals in a population do not become galled and nor do they succumb to disease; the plants can fight back in a variety of ways, some highly sophisticated.

Mycologists do not usually use the word gall to describe a growth caused by a fungus. Common terms for fungal diseases of plants are wilts, curls, scabs, spots, rots, blasting and mildew, as well as witches’ brooms, tongues, tumours and cankers. The last four are clearly galls, fitting the definition used in this book (see Chapter 1), but many of the rest are not, although some are difficult to categorise as each type merges with others. Often, the precise effect on the host plant is not known in sufficient detail to decide whether a swelling is a true gall or not–the swelling may be due only to an increase in fungal tissue or in the air spaces in the plant rather than to an increase in the size or number of plant cells. As more is learnt about the precise effects of fungal pathogens on their host plants, fungal galls may turn out to be more widespread than previously thought.

Like galls of insects and other groups, most fungal galls probably act as nutrient sinks, though of greater or lesser strength, which divert sugars and other nutrients from their normal destination (meristems, flowers, seeds, roots) to the developing pathogen. The cells of the gall remain alive and active as long as the pathogen is growing, having to supply the fungus as well as themselves. The cells are characteristic of meristematic and secretory tissues, with enlarged nuclei and many organelles (see Chapter 1), though there is no distinct nutritive tissue that is typical of insect and mite galls.

The best-known fungal galls are those that cause notorious diseases, particularly of crop plants, e.g. clubroot and white blister of brassicas, powdery scab and wart disease of potatoes, ergots of cereals, black stem rust of wheat, peach leaf curl and maize smut. Clubroot and powdery scab are slime moulds, included in the Protozoa (see Table 2.3 in Box 2.3). These are persistent diseases that are difficult to eradicate due to their resistant spores that are easily blown about by the wind and spread on transplanted plants, boots and tools; they can survive in contaminated soil for years. Spores of wart disease, caused by a chytrid fungus, are similarly difficult to eradicate and may infect new potato tubers after lying dormant in the soil for 50 years. Through strenuous efforts of control, quarantine and the breeding of resistant potato varieties, this disease is now rare. White blister belongs to the Oomycota, not considered to be true fungi, and now included in a new kingdom Chromista (Box 2.3). Besides chytrids, some gall-causing ascomycetes and basidiomycetes, particularly some rusts and smuts, are responsible for serious diseases (these are true Fungi, Box 2.3 and Table 2.3). None of the familiar mushrooms and toadstools is a gall causer, although some may cause serious disease. (Mushrooms and toadstools make up the class Basidiomycetes, with a capital B; basidiomycetes, with a small initial letter, is often used as the common name of all in the phylum Basidiomycota). Rust fungi have particularly complex lifecycles, involving up to five types of spores (described in Box 2.4). Smuts are less complex (Box 2.5). Exobasidium species specialise on heathers (Ericaceae) and are not economically important, but some cause the most elaborate of all fungus galls.

BOX 2.4. Rusts: a summary of their characteristics and lifecycles (Information from Alexopoulos et al., 1996; Buczacki & Harris, 1998)

Rusts are basidiomycetes in the order Uredinales (see Box 2.3). All are parasitic in plants (and in other fungi), and most are pathogenic, causing disease. Most of the body grows as hyphae inside the host, with haustoria, absorption organs, at intervals that project into cells and absorb nutrients from them. Blisters appear on the surface of the stems and leaves, and burst when the spores inside mature, splitting the epidermis of the host. The blisters are full of powdery spores, yellow, orange, brown or black, the orange and brown ones giving these fungi their common name. The spores may be shot forcibly into the air or are blown from the surface of the plant in air currents, dispersed by the wind. Most rusts do not seem to cause galls on their host plants. Those that do normally cause slight swellings around the blisters in which the spores develop, and a few produce gross swollen distortions of stems or leaves.

All rusts have complex lifecycles. Typically, they are ‘heteroecious’, requiring two host plants belonging to unrelated families in which to complete their development, and producing up to five types of spore. One or more spore types have been lost in some species, and others are ‘autoecious’, completing their development on one host plant only. Fig. 3 summarises the full rust lifecycle and introduces the five spore types.

The five spore types, and the structures which produce them, are conventionally designated by Roman numerals:

FIG 3. Generalised lifecycle of a rust fungus (n, haploid stage; n+n, dikaryotic stage; 2 n, diploid stage; for explanation, see text; after Alexopoulos et al., 1996).

FIG 4. Aecial peridia of Gymnosporangium cornutum projecting from aecia on rowan (Sorbus aucuparia). (Robin Williams)

O. Spermogonia bearing spermatia (= pycniospores) and receptive hyphae. The tiny haploid (n) spermatia are produced in small, often flask-shaped structures called spermogonia, usually developing on the upper surface of leaves. These spores are often exuded from spermogonia in a sweet, thick ‘honeydew’ attractive to insects, which transfer them to new host plants; the lifecycle will continue only if the new host also bears spermogonia. On the new plant, the spermatia behave like male gametes, the female gametes being the receptive hyphae, produced from the wall of a spermogonium. If a drop of honeydew containing spermatia lands near a spermogonium, a receptive hypha will grow into the drop, a spermatium will fuse with it and its nucleus will move into the receptive hypha, forming a dikaryotic cell (i.e. a cell with two haploid nuclei). This will only happen if the spermatium and receptive hypha are of complementary strains (+ and–).

I. Aecia bearing aeciospores. These spores are produced in cup-like aecia, embedded often in the underside of leaves. An aecium usually consists of a group of dikaryotic hyphae, which bud off chains of dikaryotic aeciospores (each with two haploid nuclei) from their tips. As they mature and multiply, pressure builds up in the aecium and eventually the aeciospores burst through the domed roof (the cover of the aecium is called the periderm). The torn periderm often forms a fringe around the aeciospores that may protrude some distance, and is often characteristic of the rust species (Fig. 4). Aeciospores are dispersed by the wind. If they land on leaves of the correct primary host, they germinate, producing hyphae which penetrate epidermal cells or grow through stomata.

II. Uredinia bearing urediniospores. These are the familiar rust spores, typically light-coloured though appearing dark brown en masse, often called ‘summer spores’. They develop from the tips of dikaryotic hyphae, which congregate under the epidermis of the host plant forming pustules called uredinia. As the urediniospores mature, they press against the epidermis and are released when it ruptures. Urediniospores are dikaryotic and typically have a thick spiny wall. This is the ‘repeating’ stage of rusts: while conditions remain favourable in summer, uredinia produce several crops of spores, so spreading the disease rapidly to other host individuals. Like aeciospores, urediniospores germinate to produce hyphae, which penetrate the epidermis directly or enter through stomata. Various chemical and tactile stimuli lead the hypha to a stoma; if it fails to find one within a few hours, it will die. Urediniospores do not usually survive the winter, although the mycelia developed from them in a perennial part of the host may do so. More often, it is the next type of spore that overwinters.

III. Telia bearing teliospores. Later in the summer, groups of dikaryotic hyphal cells form themselves into pustules known as telia; sometimes, old uredinia are converted into telia. Teliospores form at the tip of the hyphae, thick-walled spores composed of one, two or more cells. The pustules are black or very dark brown due to the colour of the spores inside; they are known as ‘winter spores’ because most rusts overwinter in this stage. Each cell of a teliospore is dikaryotic (with two haploid nuclei) at first; then fusion between the nuclei occurs, resulting in diploid (2 n) uninucleate cells. They normally need a period of dormancy during winter before they continue development next spring.

IV. Basidia bearing basidiospores. These spores are produced from teliospores germinating when conditions improve in the spring. A ‘promycelium’ grows out of the teliospore, one from each of its cells. The diploid nucleus moves into the promycelium, undergoes meiosis to produce four haploid nuclei, and septa grow between them so separating them into four uninucleate cells. Each cell produces a short stalk (a ‘sterigma’) with a pear-or kidney-shaped basidiospore at its tip, and the haploid nucleus squeezes through the sterigma to the basidiospore. In some species, the basidiospore becomes binucleate when its nucleus divides; others remain uninucleate. Mature basidiospores are released from their sterigmata, shot into air currents and carried away. Basidiospores are thin-walled with a simple smooth surface; they do not last long if they fail to find an appropriate host. A basidiospore may germinate directly to produce an infective hypha, or produce another crop of basidiospores first; the hypha, like germinating aeciospores and urediniospores, penetrates the epidermis of the host or enters through a stoma. It produces a mycelium, and hyphae near the surface of the host develop into spermogonia, which produce spermatia and receptive hyphae (0) ready to start the cycle again. The four basidiospores developing from one teliospore are usually of two mating types (+ and–), two of each. Each type is self-sterile, producing spermatia of one mating type that must fuse with receptive hyphae of the complementary type in order to develop further.

Most rusts with all five spore stages in their lifecycle have two host plant species; stages II, III and IV develop on the primary host, with O and I on the secondary host. The effects of the hyphae and haustoria developing from all of these spores is usually debilitating to the host, causing disease. In gall-causing rusts, it is usually the aecial stage (I) that causes the galls, and usually galls occur on only one of the host plants.

BOX 2.5. Smuts and bunts: a summary of their characteristics and lifecycles (From Alexopoulos et al., 1996; Ingram & Robertson, 1998; Buczacki & Harris, 1998; Brian Spooner, pers. com.)

Smuts and bunts, like rust fungi, are basidiomycetes but belong to different orders, the Ustilaginales, Urocystidales and Microbotryales (see Box 2.3). Like rusts, they are obligate parasites on plants; they have a saprotrophic phase, though (i.e. living on dead plant material), and are easier to culture artificially than rusts. Although some species cause devastating diseases (e.g. bunt or stinking smut diseases on wheat caused by Tilletia spp., not gall causers), far fewer are of economic importance compared to rusts, and modern control measures are mainly effective.

Cereals and other grasses are common hosts, and some develop galls. Characteristic symptoms of smuts are sooty black spore masses, replacing the florets of grasses and the anthers or ovaries of flowers. Other species affect stems, causing swollen lesions filled with black spores, or leaves covered with black streaks, which eventually split lengthways into ribbons. Some infections are systemic, hyphae growing throughout the plant and not always producing symptoms at the surface.

The lifecycle of smuts is simpler than that of rusts (Box 2.4). There is no alternation between host species and, usually, only two types of spore are involved: basidiospores (or sporidia) and teliospores, also called ustilospores. Teliospores form the dark, sooty, powdery masses which give smuts their common name. Basidiospores usually initiate the infection. Hyphae derived from basidiospores are slender, septate, often highly convoluted, and grow between and into host cells though not penetrating the host cell membrane. No haustoria are produced; instead, hyphae are much branched or coiled, providing a surface area sufficiently large for adequate absorption of nutrients from host cells. The effect on the host plant at this stage is slight, often causing only minimal stunting.

As in rusts, the hyphae of smuts contain haploid nuclei for most of the lifecycle. They may be multinucleate early in development but become dikaryotic (each cell with two haploid nuclei) before teliospores, also dikaryotic, are produced. Teliospores develop in sori, which develop in the ovaries or anthers, or form swellings in stems and leaves of the host plant. These grow at the expense of host cells, which may disintegrate as the spores multiply (as in the anther smuts); in addition, host tissues may become swollen, due to both enlargement and multiplication of cells (as in the maize smut). Teliospores are the winter or resting spores of smuts and can survive dormant in the soil for years. Their surface ornamentation is important in identification: smooth or spiny, reticulate or with small tubercles. They may exist individually or be cemented together in masses forming spore balls. Unlike in many fungi, germination of smut teliospores seems to be favoured by the low temperatures of temperate regions; many species require a period of frosty weather to break their dormancy. Fusion of the two haploid spores in the teliospore occurs at some time; meiosis occurs as the spore germinates, prior to formation of basidiospores (each with one haploid nucleus), which infect new host plants in spring. Basidiospores of compatible strains (+ and–) fuse together during infection of the new hosts, so restoring the dikaryotic mycelium.

PARASITIC PLANTS

Parasitic plants are unusual and gall-causing plants even more so. About 1 per cent of all angiosperms worldwide are parasitic and only a small proportion of these causes a gall. Their galls are all virescences or solid swellings and some are spectacular–the largest witches’ brooms, caused by the dwarf mistletoes Arceuthobium (Viscaceae) on conifers in Canada are nearly as large as the host tree. Most gall-causing plants have a wide range of hosts in several unrelated families, and their effects are more dramatic on some hosts than on others. In Britain, virescences caused by plants are rare. Two species cause proliferation of fine rootlets and root hairs: bastard toadflax Thesium humifusum (Santalaceae) infests plants of chalk and limestone grassland, and yellow-rattle Rhinanthus minor (Scrophulariaceae) attacks a wide variety of grasses.

Solid swellings are more common. Mistletoes (Viscaceae) and dodders (Cuscutaceae) cause swellings at the point where parasite and host tissues meet. The British mistletoe Viscum album inserts peg-like haustoria (absorption organs) deep into the wood of the host tree, and their growth keeps pace with the growth of the branch. The host tissues surrounding the haustoria grow to form the gall swelling. The thin pinkish stems of dodders wind around their host’s shoots, mainly heather and gorse in Britain, attaching to the host’s stem by flat plate-like haustoria, and swellings form around these. In both groups, the vascular tissues of host and parasite are closely associated in the swelling, so that nutrients can easily diffuse from host to parasite. Some mistletoes can drain the host’s resources to such an extent that they are pests of trees in forests, plantations and orchards. On the other hand, mistletoe in Britain is in demand at Christmas and is often propagated intentionally on old apple trees. And in North and Central America, the galls of some mistletoes are so intricate and elaborate that they are made into ‘wood roses’ and sold as ornaments to tourists.

THE GALLS: VIRESCENCES AND WITCHES’ BROOMS

Proliferation and bunching of plant parts occurs because the tip of the shoot (the apical bud), or the root tip, lose their normal control of growth of the plant. Gardeners have a similar effect when they pinch out the growing point of tomato or bean plants, encouraging lateral buds and side shoots to develop and so enhancing the crop. In the galls, lateral buds grow in a disorganised way and internodes may be telescoped or elongated, producing a compact bushy gall or an untidy growth formed of weak and spindly shoots, with no production of flowers, fruits or seeds. The brooming effect is probably just a by-product of the infestation and may not give the gall causer any advantage over that of an ordinary parasite. Mite and insect virescences are more highly structured than those of bacteria and fungi and most involve development of some nutritive tissue (see Chapter 1). As in the more organised and structured galls of mites and insects, the gall causer must remain alive and active for the virescence to develop.

TABLE 2.4. Proportion of virescences in Britain caused by each of the main groups of gall-causing organisms (data from Redfern et al., 2002)

Witches’ brooms on trees and shrubs and phyllanthies, chloranthies and phyllodies on herbaceous plants are the commonest types of virescences. Unusual ones are the tongue-like protuberances that sprout from distorted alder cones, wood anemones with leaves of unusual shape, and grasses with roots sprouting from swellings on stems well above ground level. Table 2.4 lists the main groups of gall causers in Britain, and the total number of galls that each causes. In column 3 is an estimate of the proportion of these that are virescences–a third of bacterial galls are virescences and 15 per cent of mite galls. Otherwise, these galls are rare; most galls show a greater level of organisation.

Witches’ brooms caused by viruses?

A virus may cause the grossly distorted catkins of willows and sallows (Salix, Fig. 5). These galls can be common on both male and female catkins, persisting on the tree all year, and are particularly obvious in the leafless canopy in winter. They are often infested with an eriophyoid mite Stenacis triradiatus, which can be present in large numbers, and this has traditionally been assumed to be the causal agent. However, virus particles have been found inside cells and the gall develops even when the mites are killed by acaricides. The mite may be a vector for the virus. Infection also may occur artificially, or via contamination of wounds, or perhaps during grafting of infected on to healthy rootstocks.

The extent of the willow catkin gall varies markedly. The catkin may be only partially transformed, producing leafy buds instead of some flowers, but is still recognisably a catkin. Or it may be totally distorted, short and thick due to lack of elongation of the axis, with all of the flowers greened and leafy or aborted altogether. The varying degrees of distortion are shown clearly in Meyer & Maresquelle (1983; Fig. 106). Virus particles have been seen in companion cells of the phloem tissue of affected catkins, cells involved in transporting sugars and other products of photosynthesis around the plant. The virus may disrupt this transport, with the gall developing as a result.

FIG 5. Distorted catkins of willow (Salix) caused by a virus or phytoplasma. An ungalled catkin is shown, far right. (Tom Higginbottom, Robert Cameron)

Bacteria and phytoplasma virescences

Bacterial virescences are not common; probably those caused by phytoplasmas are more widespread. Rhodococcus fascians (previously referred to Corynebacterium) seems to cause multiplication of buds and brooming in a number of unrelated hosts, and is probably responsible for the common swellings covered with buds in the woody stem of garden Forsythia (Fig. 6), which sometimes disfigure whole bushes (the causal organism is not proven, however). The same bacterium is found in germinating sweet pea Pisum sativum seeds, which produce short, thickened shoots with multiple buds, several from each seed.

FIG 6. A gall probably caused by Rhodococcus fascians on garden Forsythia (Redfern et al., 2002, with permission).

Witches’ brooms on trees may be caused by phytoplasmas. An early symptom in autumn is the premature development and multiplication of axillary buds, buds that normally remain dormant until the following spring. Some develop into deformed shoots, elongated and spindly or with shortened internodes and with pale undersized leaves, while others form an untidy mass of fine roots intermingled with the shoots. The clover phyllody on Trifolium repens (Fig. 7) is caused by a phytoplasma and is probably the same organism that causes ‘green petal’ disease in garden strawberries (Fragaria x ananassa). Both are transmitted from plant to plant by leaf hoppers. The phytoplasma transforms sepals, petals, stamens and carpels into a bunch of abnormal leaves and causes young leaves to be stunted, yellowed and misshapen, and no seeds or fruits are produced.

FIG 7. Phyllody on white clover Trifolium repens caused by a phytoplasma (from Redfern et al., 2002, with permission).

Fungal virescences

Several different groups of true fungi cause galls and a few of these are virescences (Table 2.3 in Box 2.3). The spectacular witches’ brooms on birch (Betula, Fig. 8) are caused by the ascomycete Taphrina betulina (T. turgida is a synonym), though birch brooms may also be caused by phytoplasmas. Infection of a bud or young shoot leads to a proliferation of buds, and each produces a spindly twig with leaves that are often abnormal in size, shape and colour. Continued production of buds and growth over many years can produce brooms of immense size. The fungus is present as a perennial mycelium inside the broom and produces a bloom of ascospores on the leaves. Other Taphrina species cause brooms on other trees, e.g. T. wiesneri (= T. cerasi) on wild cherry Prunus avium and dwarf cherry P. cerasus. The brooms do not have much effect on the vigour of the trees, although they may affect the fruit crop. Other species of Taphrina are included in Chapter 3.

FIG 8. Witches’ brooms caused by Taphrina betulina on birch Betula.

FIG 9. Tongue gall on female cone of alder Alnus glutinosa caused by Taphrina alni. (Michael Chinery, Tom Higginbottom)

Taphrina alni (= T. amentorum) causes an unusual virescence on the female cones of alder Alnus glutinosa. The cone is distorted, with some of its bracts and ovaries enlarged, and the fungus projects as one or more tongue-like processes, often known as ‘languets’. These are green at first, then red or purple and later brown (Fig. 9). This gall has been very rare in Britain, known only in Cornwall in the 1940s. Since 1999, though, it has had a population explosion, becoming common in some years and appearing in many sites across Britain. The reason for this sudden upsurge is obscure; it would be interesting to follow its progress over future years.

Rust fungi are responsible for a number of witches’ brooms. A twiggy broom caused by Melampsorella caryophyllacearum occurs on silver fir Abies alba and sometimes on other Abies species. This can grow for many years, sometimes reaching an impressive size, with several occurring on one tree. Each starts as a swelling (the ‘cauldron’) that originates deep within a branch and breaks through superficial tissue layers. The swelling gives rise to one or more shoots with multiple buds, which sprout to produce the bushy broom, with needles shorter than normal. Aecia, which produce aeciospores (see Box 2.4), eventually appear on the underside of the needles, forming two rows of little cups, one each side of the midrib.

Rusts cause other types of virescences, too. The aecial stage of Uromyces pisi-sativi causes shoots of cypress spurge Euphorbia cyparissias to alter. The internodes lengthen and the leaves become short, thick and yellowish, quite different from their normal strap-shape. Fungal hyphae ramify through the galled leaf, eventually producing aecial cups on the surface, which discharge the aeciospores. Leaves of wood anemone Anemone nemorosa and cultivated garden anemones A. coronaria are altered in shape by the aecial stage of Tranzschelia discolor. Normally, the leaf is deeply cut with narrow elongate lobes. When galled, the lobes are thick and broad, with aecial cups eventually erupting on the under surface. The effect of the fungus can be slight, modifying only part of a compound leaf, or can be much more extensive. The leaves are sometimes so altered that the German botanist Dillenius in the eighteenth century described them as a separate species, the mysterious ‘Conjurer of Chalgrave’s Fern’ (see Chapter 14).

Virescences caused by smut fungi are rare and none are known in Britain.

Mite and insect virescences

Few virescences in Britain are caused by animals and most of these are mite galls (see Table 4.1 in Box 4.1, Chapter 4). About 25 species of eriophyoid mites and a few insects cause phyllanthies on a dozen or so dicotyledon families (e.g. dead-nettles and mints, Lamiaceae, and legumes, Fabaceae), and on grasses (Poaceae) and rushes (Juncaceae) amongst the monocotyledons. Apart from Aceria tenuis, which affects several genera of grasses, most of these virescences are specific to a host species or genus. Nomenclature of eriophyoid species is continually changing as more becomes known about this neglected group, and it may turn out that Aceria tenuis is an aggregate of several closely related species, each specialising on a particular genus or species of grass. Mite and insect virescences are more highly organised than those caused by viruses, bacteria, fungi and plants, and usually include some development of nutritive tissue (see Chapter 1). Their extent is affected by the number of feeding individuals–a large population will cause a large virescence.

The structure and development of most mite and insect virescences and the lifecycle of their causers are not well known. Exceptions are the phyllanthies of the mite Aceria lycopersici and the psyllid Livia juncorum, described here. Mite virescences, like mite galls in general, are often extremely hairy, covered with a thick white or grey pubescence. The mites live amongst the hairs, and depend on the high humidity produced there. General information on eriophyoid mites is included in Chapter 4 and Box 4.1 and the range of galls they cause is depicted in Fig. 38. For psylloids, see Box 4.4.

FIG 10. Galls of Aceria lycopersici on woody nightshade Solanum dulcamara (from Redfern et al., 2002, with permission).

Aceria lycopersici (Eriophyoidea) causes a phyllanthy on woody nightshade Solanum dulcamara, transforming a single flower, or the whole inflorescence, into a leafy mass (Fig. 10). Each leaf is unusually small with inrolled margins and is covered with silvery white woolly hairs. The gall is most extensive when the shoots are attacked when very young, long before the floral parts are recognisable. The parts that should develop into sepals, petals and stamens remain small and are leaf-like, and additional buds develop in their axils, becoming leafy and hairy. The shoot axis thickens and fails to elongate so that the modified buds remain bunched together. Inside each affected bud, a layer of nutritive tissue develops, about four cells thick. The mites pierce these cells and suck out their contents, destroying them one by one as they feed (see Meyer & Maresquelle, 1983, Fig. 8, for a good description of these galled tissues).

The lifecycle of A. lycopersici is known in more detail than that of most gall mites. Females appear on the plants in spring and search for new leaf and flower buds. They pierce individual cells, moving frequently, never feeding for long at the same place. Feeding stimulates the appearance of the first gall symptoms, the curling of the margins of the young leaves and the development of hairs. After some hours, spherical, pearly, translucent eggs appear amongst the hairs, a female laying one every day or two. The high humidity amongst the hairs of the young gall is necessary for survival of the egg, which loses water easily through its permeable shell. After five to seven days, the egg hatches and the young mite grows and moults twice. The first instar lasts for two or three days, the second for four or five days. Thus, the lifecycle from egg to adult takes only about 12 days when the climate is ideal and food is plentiful in spring and early summer. Most of this first generation develops into females, which start laying eggs when a day or two old. The few males deposit spermatophores, packets of sperm, which are picked up by some of the females, and fertilise a proportion of their eggs. As in most eriophyoid mites, reproduction is partly parthenogenetic (‘arrhenotoky’), i.e. unfertilised eggs develop into males and fertilised eggs into females (see Box 4.1). In the several generations that develop through the summer, the proportion of males and females varies, though there is usually a preponderance of females. Females continue to lay eggs as long as conditions are favourable. They, the females, can survive outside the gall, piercing cells of fully formed leaves and causing small necroses (scar tissue), 0.3 mm in diameter. Their survival rate, though, is only about 30 per cent of the females inside galls. While free-living, they may disperse to other buds and young leaves and initiate new galls. Sometimes they lay eggs outside the galls, but the larvae do not usually survive–humidity is too low and there is no specialised nutritive tissue. As the climate deteriorates in late summer, newly fertilised females migrate to overwintering sites, crevices in bark and beneath the scales of dormant buds, and the rest of the population dies. The females require a period of low temperatures before they can reproduce the following year. Although they may vary in size through the year, the overwintering generation is no different in morphology from the summer females. There is no alternation between the protogynes and deutogynes typical of many eriophyoid mites (see Box 4.1).

A. lycopersici is normally specific to woody nightshade. It can, however, produce similar galls when experimentally transferred to crop plants belonging to the same family, Solanaceae (e.g. tomato, pimento and potato). This does not normally happen in the wild but, if it were to do so, the mite could become a serious pest.

The best-known insect virescence, the tassel gall on rushes (Juncus), is caused by Livia juncorum (Psylloidea: Psyllidae). This gall (Fig. 11) affects Juncus articulatus and, perhaps, other species of small rush. Like the mite galls above, the extent of the virescence varies, depending on the stage of flower development at the time of infestation and the number of psyllids involved. A few flowers in the panicle may become greened and leafy, or the whole inflorescence may be affected, and develop the reddish colour characteristic of the gall. In a heavily attacked inflorescence, the flower parts are grossly enlarged and thickened, showing no trace of their floral origins. Leaf buds develop in their axils and become affected too, which, with the lack of elongation of the shoot, results in a bunch of thick, crowded, modified leaves. The psyllids live between the affected parts (Meyer & Maresquelle, 1983, Fig. 105, contrasts the structure of a normal and a galled inflorescence). Unusually for an insect gall, there is no development of nutritive tissue in this gall. The psyllid’s mouthparts, like aphids’, are a bundle of stylets that are inserted into the plant, and move between cells until they reach and pierce a sieve tube of the phloem tissue. Thus, easily digestible food is acquired on tap and specialised nutritive tissue is unnecessary. Vascular strands probably multiply in this gall, so facilitating feeding by the psyllids.

FIG 11. Tassel gall on Juncus articulatus caused by Livia juncorum. (Tom Higginbottom)

FIG 12. Gall of Mayetiola graminis on Poa nemoralis (from Redfern et al., 2002, with permission).

Livia juncorum has an annual lifecycle. The adult psyllids overwinter in moss and debris; nymphs normally do not survive after the first frosts of autumn. In the spring after mating, females lay eggs in a row under the newly developing leaf sheaths, each egg attached by a short stalk inserted into the epidermis. Egg-laying continues until the end of the summer. Eggs hatch in about two weeks, the hatchlings start feeding, and the first signs of gall formation appear in May. There are five nymphal instars, and adults appear about 30 days after oviposition. In northern France, three or four generations follow, development continuing as long as conditions remain favourable.

In Britain, generations reduce in number from south to north. The first two nymphal instars can initiate a gall but only the first instar can cause a fully formed gall. Its extent, i.e. the number of galled shoots in the inflorescence, depends on the weather and on the size of the psyllid population. If spring is early and followed by a warm summer, plant and psyllids become active early and large numbers of psyllids build up, so inducing an extensive gall. In less favourable conditions, swelling is limited and the gall is much less obvious.

Virescences caused by gall midges (Diptera: Cecidomyiidae) are unusual. Mayetiola graminis induces a bunch of roots on stems of wood meadow grass (Poa nemoralis), well above the level of the soil (Fig. 12). The larva feeds in a swelling above a node, in a chamber between the stem and leaf sheath, and the roots grow out from the free side of the stem. These, like normal roots, have a central core of vascular tissue and develop from deep within the stem. A layer of nutritive tissue several cells thick develops from the epidermis and cortex cells beneath the larva. The swelling is due mainly to enlargement of pith cells, which develop unusually large nuclei and fill the lumen (usually hollow) of the stem. The first instar larvae initiate the gall, hatching from eggs laid on the upper surface of the young leaves in May. Two or more larvae crawl into the space between the leaf sheath and the stem; they start feeding and the gall develops around them. They grow through two instars. Then the skin of the full-grown second instar hardens into a light brown puparium, 4–5 mm long, inside which the third instar, and later the pupa, develop. Mayetiola is unusual in that the third instar does not feed. The larvae are full-grown in August and overwinter in their puparia. They pupate next spring and adults emerge in May.

CHAPTER 3

Solid Swellings

SWELLINGS ON PLANTS, WHICH are more or less solid, without regular chambers or cavities, are the subject of this chapter. Like virescences (Chapter 2), their growth is unlimited and their size varies, and they often grow larger on younger than older hosts. They range from single enlarged cells, in algae, to growths of a metre or more on trees; crown gall, caused by Agrobacterium tumefaciens, can be huge. Most of the organisms causing these galls are microscopic: viruses, bacteria and fungi, and a very unusual animal (a rotifer). At the other extreme are parasitic plants, such as mistletoe Viscum album and dodders Cuscuta species, which cause swellings at the point of entry into their host plant. Viruses, bacteria and the rotifer live inside the cells of the host, while fungi and parasitic plants mainly grow between the cells. Gall induction by all of these groups is unusual; most of the species dependent on living hosts are straightforward parasites, which may cause spots, curls, wilts or rots, but do not normally induce swellings in their hosts. Like ordinary parasites, the gall causers of this chapter often exist in the host outside their galls, not provoking any symptoms. Most fungi, for example, only cause swellings where hyphae accumulate beneath the surface before producing spores. Viruses and bacteria may cause secondary galls, some distance away from the site of infection and devoid of inducing organisms.

The swellings are described in turn, from the simplest, caused by a rotifer Proales werneckii, to the relatively elaborate growths caused by the basidiomycete fungus Exobasidium, although none of these are as structurally complex as some insect galls. Several are economically important pathogens, having major effects on agricultural crops in Britain and elsewhere (see also Chapter 13). Background information on the biology of the non-animal groups (viruses, bacteria, parasitic plants and fungi) is given in the previous chapter and in Boxes 2.1 to 2.5.

SINGLE SWOLLEN CELLS

These are the simplest of galls and are caused by microscopic organisms. Most occur on small hosts, algae or fungi, or are hidden away underground causing swellings of plant root hairs. As a result, they and their inducing organisms are little known. Here, one example is given, caused by a strange rotifer, the only gall caused by an animal to be included in this chapter.

Proales werneckii (Rotifera: Proalidae) causes enlargement of the cells of the alga Vaucheria. This is the only gall-causing rotifer known and the only animal known to gall freshwater algae. It parasitises a number of species and is widely distributed from Devon and Sussex to central Scotland, but it is rarely recorded, probably because algae are neglected by cecidologists. The galls were first illustrated in 1787 on Vaucheria dilatata by Vahl, who thought they were normal parts of the alga (they could easily be mistaken for antheridia, the male sex organs). Early in the nineteenth century, Roth described two varieties of V. dilatata, based on galled and ungalled filaments. They were recognised as galls in 1803 and ascribed to an ‘insect’ Cyclops lupula (Cyclops are tiny Crustacea belonging to the order Copepoda). In 1834, Ehrenberg recognised the causer as a rotifer that he described as a new species Notommata werneckii, which was assigned to the genus Proales in 1889 (these and the following details are from Spooner, in Williams, 1994).

The young female rotifer induces a gall by biting through a filament of the alga near to the growing tip where the membrane is thinnest, and crawling inside. It moves about at first, then settles and the gall develops–perhaps the swelling is caused by a secretion from the rotifer’s large salivary glands. Several females may enter one filament with each inducing a gall. The galls grow quickly as long as the female feeds continuously. Galls normally develop on one side of a filament, though they are occasionally terminal in position (Fig. 13). Dome-shaped at first, they usually elongate to reach a maximum length of nearly 1 mm in eight to ten days. Mature galls are adorned with horn-like projections, varying in number from a pair to 16, mostly extending from the free end. The body of the gall is dark green and packed with chloroplasts, with the projections almost colourless. The rotifer feeds on the cell contents but avoids the chloroplasts. It appears black, due to faeces that are retained within its body so avoiding contamination of its food supply. When mature, it lays 30–80 eggs and then dies, leaving the gall full of eggs. Then the chloroplasts lose their green colour, the gall becomes transparent and the eggs can clearly be seen. These are released when the filament dies and rots.

FIG 13. Galls of Proales werneckii in Vaucheria dilatata (original from Vahl, 1787; reproduced with the kind permission of the Director and the Board of Trustees, Royal Botanic Gardens, Kew).

SOLID SWELLINGS CAUSED BY VIRUSES

One virus gall is well known–the wound tumour virus gall (WTV) (see Chapter 2 and Box 2.1 for background information on viruses). WTV causes swellings of various sizes that look externally very like small crown galls (caused by Agrobacterium tumefaciens, see below). Tomato, potato and tobacco plants (all Solanaceae) seem to be particularly susceptible, and the legume Melilotus alba (Fabaceae; Fig. 14) is regularly used as an experimental host plant. Common symptoms are tumours in stems and roots and swellings of veins on the undersides of leaves. Different strains of the virus differ in their ability to produce galls–host plants may suffer little ill effect or be so severely damaged that they die before reproducing. The swellings are generally associated with wounds, for example, those caused by feeding leafhoppers (e.g. Agallia and Agalliopsis), which are common vectors, or tissue damage at the point where lateral roots emerge from the main root, where normal growth causes cells to be crushed.

FIG 14. Wound tumour virus galls induced in the laboratory on the stem of white sweet clover Melilotus alba (after Matthews, 1991).

Most WTV infection starts in the phloem sieve tubes of stems or roots and spreads to adjacent tissues. The virus causes the cambium of vascular bundles to become unusually active and the phloem parenchyma cells to de-differentiate so that they become meristematic. Both produce phloem tissue that is abnormal, with malformed cells, so that the normal function of the phloem is impaired as it fails to transport sugars around the plant efficiently. The adjacent xylem is affected too and becomes less efficient at transporting water and mineral nutrients. In young tumours, there may be some organisation of cells into tissue layers: an outer zone of abnormal xylem elements, a central zone of actively dividing cells, and pseudo-phloem (without sieve tubes and companion cells) on the inside. Older tumours become disorganised, with no recognisable cell layers.

WTV induces the plant to produce an excess of growth hormone, overriding the normal regulation of growth. A secondary effect on Melilotus alba is the reduction in the number of root nodules induced by the bacterium Rhizobium (see below). They become swamped by vigorous growth of the WTV gall, preventing the plant from gaining benefit from the activity of the bacterium (see Box 2.2).

SOLID SWELLINGS CAUSED BY BACTERIA

Galls of three bacteria species are described here, all belonging to the same family, Rhizobiaceae; background information on the biology of bacteria is given in Chapter 2 and Box 2.2. Crown gall, caused by Agrobacterium tumefaciens, is the commonest and most widespread of all gall-causing bacterial diseases, found on more than 600 host species. Hosts are wild plants as well as cultivated species, and include trees, soft fruits, vegetables and the grapevine, as well as ornamental roses, rhododendrons and garden flowers. They include dicotyledons and conifers but not monocotyledons, and crown gall does not affect cereals and other grasses. The bacterium exists as several strains, most of which have a restricted host range, and some are more virulent than others. Most, though, do not seriously affect the host plant. The disease was first recognised in 1853. The causal agent was shown to be a bacterium by Erwin F. Smith in 1907, although the role of plasmids (see Box 2.2) was not understood until the late 1970s. Active research continues, with new discoveries on the interaction between pathogen and host at the molecular level regularly being made.

Crown gall tumours vary from the size of a pea to enormous growths in trunks and branches of trees. They are irregular in shape, fleshy when young, becoming hard (Fig. 15) and with no distinct internal organisation. They consist of a mass of parenchyma cells, supplied by disorganised vascular tissue which links up with the normal vascular bundles of the stem. The bacteria survive free-living in the soil and infect a plant through a wound, often at soil surface level at the junction of stem and root, i.e. in the ‘crown’ (hence the common name). Wounds are caused naturally at the point where side roots emerge from the main root, or can be caused by subterranean chewing insects or nematodes–these aid transmission though are not essential. Infection via grafts is common, particularly in fruit trees and roses. Dispersal by wind and rain enables above ground wounds to become infected too. Galls can develop anywhere in the plant and, because they are induced by plasmids, secondary tumours without bacteria develop even after the initial colonists have died out. Thus, bacteria-free grafts can infect a new host, as long as plasmids are present.

FIG 15. Crown gall caused by Agrobacterium tumefaciens on (a) bramble Rubus sp. (from Redfern et al., 2002, with permission) and (b) branches of horsechestnut Aesculus hippocastanum. (Keith Harris)

A related species, Agrobacterium rhizogenes, causes hairy root disease, a gall consisting of abnormal root-like processes on aerial parts of the plant, and it also infects a wide range of hosts. It behaves in a similar way to crown gall, with part of its plasmid (an Ri, root-inducing, plasmid) being inserted into the host’s DNA and coding for excess production of plant hormones and of opines (see Box 2.2). Some strains of A. rhizogenes produce an antibiotic, agrocin, which is toxic to A. tumefaciens and has been used as a method of biological control for some strains of crown gall (marketed as ‘No Gall’ and ‘Galltrol’). It can control crown gall on peach trees, roses and other nursery-grown crops, but is not effective against all strains (e.g. the strain on grapevine).

The common and familiar nodules on roots of legumes (Fabaceae) are caused by a small group of unusual bacteria belonging to the genus Rhizobium. This gall association is different from most others in that both partners benefit, although neither is completely dependent on the other. Rhizobium can survive for long periods (e.g. ten years) free-living in the soil, but its numbers gradually decline if the association fails to occur. The host legume survives well without bacteria if the soil is fertile; in infertile soil, especially one lacking sufficient nitrates, seedlings die and growth of established plants is poor if galls fail to form. But the relationship may not always be of mutual benefit, and can change if environmental conditions alter. In fertile soil rich in nitrates, the nodules are digested by the plant, along with the contained bacteria. Conversely, if seedlings are weak and soil conditions unfavourable, Rhizobium becomes pathogenic and can destroy root cells and kill the seedling. Like other gall causers, the bacteria acquire sugars and other nutrients from the host plant, and also they gain some protection from generalised predators and extremes of climate. Rhizobia are different from most other gall causers (and most other organisms) in that they ‘fix’ atmospheric nitrogen, converting it into soluble nitrogen salts that are passed on to the plant and used to synthesise proteins. When the plant dies and decomposes, these nitrogen salts are released into the soil, so increasing its fertility.

The changes at cellular level that result in gall formation are fairly well known for Rhizobium nodules, though the processes involved are not fully understood. Bacterial cells multiply in the soil around the root of a legume seedling and cause its root hairs to curl and branch–this may be due to the amino acid tryptophan, released by the root and converted into the auxin, indole acetic acid (IAA), by the bacterium. Bacterial cells become attached to the root hairs and penetrate them, each forming an infection thread that grows along the hair to the root. Changes in hormone levels in the root cells, controlled by a plasmid in the same way as in Agrobacterium tumefaciens (see Box 2.2), stimulate cell division and formation of the nodule. Unlike crown gall, though, tissues of the nodule are organised into distinct zones (Fig. 16). Zone III contains bacterial cells enlarged and transformed into ‘bacterioids’, which have the ability to fix gaseous nitrogen and convert it into ammonium ions. Zone I contains typical meristematic cells of the host, which divide causing the nodule to grow. Zone II is the region of infection, with host cells enlarging. In Zone IV, bacterioids and root cells break down, having reached the end of their useful life. The outer region of the nodule contains growing host plant cells, including xylem and phloem connected to the plant’s vascular system, which allow water, mineral ions and carbohydrates to be transported to the bacterioids, and nitrogenous salts produced by them to travel in the opposite direction.

Bacterioids in the nodules are capable of fixing gaseous nitrogen because they produce the enzyme nitrogenase. The enzyme is damaged and broken down by oxygen, a conundrum for the plant since oxygen is essential for survival of its root cells. So, the nodule needs to be highly organised biochemically, as well as structurally. This is achieved by the red pigment, haemoglobin (used as an oxygen carrier in the blood of vertebrates and other animals but not normally found in plants). Haemoglobin molecules combine with the oxygen diffusing into root cells from the soil, and prevent it from reaching the bacterioids, but release it into the root cells, as it is needed. The plant and bacterium are so well integrated that haemoglobin is synthesised jointly–Rhizobium produces the oxygen-binding ‘haem’ portion of the molecule, while the plant synthesises the protein ‘globin’. How this close association evolved is a matter of speculation; it may be the result of the transfer of bacterial genes into the plant’s genome sometime in its evolutionary past.

FIG 16. Diagrammatic section through a Rhizobium nodule on the root of lucerne Medicago sativa (after Meyer & Maresquelle, 1983). For explanation of the four zones, see text.

SOLID SWELLINGS CAUSED BY PARASITIC PLANTS

In Britain, species of two plant families cause galls on other plants: mistletoes (Viscaceae) on apple trees (Malus), limes (Tilia), hawthorns (Crataegus) and poplars (Populus), and dodders (Cuscutaceae) on gorse (Ulex), heather (Calluna) and stinging nettles (Urtica). Some of the enlargement is due to the parasite’s tissues inside the host, but much is caused by multiplication of the host’s cells. Although the tissues in the swelling are not highly organised, the vascular tissue of the parasite grows towards that of the host, and the xylem elements become so closely associated that water and nutrients pass from host to gall causer. The phloem elements are less closely associated and transfer of sugars to the parasite is probably less important, at least in mistletoes–these mistletoes are green (i.e. contain chlorophyll), allowing them to photosynthesise and to synthesise their own carbohydrates. Dodders have thin pinkish stems that twine around their hosts’ stems, attached by haustoria, and minute leaves reduced to scales. The plants contain very few chloroplasts and must be more dependent on the photosynthesis of their hosts than the mistletoes.

SOLID SWELLINGS CAUSED BY FUNGI

Most solid non-chambered galls are caused by fungi (‘fungi’ is used here to mean all groups traditionally studied by mycologists). Background information is given in Chapter 2 and Boxes 2.3, 2.4 and 2.5. The galls come in a variety of shapes: small swellings and cushions on leaves and stems, grotesque swellings on roots, and growths which distort whole fruits or replace the grain in grasses. The rest of this chapter describes examples from a range of families, selected for the spectacular galls that they cause or for the historical importance they have had. Fungi have complicated lifecycles; these are described in diagrams and in the text (but the detail can easily be skipped).

Clubroot or ‘finger and toe’ disease of brassicas, caused by Plasmodiophora brassicae (Plasmodiophoraceae)

P. brassicae can infest many Brassicaceae, attacking both wild species and cultivated cabbages, cauliflowers and their relatives, and garden flowers such as stocks and wallflowers. It is particularly a problem in gardens with wet acid soils. Although the species sometimes attacks plants in other families too, the gall is restricted to brassicas. The roots become swollen and misshapen (Fig. 17) and, above ground, the leaves wilt and become bronzed, the stems are stunted and eventually the whole plant collapses. Later, soft rot bacteria and fungi may affect the galls, and they decay into a slimy, evil-smelling mess.

As is characteristic of this family, the body of P. brassicae is a plasmodium, bounded by a membrane and containing many nuclei, but without cell walls. A generalised plasmodiophorid lifecycle is shown in Fig. 18. A resting spore (Fig. 19) in the soil germinates to form a primary zoospore with two flagella (Fig. 18 A, B). The zoospore swims to the appropriate host, attracted by specific chemicals leaking from the root, and infects a root hair using a remarkable mechanism. It attaches itself to the surface of a root hair, loses its flagella and forms a rounded cyst (Fig. 18 C, Fig. 20). A short hypha emerges from the cyst and clamps on to the root hair. Inside the cyst, a bullet-shaped ‘stachel’ develops with the nucleus of the zoospore enclosed in cytoplasm attached behind, and is fired through the wall of the root hair. Inside the cell wall, the membrane of the root hair remains intact, and encloses the pathogen–at this stage it resembles a small spherical amoeba. Although the infection process takes several hours, firing of the pathogen into the root hair is complete in only a few minutes. The nucleus of the zoospore divides to produce many daughter nuclei, and the zoospore enlarges using nutrients absorbed from the host cell. It is now a plasmodium; several plasmodia may grow in the same host cell, filling the available space in a cell now much enlarged. Eventually the plasmodium divides into a mass of secondary zoospores (Fig. 18 D, E), each with a single haploid nucleus and flagella, and these are released to infect more root hair cells (Fig. 18 F). Some of them attack new cells immediately (G) and repeat stages D–F but others, from several plasmodia, seem to act as gametes, sex cells. They fuse together in pairs forming larger spores each with two haploid nuclei (H, one nucleus from each parent), and these infect cells of the root cortex of the host. The cytoplasm of the spore grows, and the nuclei divide so producing a new, multinucleated, secondary plasmodium (I). It is this stage that causes the gall; the plasmodia stimulate the host’s cell to enlarge and divide, so producing the clubroot symptoms. Inside each plasmodium, nuclei from the different parents fuse in pairs to form large diploid nuclei (J). These soon divide, by meiosis, to produce haploid daughter nuclei (K). Cytoplasm, a membrane and a thick wall develop to enclose each daughter nucleus, and these form a mass of resting spores (L). Resting spores are released into the soil when the root dies and rots, and can survive dormant for eight or more years (some have been found to be viable after 40 years!). The cycle starts again when these spores germinate. P. brassicae is haploid for most of its lifecycle, except for the brief period when nuclei from two parents fuse (Fig. 18 J).

FIG 17. Clubroot gall of Plasmodiophora brassicae (from Redfern et al., 2002, with permission).

FIG 18. Generalised lifecycle of Plasmodiophora brassicae (modified after Alexopoulos et al., 1996). Only the parasite, not the host cell, is shown; for explanation, see text.

FIG 19. Resting spore (3–5 µm diameter) of Plasmodiophora brassicae (after Ingram & Robertson, 1999). It has a spiky wall (dark stipples), lipid droplets (open shapes) and a nucleus in the centre.

FIG 20. Infection mechanism of zoospore of Plasmodiophora brassicae (after Ingram & Robertson, 1999).

The clubroot symptoms appear because the balance of the host’s growth hormones (auxins and cytokinins) is upset. The effect of this is to increase the numbers and size of root cells, with the result that the main and lateral roots become grossly swollen. The rest of the plant is affected too. The vascular tissues of the root are disrupted and this affects transport of water and nutrients to the shoot, causing the leaves and stem to wilt. Clubroot was first recorded in Britain in 1795, in Scotland; watering cabbages with the bathwater of syphilitic patients was thought to be the cause–so explaining the strange shape of the affected roots! The disease is easily spread in transplanted plants and on boots and tools and control is difficult. Growing brassica crops in the same plot only once in five years’ rotation helps to reduce the abundance of surviving zoospores in the soil, and some host plant varieties are fairly resistant. Raising the soil pH and applying fungicides can help too but never gets rid of all spores. Traditional remedies might work–applying rhubarb, manure, soot, mothballs or eggshells to the planting holes, but there is no scientific basis to any of these!

Powdery scab of potato, Solanum tuberosum, caused by Spongospora subterranea (Plasmodiophoraceae)

This is a common and widespread disease, causing rough, brown, raised scabs on the surface of potato tubers. It is particularly common in wet soils, where lesions may enlarge to produce knobbly outgrowths (Fig. 21), and these sometimes distort the whole tuber. Powdery scab also attacks the roots of tomato, in the same family as potato, the Solanaceae.

FIG 21. Powdery scab of potato, Solanum tuberosum, caused by Spongospora subterranea (from Redfern et al., 2002, with permission).

S. subterranea has a lifecycle similar to that of Plasmodiophora brassicae, to which it is closely related. Powdery spore balls are released from the scabs; they contain resting spores that can survive in the soil for ten years or more. Germination is stimulated by chemicals released into the soil by the host, and produces bi-flagellated zoospores that swim up the chemical gradient to the host’s roots and infect root hairs in much the same way as in P. brassicae. Plasmodia develop in the root hairs and produce more zoospores, and these infect the tubers through ‘eyes’, lenticels and wounds; they develop into more plasmodia which cause the scabs.

Control of powdery potato scab is difficult; some potato varieties are more resistant than others, though none is immune. S. subterranea is a vector of the virus disease ‘potato mop top’ (not a gall), which can survive in resting spores for many years. Good husbandry seems the best treatment: improve the drainage of wet soils, do not grow potatoes in the same ground year after year, and burn potato tops and peelings instead of composting them.

White blister of brassicas, caused by Albugo candida (Albuginaceae)

Albugo candida is the best-known species of its family and illustrates the typical lifecycle and habits. It exists as many strains, each infecting a particular variety of cultivated Brassica (cabbage, cauliflower, horseradish, etc.), but is probably most familiar to naturalists on shepherd’s purse Capsella bursa-pastoris, where it forms a white encrustation on stems, leaves, flowers and seed pods, thickening and distorting these parts (Fig. 22).

Much of the lifecycle (Fig. 23) has been known for a long time, worked out by de Bary in the 1860s. The mycelium of the fungus ramifies between cells of the leaf or stem, acquiring nutrients via haustoria that penetrate the cell walls (A; Meyer & Maresquelle, 1983, and Alexopoulos et al., 1996, provide electron micrographs that illustrate the fine detail of these haustoria). Groups of hyphae become club-shaped sporangiophores, which bear the spores, and these clump beneath the epidermis causing swellings. Chains of sporangia develop at the tip of each sporangiophore and detach as they become mature; they cause the epidermis to bulge upwards (B) to form shiny white blisters, characteristic of the early stages of the disease. The epidermis eventually ruptures, releasing white sporangia (C) that form a powdery crust on the surface, and are dispersed by wind, rain splash and visiting insects. Mortality during dispersal is high; the sporangia must land in a suitable place and weather conditions must be favourable (warm and wet) if they are to develop further. If conditions are right, several zoospores form in each sporangium (D), each with a nucleus and a pair of flagella (E). Zoospores swim to a suitable spot on the host plant, lose their flagella and encyst (F), before germinating and sending out germ tubes (G), and these penetrate the host through stomata. The germ tubes develop into hyphae that grow through the mesophyll of the leaf or through the cortex of the stem, extending haustoria into adjacent cells (A).

FIG 22. Albugo candida on shepherd’s purse Capsella bursa-pastoris (from Redfern et al., 2002, with permission).

FIG 23. Lifecycle of Albugo candida (after Alexopoulos et al., 1996). See text for explanation.

This is the asexual lifecycle; the pathogen’s cells contain diploid nuclei and the cycle will continue throughout the summer as long as conditions are favourable. The sexual phase, involving male and female gametes, occurs in late summer and autumn and enables the organism to overwinter as resistant spores in the soil after the host plants have died back. The sex organs, oogonia (female) and antheridia (male), develop on hyphae inside the plant, and meiosis occurs within them, so that the gametes are haploid. There are several haploid nuclei inside each oogonium but only one survives. An antheridium attaches itself to the oogonium (Fig. 23, H), forms a fertilisation tube, and the male nucleus plus some cytoplasm passes into the oogonium (I). The male and female nuclei and cytoplasm fuse to form the zygote or oospore (J), so regaining the diploid condition. Oospores develop thick, ornamented walls (K), able to withstand harsh winter conditions, and are released into the soil when the plant dies and rots; there they remain dormant for several months. In spring, each oospore produces several bi-flagellated zoospores (L, M) and these swim to a new host plant, germinate (N) and enter the host, so starting the asexual phase of the lifecycle again. In contrast to the Plasmodiophoraceae, most of the lifecycle of A. candida is diploid.

White blister on C. bursa-pastoris causes thickening of much of the plant above ground. If flowers are affected, the normal structure of the stamens is completely modified: the anthers degenerate and produce rudimentary leafy outgrowths, vascular tissue multiplies, and no pollen is produced. Control of the disease on brassica crops is relatively easy using copper-containing fungicides (e.g. Bordeaux mixture) and burning the affected plants.

Wart disease of potato, Solanum tuberosum, caused by Synchytrium endobioticum (Synchytriaceae)

Known in Britain since about 1860, wart disease (or black wart disease) is a devastating disease of potato tubers. It causes green cauliflower-like growths on parts of the plant above ground and close to the soil surface, but it is the large warty galls that destroy the tubers (Fig. 24) for which it is best known. These are whitish at first and later blacken and decay.

FIG 24. Gall of Synchytrium endobioticum on a potato tuber Solanum tuberosum (from Redfern et al., 2002, with permission).

FIG 25. Lifecycle of Synchytrium endobioticum (modified after Curtis, 1921, described in Alexopoulos et al., 1996). See text for explanation.

The lifecycle of S. endobioticum is complex (Fig. 25). In spring, resting spores germinate to produce zoospores, each with a single posterior flagellum (Fig. 25 L). The zoospore swims through wet soil and penetrates an epidermal cell of a tuber, through an ‘eye’ (A), losing its tail in the process. Each zoospore grows within its host cell (B; the zoospore is now called a prosorus), the cell enlarges and surrounding cells are stimulated to divide and enlarge. The nucleus of the prosorus divides to form a multinucleated sorus (C), which becomes a sporangium when its cytoplasm divides and rounds up around each nucleus (D), forming more zoospores (E). These are released to infect more host cells, so causing further enlargement of the tuber. Several generations of this asexual cycle (the left-hand cycle in Fig. 25) occur throughout the summer, as long as conditions remain warm and wet, and it is this stage that produces the large warty galls characteristic of the disease. Inside, the galls consist mainly of large parenchyma cells without any differentiation into tissue layers. If conditions become less favourable, especially if soils dry out, the sorus, instead of becoming a sporangium (Fig. 25 D), becomes a gametangium, i.e. the spores it produces become gametes (F). These fuse in pairs, forming bi-flagellate spores with two haploid nuclei (G), which soon fuse. Each zygote invades a new host cell (H), loses its flagella and becomes a resting sporangium (I), with a thick wall (J) resistant to decay. Resting sporangia are released into the soil when the tubers turn black and rot during autumn and winter. When conditions improve next spring, the nucleus of the resting sporangium divides, probably by meiosis, to form the next generation of zoospores (K, L), which are released to infect the new season’s crop of potatoes.

Resting sporangia can remain viable in the soil for 50 years or more and are easily spread via contaminated soil and on boots and gardening tools, as well as in diseased tubers used as ‘seed’. Since outbreaks of the disease in the early twentieth century, which caused devastating losses of potato crops especially in northwest England and Scotland, law in Britain enforces stringent measures for its control. No potatoes can be grown in contaminated soil, and only immune cultivars can be grown in a safety zone around contaminated areas. Commercial growers must grow immune cultivars, although the rules are less restrictive for amateur gardeners. Nowadays the gall is rare, though it does turn up from time to time in gardens and allotments. The disease is found all over the world where potatoes are grown and has proved impossible to exterminate, due to its ability to mutate into new strains.

S. endobioticum can infect wild species of Solanum, e.g. woody nightshade S. dulcamara and black nightshade S. nigrum. It does not seem to attack other cultivated Solanaceae, though, such as tomatoes Lycopersicon esculentum. Other Synchytrium species gall wild plants from a variety of families: S. mercurialis causes small, glassy yellow raised blisters on dog’s mercury Mercurialis perennis, S. taraxaci causes small, yellow, orange or red bumps on leaves and stems of dandelion Taraxacum officinale and S. anemones produces tiny hard black galls anywhere on the shoots of wood anemone Anemone nemorosa.

Almond and peach leaf curl on Prunus species, caused by Taphrina deformans (Taphrinaceae)

All Taphrina species have a restricted host plant range, attacking selected host species within just one genus. T. deformans is the best known of the gall causers; it infests peach and nectarine Prunus persica and, less commonly, almond P. dulcis. Soon after bud burst in the spring, the leaves thicken, blister and curl downwards (Fig. 26), becoming yellow, then red with the upper surface covered with a pale bloom of spores, before turning a dirty brown and falling prematurely. The fungus also affects young shoots, flowers and fruits, particularly producing spore bloom on the smooth skins of nectarines. Severe infestations occur when the weather in spring is cold and wet; then trees lose their vigour, fail to produce fruit and can become totally defoliated. Hot, dry, sunny weather checks the spread of the disease.

FIG 26. Peach leaf curl caused by Taphrina deformans on peach Prunus persica. (Michael Chinery)

The lifecycle of T. deformans (Fig. 27) is typical of its genus and, for a fungus, is fairly simple. In summer, ascospores are released from the bloom on the gall and are dispersed by wind and rain; they will germinate on the surface of shoots if they land on a suitable host tree. They bud (A) producing sprout cells (B), each with one haploid nucleus, and continue to bud off new cells while the weather is mild and damp, forming a yeast-like colony that can be a considerable size. These cells develop thick resistant walls and overwinter under bud scales and in crevices in the bark. In spring, the overwintering spores germinate and produce hyphae with cross-walls, with pairs of haploid nuclei in each cell (i.e. they are dikaryotic, C). This is the infective mycelium; the hyphae infect new leaves as they appear in spring, penetrating deep between the mesophyll cells. The hyphae may push through cell walls into the cells, invaginating the cell membrane, but they do not form haustoria. The host cells respond by enlarging and multiplying, more so on one side of the leaf than the other, which produces the characteristic leaf curl; these cells actively supply nutrients to the growing parasite. Hyphae mass beneath the cuticle of the host plant and those near the surface enlarge to form a layer of ascogenous cells (i.e. cells that will form the asci, D), each containing one or more pairs of large haploid nuclei. The paired nuclei fuse (E), the resulting zygote divides, while the ascogenous cell elongates (F). The daughter nuclei (still diploid) move apart, and the cytoplasm divides to produce an upper cell that becomes the ascus cell (which will produce ascospores), and a lower stalk cell (G). The ascus nucleus now divides by meiosis (so the diploid phase is brief) to produce four haploid daughter nuclei (H), which divide again, producing a young ascus with eight haploid nuclei (I). The contents of the stalk cell by now have disintegrated. As the asci enlarge and elongate, they push up the cuticle of the host. Eventually they break through the cuticle and produce a compact layer of mature asci on the leaf surface, each ascus containing eight mature ascospores (J). The ascus layer and its spores produce the characteristic pale grey bloom on the leaves. Pressure builds up inside each ascus, which bursts, so shooting ascospores for some distance into the air. Air currents disperse them to new hosts where each starts budding (A) to begin the lifecycle again.

FIG 27. Lifecycle of Taphrina deformans (after Alexopoulos et al., 1996). See text for explanation.

Peach leaf curl is a difficult disease to eradicate. The best treatment is to burn affected parts early in the season before the ascospore bloom develops. In addition, a copper-based fungicide can be sprayed just after leaf fall or in early spring to destroy the yeast stage on the surface of the bark.

Other Taphrina species cause similar leaf curls, e.g. T. tosquinetii on alder (Alnus glutinosa) and T. populina on black poplar Populus nigra and its hybrids. T. populina causes upward bulges in the leaf blade, the underside lined with golden yellow asci. Several separate bulges may form on each leaf blade though usually they do not affect the whole leaf. A more dramatic gall is caused by Taphrina pruni on Prunus (blackthorn P. spinosa, wild plum P. domestica). The fruits are enlarged and distorted ‘pocket plums’ (Fig. 28) and often the shoots are stunted and swollen, and discoloured with small, distorted, strap-shaped leaves. Pocket plums become covered with an off-white bloom of ascospores, then shrivel completely and drop off. Inside, the fruit is hollow with no trace of a stone. As often occurs with fungal galls, pocket plums are normally not very common, but there are occasional years when every fruit seems to be affected.

FIG 28. Pocket plums on blackthorn caused by Taphrina pruni. (Michael Chinery)

Ergot disease of grasses caused by Claviceps purpurea (Clavicipitaceae)

Ergots (the name derived from the French argot = a cock’s spur) are fungal growths, purplish or dark violet-black in colour when ripe, which replace the grain of individual florets in the flowering heads of grasses (Fig. 29). Many wild grasses are hosts and C. purpurea exists in a number of forms, each specialising in a different group of grasses. In large-flowered species, such as marram grass Ammophila arenaria and cord grass Spartina, the ergots grow particularly large. Before the widespread use of fungicides and modern seed-cleaning techniques, the disease was very common in cereals, especially in rye crops Secale cereale. Ergots often contaminated the flour and caused ergotism or ‘St Anthony’s fire’–this had profound effects on humans and their domestic animals especially between the fourteenth and eighteenth centuries, and probably has affected people throughout history (see Chapter 14).

FIG 29. Ergots in florets of couch grass Elytrigia repens. (Michael Chinery)

There was some doubt as to whether the ergot is a true gall because, when shed from the plant, it is formed entirely of fungal tissue. However, the ovary of a very young floret is invaded by hyphae that grow upwards from the base of the floret and this apparently does cause enlargement and multiplication of young cells of the floret. The ergots look like galls, too, and deserve to be included here. The species has an interesting history (see Chapter 14).

The lifecycle of Claviceps purpurea is shown in Fig. 30. The ergots (sclerotia, A) fall to the ground with the ripened grain in late summer or autumn and overwinter. They are tough and, like seeds, may lie dormant for several years; often they require exposure to cold temperatures before they can develop further. In spring, they germinate, each producing several rounded knobs on stalks called stromata (B), which are nearly a centimetre tall and easily visible to the naked eye. Tiny cavities (perithecia) develop over the surface of each knob and inside these sexual reproduction takes place. Each perithecium contains an ascogonium (the female sex organ), which contains several haploid nuclei (C). From its base, hypha-like antheridia (male sex organs) develop, and these also contain haploid nuclei. The tip of one antheridium fuses with the ascogonium (D) and a nucleus, acting like a male gamete, passes into the ascogonium and fuses with one of its nuclei to form a zygote, the ascus mother cell (E). Several ascus mother cells form inside each perithecium, and each develops into an elongated ascus (F). Its nucleus divides by meiosis to form eight haploid ascospores, each long and thread-like (G). So, the diploid part of this lifecycle is short, lasting only a few weeks in spring. When the host grass flowers in early summer, the ascospores are discharged forcibly from the asci and perithecia; they are thrown upwards away from the ground, carried by air currents or raindrops and, with luck, are caught by a ripe stigma of the correct species of grass. Each stigma is receptive for only a brief period and synchrony between fungus and host is essential for survival. On the stigma, the ascospore germinates (Fig. 30, H), and sends a hypha down the style to the ovary in the same way as a pollen tube grows from a pollen grain. A branched mycelium (I) develops, utilising nutrients intended for the growth of seeds and destroying the tissues of the ovary as it grows, and replaces the ovary with a soft, white mycelial mat (J). This develops special short hyphae (conidiophores) that bud off tiny oval spores called conidia from their tips. The conidia (K) become mixed with a sweet sticky ‘honeydew’, which exudes in droplets from the grass florets. The honeydew attracts insects, which pick up the spores and carry them to new hosts, where the spores germinate (L) and produce new mycelial mats and more conidia. This asexual cycle can continue as long as conditions are favourable and there are suitable grasses in flower. Meanwhile, the mycelial mat (J) hardens and darkens, and transforms into a pink, purple or nearly black sclerotium (ergot, A). One inflorescence may bear several ergots amongst the healthy grain; they are larger and often fall to the ground before the grain, or are knocked off during harvesting. They overwinter in the soil thus completing the lifecycle.

FIG 30. Lifecycle of Claviceps purpurea in rye Secale cereale and other cereals (after Alexopoulos, et al., 1996). For explanation, see text.

Ergots are toxic to mammals, humans as well as domestic animals, and have caused serious diseases in the past (now largely controlled, see Chapter 13). The toxicity is caused by several alkaloids, which produce the purple colours, and these are concentrated in the outer layers of the ergot. They protect the ergot from desiccation and damage by sunlight, and discourage predation by mice and voles during the overwintering period in the soil. Ergot contamination of the grain can be controlled: effective seed cleaning will remove ergots from harvested grain, and numbers in the soil can be reduced by rotating cereal crops with non-cereals, or by deep ploughing to bury the ergots, and by removing grass weeds which could act as alternative hosts. Fungicides work too but only if applied in the brief period when the grass flowers are ripe and their stigmas exposed.

Black stem rust disease on wheat Triticum and common barberry Berberis vulgaris caused by Puccinia graminis (Uredinales: Pucciniaceae)

This rust fungus is a pathogen of many grasses, wild species as well as the cereals wheat, barley Hordeum distichon, rye Secale cereale and oats Avena sativa and, particularly on wheat, it has caused disease epidemics worldwide (see Chapter 13). Its galls are not very distinct and develop only on the secondary host, common barberry (and on Mahonia species abroad), and are now rare in Britain. Puccinia graminis is a complex of microspecies that consists of several morphologically identical formae speciales, and these attack particular species of grass (e.g. f.sp. tritici on wheat, f.sp. avenae on oats and f.sp. secalis on rye). All of them have the same secondary host, the common barberry. The gall is caused by the aecial stage (see Box 2.4, Chapter 2, for a summary of the five spore stages of rusts). The galls appear as slightly raised cushions on the underside of barberry leaves and on fruits; they are pale yellow at first and later purple-red and bear bright orange cup-like aecia. Orange spots also occur on the upper surface of the leaves (these are the spermogonia, Fig. 31). The uredinial and telial stages occur on the leaves of the grasses where they appear as rust-red or black streaks and spots.

The lifecycle of P. graminis on wheat and barberry is complex with all five spore stages and was one of the first rust cycles to be worked out in detail. In mid-and late summer, teliospores (Fig. 31 A) are produced on wheat or wild grasses; they are two-celled and thick walled, with a diploid nucleus in each cell, and they overwinter on stubble and dead leaves. In spring, each cell of the spore germinates and produces a promycelium and four tiny basidiospores (B). The nucleus moves into the promycelium, divides by meiosis and the four haploid daughter nuclei move into the basidiospores. There are two types of spore (+ and–), which are equivalent to ‘male’ and ‘female’–this is the start of the sexual stage of the black stem rust. The basidiospores are forcibly thrown from the stubble surface and are dispersed by the wind. If they land on suitable leaves of a barberry bush, and if environmental conditions are favourable, they will germinate. Each basidiospore produces a hypha that penetrates the leaf epidermis and develops into a mycelium, growing through the mesophyll of the leaf. Spores are so common that usually several of each type (+ and–) land on the same leaf so that hyphae of both types are present inside. A few days later, hyphae nearest to the upper epidermis of the leaf develop spermogonia (C), flask-shaped structures that open to the outside. Hyphae in the flask produce chains of minute spermatia, which when mature are exuded in droplets of ‘honeydew’ through the top of the flask. Each spermogonium also produces receptive hyphae from its wall, which grow out of the opening and await fertilisation, a process facilitated by insects. A fly or ant, attracted by the honeydew, picks up spermatia on its mouthparts and these are brushed off by receptive hyphae protruding from the next spermogonium it visits. If they are of complementary strains, one + and the other–, the spermatium and receptive hypha fuse and the nucleus and cytoplasm of the spermatium pass into the receptive hypha (C). Spermatia and receptive hyphae can both be either + or–, making sex in rusts a complicated process.

FIG 31. Lifecycle of Puccinia graminis (after Alexopoulos et al., 1996). Open and closed circles represent nuclei of complementary (+ and–) types. Stages A–B occur on wheat or wild grasses; C–D on barberry; E–F on wheat or wild grasses again; for further explanation, see text.

While this is happening on the upper surface of the leaf, the mycelium growing from the basidiospore will have spread through the leaf. Nuclei from the spermatia travel via the receptive hyphae and this mycelium to the lower surface of the leaf. Hyphae near the lower epidermis now develop into aecia, now dikaryotic with + and–nuclei, and bud off chains of aeciospores (D), also dikaryotic, with two nuclei, one + and one–. The nuclei have travelled down the hyphae from the upper surface–but exactly how they move is obscure. Rust hyphae are divided into cells, so the nuclei must cross several cell membranes before arriving in an aecium. So, spermogonia and aecia develop on the same barberry leaves but only the aecial stage on the underside causes a gall.

Aeciospores are spiny; they mature in the aecium and eventually break through the lower epidermis of the barberry leaf to be blown away in the wind (perhaps they are dispersed by insects also). If they land on a suitable wheat plant, they produce hyphae, which penetrate the leaf epidermis and a dikaryotic mycelium develops inside. Eventually, uredinia (E) develop beneath the epidermis and bud off urediniospores on stalks. Urediniospores are dikaryotic, oval in shape, and are also spiny. As they mature, pressure builds up beneath the epidermis, eventually rupturing it and causing rust-red streaks running parallel to the leaf veins. In favourable summer weather, several generations of urediniospores are produced and infect new wheat plants, producing clouds of rust-coloured spores that spread from field to field. As the grain is ripening, uredinia start to produce a few teliospores amongst the urediniospores (F), each with two dikaryotic cells. In a few weeks, more and more teliospores are produced at the expense of urediniospores until the uredinia have become telia. The teliospores are dark red and, en masse, they appear as black streaks on the wheat stems and leaves. Inside the teliospore, the two haploid nuclei in each cell fuse, so forming the diploid spores that overwinter on stubble (A), completing the lifecycle.

In Britain, the galls on barberry are much rarer than they used to be (see Chapter 13), and are much more difficult to find than the black stem disease stage on grasses (though this is also less common than previously and nowadays is most likely to be found on rye grasses Lolium species). The galls occasionally turn up in southwest England where, it seems, they probably have developed from airborne spores blown in from Spain rather than from home-grown spores.

Galls on juniper Juniperus communis, hawthorn Crataegus monogyna and rowan Sorbus aucuparia caused by Gymnosporangium species (Uredinales: Pucciniaceae)

Gymnosporangium rusts cause remarkable and attractive galls of quite different appearance on each of their hosts. Juniper is the primary host of both G. clavariiforme and G. cornutum and their secondary hosts are hawthorn and rowan, respectively. The gall on juniper caused by G. clavariiforme is shown in Fig. 32.

The lifecycle is slightly less complex than that of the wheat–barberry rust. The telial stage affects the primary host juniper, causing spindle-shaped swellings in the branches on which telia and teliospores develop. Each teliospore has two cells and a stalk, and masses of these extend from the surface of the gall in spring, forming small, hard, yellowish-brown projections. When wetted by rain, their size and colour are transformed: the stalks rapidly absorb water and swell into bright orange-yellow, gelatinous, tongue-like structures, up to 1 cm long and with teliospores all over the surface. Basidiospores develop from the teliospores and are blown from the galls by the wind. If they land on a suitable alternate host, different galls develop in the leaves. On rowan, orange patches of G. cornutum develop in the thickness of the leaf. Black spermogonia appear on the upper surface in early summer, with characteristic horn-shaped aecia projecting from a swelling on the underside (Fig. 4 in Box 2.4, Chapter 2) in summer and autumn. These galls are common in Scotland, much rarer further south. On hawthorn, aecia of G. clavariiforme cause colourful swellings of the leaves, petioles and fruits (Fig. 33) and develop into telia–there is no uredinial stage in Gymnosporangium and no urediniospores. Aeciospores are blown by the wind to infect new junipers. Telia and teliospores continue to be produced by infected junipers for many years, because the mycelium within is perennial–the tree is infected for life.

FIG 32. Orange ‘tongues of fire’ (the telia) of Gymnosporangium clavariiforme on Juniperus communis (from Ingram & Robertson, 1999, with permission).

The effect of these Gymnosporangium species on their hosts is not severe. The related North American cedar–apple rust G. juniperi-virginianae, on the other hand, has caused extensive loss of apple crops in the USA. It attacks red cedars (Juniperus species, not true cedars) and apples or crab apples (Malus species). It was one of the first rusts to be grown in artificial culture in the 1950s, a difficult process previously thought to be impossible because rust fungi are obligate parasites and, in the wild, they cannot survive outside their hosts.

White pine blister rust Cronartia ribicola (Uredinales: Cronartiaceae) on five-needled pines, e.g. Weymouth pine Pinus strobus and currants Ribes

Cronartia ribicola is a destructive pathogen of five-needled pines. It alternates to blackcurrants and gooseberries (Ribes species, the primary hosts) but on these it only causes limited defoliation. On the pine trees, it causes canker galls on the branches and trunks; the galls enlarge each year and eventually encircle the branch or stem. The effect, at the least, is a reduction in the tree’s growth rate, but often the branch is killed or even the whole tree. The fungus probably originated in northeast Asia on Pinus cembra, a native tree that is tolerant of the disease, and is likely to have been imported into western Europe in the eighteenth and nineteenth centuries with exotic pines and currants destined for parks, botanical gardens and commercial forests. When it reached North America in the early twentieth century, it caused an economic disaster.

FIG 33. Aecial gall of Gymnosporangium clavariiforme in fruit of hawthorn Crataegus monogyna (from Redfern et al., 2002, with permission).

The lifecycle follows the usual rust pattern. The basidiospores develop on currant bushes, disperse in the wind and infect pine needles, producing hyphae that grow through the vascular tissue of the leaves and into the branches and trunk. These hyphae develop into a perennial mycelium that ramifies through the whole tree. In spring, small yellowish swellings appear in the bark and produce spermatia and, by the summer of the following year, these swellings have developed into large white or orange blisters bearing clusters of aecia. The blisters enlarge each year to form the woody cankers, which often exude resin that runs down the branch. The damage is done by the hyphae within the vascular tissue. These kill the cambium cells, perhaps after they have produced malformed bark and wood tissue, and prevent the vascular tissue from functioning. Orange aeciospores are released when the aecia burst, and can be dispersed several hundred kilometres by the wind to infect cultivated blackcurrants, gooseberries and flowering currants, as well as wild species. On Ribes, the aeciospores germinate to produce hyphae that grow through the leaf and, beneath the lower epidermis, these form uredinia on swellings that look like little yellow-brown igloos. They break through the epidermis, releasing urediniospores that infect more currant bushes throughout the summer. Telia, also developing on the undersides of the leaves, form brown, elongated, curved, bristle-like columns of cells, which produce the teliospores. These germinate in situ, producing basidiospores that disperse in the wind to reinfect the pine trees. So, galls develop on both hosts but only on the pine are they a symptom of a serious disease.

Maize smut disease on Zea mays caused by Ustilago maydis (Ustilaginales: Ustilaginaceae)

Ustilago maydis is the cause of the grossly distended grain on malformed maize cobs (Fig. 34), known since the sixteenth century throughout the warmer parts of Europe and, in recent years, becoming common in southern England. Inflorescences may be infected before the cobs develop and stems and leaves may also develop the characteristic swellings. Most damage to the host is done when the ovaries are infected, with galls replacing the grain, sometimes destroying cobs completely so that no seeds are produced. The swellings are silvery-white at first, and become dark brown as the mass of black teliospores (also known as ustilospores) develops inside (see Box 2.5, Chapter 2, for a general description of the lifecycle of smut fungi).

FIG 34. Galls on Zea mays caused by Ustilago maydis in grains of a corn cob. (Michael Chinery)

Fig. 35 summarises the lifecycle of maize smut–this is simpler than that of rusts, with one host and only two types of spore. Sooty masses of teliospores burst out of the galls when they split open in late summer and autumn; they are blown about by the wind or dispersed by rain, and land on the soil where they overwinter. The teliospores are dikaryotic at first and the two nuclei soon fuse to form a diploid zygote. They may germinate the following spring or lie dormant for several years before germinating. Similar to the rusts, the teliospore germinates into a promycelium and its diploid nucleus divides by meiosis to produce four haploid basidiospores, two of each type, + and–. The basidiospores are dispersed by wind and rain and a few land on young maize plants. Further development requires fusion of compatible (+ and–) basidiospores, which occurs on the surface of young leaves. The fused spore germinates and produces an infection hypha, which penetrates the epidermis of the leaf or enters via a stoma or a wound. The hyphae grow between cells towards a developing inflorescence, and produce galls that erupt from the stem and eventually infest the ovaries as these develop on the cobs in summer. Teliospores then develop in the galls and burst out of them when they split open in late summer and autumn to overwinter in the soil.

FIG 35. Lifecycle of maize smut Ustilago maydis on sweetcorn Zea mays (simplified after Buczacki & Harris, 1998).

There are two distinct phases in the lifecycle of U. maydis. The first is a unicellular, haploid phase that multiplies by budding in a yeast-like manner (the promycelium and basidiospore stage). This phase is non-pathogenic and can be grown in a petri dish in the laboratory. Second is a filamentous, dikaryotic phase, formed after fusion of two compatible haploid basidiospores; this is pathogenic and requires a maize plant in which to develop. Without a gall, teliospores are not produced and no meiosis occurs to produce basidiospores.

Other smut fungi have similar lifecycles to that of maize smut, though there are some interesting variations–for example, the anther smuts (these are not true smuts, see Table 2.3 in Box 2.3, Chapter 2). Microbotryum violaceum (Microbotryales: Microbotryaceae) is common on campions Silene in Britain. There are few symptoms until black telia appear instead of anthers, replacing the pollen with a mass of powdery black teliospores and sterilising the flower. Dispersal of the spores is more precise than in the maize smut: foraging insects collecting pollen become covered with teliospores and transport them to the next flowers they visit. The smut, moreover, transforms the ovaries of the flower into more stamens, so inducing the plant to produce extra pollen that it replaces with more spores. Infection in Microbotryum species (though not in U. maydis) is systemic and permanent and causes lifetime sterility of the host.

Galls of Exobasidium (Exobasidiales: Exobasidiaceae) on bilberry Vaccinium myrtillus and other Ericaceae

These galls are the most elaborate of any fungal galls. The Exobasidiales is a small order of basidiomycete fungi with affinities to the smuts (Ustilaginales) and most species are parasitic on heathers and their relatives. Exobasidium is the largest genus, causing galls on bilberry, cowberry (V. vitis-idaea) and several genera introduced in Britain (Rhododendron including azaleas, and Camellia). The galls are swellings, often reddened, on leaves, flowers or stems; they are mainly due to cell enlargement and sometimes enlarge the plant part to many times its normal size (Fig. 36). The swelling is composed of large parenchyma cells, with enhanced vascular tissues supplying the surface layers where the basidia and basidiospores develop.

The lifecycles of exobasidiums are not well known, probably because few hosts of economic importance are affected. In E. myrtilli, infection is systemic: plants are infected for life with new shoots becoming infected from the start of their growth. The mycelium grows through the rhizomes, stems and leaves of bilberry, with hyphae penetrating between cells and showing little outward sign of their presence until the galls appear. Nutrients are absorbed from host cells via distinctive lobed haustoria. On the surface of the gall, in or under the epidermis, the basidia produce haploid basidiospores that break through the epidermis and form a white, powdery layer on the surface, and are dispersed in the wind. If they land on a suitable host, the basidiospores germinate to produce infection hyphae, which penetrate the host epidermis directly or via stomata (in some species, secondary spores are produced from the basidiospores, and these produce the infection hyphae). Some exobasidiums are not perennial and must infest a new host plant each year. Probably basidiospores can survive dormant beneath bud scales of the host over the winter, before germinating in spring when temperatures rise.

FIG 36. Gall caused by Exobasidium myrtilli on bilberry Vaccinium myrtillus (from Redfern et al., 2002, with permission).

PART TWO

Open Galls

The galls in Chapters 4, 5 and 6 are open galls–an opening through which the gall causers can leave is part of the structure of the gall. The aperture may be permanently wide open, but more often is blocked with hairs or is temporarily closed, imprisoning the gallers. Generally, the causers of these galls cannot chew or burrow their way out. They escape through the aperture or when changes in the gall, i.e. in the plant, provide an exit. Open galls contrast with the closed chambered galls of Chapters 7, 8 and 9; in these, the gallers chew or burrow their way out or wait until the plant rots to release them.

Within each of the three open-gall chapters, the examples described illustrate a series of increasing complexity. Erinea, folds and rolls tend to be the simplest (Chapter 4): erinea are felt-like patches of glandular hairs, usually on the underside of leaves, and rolls and folds affect the leaf margins. Pits, blisters and pouches (Chapter 5) involve shallow or deeper invaginations of the leaf blade. The distinctive galls affecting buds and shoot tips, variously known as big buds, cigar galls, pineapple galls, rosettes and artichokes, make up Chapter 6. Most galls can be readily assigned to a descriptive category but there are some that are intermediate and do not fit easily into a particular section.

A variety of animals causes open galls: gall mites, thrips, bugs (psylloids,* aphids and scale insects), sawflies and gall midges. All of these groups cause leaf folds and rolls and gall mites, in addition, cause erinea; so, all appear in Chapter 4 and the same groups, with some omissions, cause the galls in Chapters 5 and 6. Background information on each group, including its classification and biology, is included in Chapter 4 and in Boxes 4.1–4.8. The remaining gall-causing insects, and nematodes which are unusual gall causers, cause closed galls and background information for them can be found in Chapters 7, 8 or 9. Although gall induction is common amongst eriophyoid mites, gall midges and the more ancient families of aphids, it is unusual in the other groups. Aphids (the more recent ones), psylloids, scales, thrips and sawflies that cause these open galls belong to families whose species are mainly free-living, and the gall causers are not very different from their non-galling relatives. Apart from the sawflies in which the galling species are closely related, gallers tend not to be closely related and it is clear that this way of life has arisen many times during the course of evolution. Sometimes, too, very different organisms cause galls with a similar appearance and structure.

Most of the galls included in Chapters 4 and 5 develop in new leaves, just bursting from their buds, at a time when they are actively growing and not yet fully differentiated. Those in Chapter 6 develop in buds, with tissues that by definition are young and undeveloped. Feeding by the gall causer ensures that the leaf or bud is typically modified and that the characteristic gall develops. Most of the insects and mites involved have piercing mouthparts: newly hatched nymphs in the psylloids, scales and aphids, and adult females in gall mites and thrips. In gall midges, the first instar larvae initiate the galls and feed probably without piercing cells. In all sawfly galls, the ovipositing female causes the initial reaction of the leaf and the gall may grow to almost its full size before the egg hatches; in most species, though, the caterpillar must feed to ensure that the gall develops to maturity.

CHAPTER 4

Erinea, Folds and Rolls

ERINEA AND FOLD AND roll galls include the simplest of galls, though even these vary considerably in complexity. The simplest are folds and rolls similar to those of caterpillars that fasten their rolls with silk, or of aphids that crumple and distort young leaves, and these hardly warrant inclusion as galls. At the other extreme, some approach the complexity of pouches and may even become closed, with their original opening sealed over. Erinea vary in the shape and structure of the glandular hairs and in the degree of modification of the leaf blade, and sometimes become incorporated into a leaf fold or roll. Two groups of mites and insects from four orders cause these galls and are introduced below, followed by examples of galls caused by each of the groups.

THE GALL MITES (ERIOPHYOIDEA AND TARSONEMOIDEA)

The general biology and classification of eriophyoids, the larger of these two groups, is described in Box 4.1. The name, Eriophyoidea, comes from the Greek erio (= wool) and phyes (= living) and refers to the erinea, woolly patches on undersides of leaves that are unique to these mites. All eriophyoids are herbivores and most are specific to a host plant species or to related species in the same genus. They feed on cells of the plant epidermis one at a time, piercing them with their needle-like stylets and sucking up the cell contents. In the gall causers, this induces surrounding cells to modify and form patches or a layer of nutritive tissue, or to become erineum hairs. A few mite species cause witches’ brooms and phyllanthies, described in Chapter 2. Many eriophyoids are not gall inducers, though. They are vagrants (wandering species) that cause discolouration, stippling, russetting or bronzing, perhaps with curling and shrivelling of leaves if infestation is heavy. Sometimes they carry disease organisms. Blackcurrant reversion disease (probably a virus) is transmitted by the big bud mite Cecidophyopsis ribis on blackcurrant (Ribes nigrum, see Chapter 6), and the bacterium Xanthomonas juglandis causes walnut blight on walnut (Juglans regia), transmitted by Aceria tristriata, which causes rounded pustules along the leaf veins. Fig. 38 illustrates the range of galls caused by mites, from erinea, flat on the surface or in hollows, to leaf rolls, blisters, pouches and enlarged buds, the most complex with distinct layers of tissues.* Most of these can vary considerably in size–the larger the population of mites, the larger the gall. A few species exist as more than one strain, each causing a different type of gall on the same plant. One race of Colomerus vitis, for example, causes erinea on the leaves of grapevine (Vitis vinifera), a second rolls the leaf margins, while a third causes enlarged and distorted buds. Hairiness is characteristic of eriophyoid galls, glandular hairs inside or elongate hairs surrounding the gall opening, and some galls are hairy inside and out.

BOX 4.1. Biology of eriophyoid mites (Information from Westphal, in Shorthouse & Rohfritsch, 1992; Boczek & Griffiths, in Williams, 1994; Lindquist et al., 1996; Oldfield in Raman et al., 2005)

Mites belong to the Acari, one of the classes of Chelicerata (= Arachnida), which also includes spiders, harvestmen and scorpions. Only two superfamilies of Acari include gall causers: a few species in the Tarsonemoidea (Box 4.2) and many species of Eriophyoidea, a much larger group. Worldwide, there are over 300 genera and nearly 4,000 species in the Eriophyoidea.

Eriophyoids are highly modified mites, with pale-coloured, elongated, worm-like bodies only about 0.1–0.4 mm long, and with only two pairs of legs (Fig. 37), thus easily distinguishable from other mites. Their morphology is fairly constant, however, making recognition of species difficult (certainly the job of a specialist). Most species, though, are specific to a particular host plant and cause characteristic damage, so that species can often be recognised at secondhand.

FIG 37. Generalised eriophyoid mite (from Redfern et al., 2010, with permission).

The Eriophyoidea is divided into three families (Table 4.1) according to the structure of the dorsal shield and the mouthparts, and the number and arrangement of setae on the dorsal shield. Most gall-causing species belong to the Eriophyidae.

The mouthparts of eriophyoids are characteristic and all species feed in a similar manner. The chelicerae (jaws) are fine stylets, surrounded by a pair of pedipalps that are more or less fused to form a suction cup, and the mite feeds by piercing and sucking. The suction cup is closely applied to the surface of a leaf, helped by arching of the body, which is anchored by a sucker at the posterior end and by the adhesive tips of the legs. The chelicerae move against each other and pierce individual cells, with mites sucking for periods of a minute or so up to several hours or one to two days; the longer the feeding, the more elaborate the gall.

The generalised lifecycle of eriophyoid mites is fairly simple: an egg, two immature instars (known as a larva and a nymph) that resemble adults, and the adult male or female, with females always commoner than males. They are ‘haplodiploid’ organisms: males are haploid, produced from unfertilised eggs and with one set of chromosomes from the mother (‘arrhenotokous parthenogenesis’), and females are diploid, developing from fertilised eggs with a set of chromosomes from each parent. ‘Thelytokous parthenogenesis’ may occur in some species (i.e. diploid females also produced from unfertilised eggs) but this has not yet been proven. Many gall-inducing species, at least those inhabiting deciduous host plants in temperate climates, produce two types of female: the overwintering ‘deutogynes’ that lay eggs in spring in newly developing buds or leaves, and ‘protogynes’ that develop from them and produce several generations in summer. These two forms are sometimes so different that they have formerly been assigned to different genera. A proportion of the protogynes of some species are ovoviviparous, i.e. the egg hatches inside the female’s body and the larva escapes probably through her genital opening. This occurs in Eriophyes laevis, which causes the common pouch galls on alder Alnus glutinosa leaves, Vasates quadripedes, a mite recently introduced to Britain that causes pouch galls on leaves of silver maple Acer saccharinum, and Phytoptus avellanae, the cause of big bud on hazel Corylus avellana, and probably will be found in other species, too. The sexes do not mate directly. The male deposits sacs of sperm (spermatophores) on stalks on the plant surface and the female mounts one, enabling the sperm to enter her genital opening. They are stored, available to fertilise her eggs later (mite sperm, unlike those of other arachnids, insects and most other animals, do not have flagella). A female lays 50 or more eggs and the population increases rapidly through several generations involving protogyne females during favourable conditions in the summer. Mites leave the galls in late summer and the second type of female, deutogynes that have already been fertilised, overwinter in crevices in the bark or under bud scales. Next spring, the deutogynes emerge to found the next generation of galls. Dispersal between plants is passive, mostly in the wind or on the bodies of insects, birds and other animals.

TABLE 4.1. Number of genera worldwide (and approximate number of species) of the three families of the Eriophyoidea, with examples of gall causers and notes on hosts, habits, etc. (after Amrine & Stasny, 1994; Lindquist et al., 1996; Oldfield, in Raman et al., 2005)

FIG 38. The variety of eriophyoid galls, indicating a possible sequence of development (modified after Westphal, in Shorthouse & Rohfritsch, 1992). British examples (listed below) are included for each type apart from 24, which does not occur in Britain. The gall mite (black areas) and woody layer (crosses, in 22) are shown. 1. Eriophyes leiosoma on Tilia; 2. Aceria varia on Populus; 3. Aculus hippocastani on Aesculus (often leaf is bulged upwards); hairs are multicelled; 4. Eriophyes distinguendus (new name for E. paderinus) on Prunus; 5. Acalitus brevitarsus on Alnus; 6. Aceria nalepai (new name for Eriophyes inangulis) on Alnus; 7. Colomerus vitis on Vitis (hairs single-celled when young, multicelled later); 8. Aceria erinea on Juglans; 9. Aceria ulmi (new name for Aceria filiformis) on Ulmus; 10. Aceria tristriata on Juglans; 11. Aculops macrotrichus (was in Aceria) on Carpinus; 12. Aculus convolvuli (was in Aceria) on Convolvulus; 13. Epitrimerus trilobus on Sambucus; 14. Phyllocoptes goniothorax on Crataegus; 15. Phytoptus tetratrichus (was in Phytocoptella) on Tilia; 16. Aceria macrorhyncha on Acer; 17. Aculus laevis on Salix; 18. Eriophyes viburni on Viburnum; 19. Phyllocoptes eupadi (new name for Eriophyes padi) on Prunus; 20. Eriophyes tiliae (was in Phytoptus) on Tilia; 21. Eriophyes laevis (was in Phytoptus) on Alnus; 22. Aceria campestricola (new name for the misnamed Eriophyes ulmicola) on Ulmus; 23. Eriophyes similis on Prunus; 24. Eriophyes dichrostachia on Dichrostachia (an acacia, not British); 25. Trisetacus pini on Pinus; 26. Phytoptus avellanae (was in Phytocoptella) on Corylus; 27. Aceria lycopersici (new name for A. cladophthirus) on Solanum.

Tarsenomoidea (Box 4.2) is a much smaller group of mites than the Eriophyoidea and the few galls that they cause are not well known. Some species are crop pests. Their damage ranges from streaks and mottles on leaves and stems to bulges on leaf veins, thickened pits, shortening of internodes and brooming. Tarsonemids, even the gall causers, tend to have wide host ranges. Steneotarsonemus spirifex attacks cereals and other grasses: it causes spiral deformation of the stem inside the sheath, malformed inflorescences and spikelets without grain, types of damage which probably can be classified as galling. Steneotarsonemus nitidus is an inquiline in the pouch galls of Eriophyes laevis on alder (Alnus glutinosa). It is a ‘cleptoparasite’: it invades the galls, feeds on the nutritive hairs within, multiplies and drives away the rightful occupants. The hairs provide thin-walled cells, suitable for piercing by the short stylets of the tarsonemid. Perhaps cleptoparasitism, via the specialised hairs of eriophyoid galls, was the route by which the tarsonemids developed the ability to feed on plants (most of them are fungal feeders).

GALL THRIPS (THYSANOPTERA)

For information on the biology and classification of thrips, see Box 4.3. Most thrips galls are curls, folds or rolls of leaves, or leaf pouches or rosettes, and nearly all have permanent openings to the outside. The most complex have layers of supporting tissues. A few species cause swollen bladders in buds or, rarely, enclosed swellings in young twigs. Most gall causers are restricted to trees and shrubs in the tropics and warm temperate regions and are common in India, southeast Asia and Australia. There are no thrips galls in Britain. Some plant genera are particularly susceptible to galling by thrips, e.g. figs (Ficus, Moraceae), peppers (Piper, Piperaceae) and acacias (Acacia, Mimosaceae). On Australian acacias they are common on the phyllodes, the flattened and extended leaf petioles that behave as leaves in species that have lost their leaf blades. These plants are major constituents of the flora of their areas and offer a widely available source of food, so it is not surprising that thrips have specialised on them.

BOX 4.2. Biology of tarsonemoid mites (Information from Jeppson et al., 1975; Channabasavanna & Nangia, in Ananthakrishnan, 1984)

The superfamily Tarsonemoidea belongs, like the Eriophyoidea (see Box 4.1), to the arthropod class Acari, and is divided into three families; only the Tarsonemidae includes phytophagous species and many of these attack grasses (Poaceae). The few galls caused by them are rudimentary, usually small, difficult to find, and hardly qualify as galls.

All tarsonemid mites are small, only 0.1–0.3 mm long. They are similar to most other mites, and unlike eriophyoids, in that most adults have rounded or ovoid bodies, a relatively hard integument and four pairs of legs. Some species in the genus Steneotarsonemus, which includes gall-causing species in Britain, are more eriophyoid-like with an elongate body, but retain the four pairs of legs with the third and fourth pairs placed near to the rear end. Perhaps a species of this type evolved to produce the Eriophyoidea, losing its two hind pairs of legs.

Like other Acari, tarsonemoids have two pairs of mouthparts, the piercing chelicerae and segmented pedipalps. The chelicerae are simple stylets, well adapted to piercing fungal hyphae and algae, but not capable of penetrating plant cells with thick walls and waxy cuticles. Some species ingest whole fungal spores. On plants most are restricted to succulent tissues with thin cell walls, as in meristems.

Males are generally smaller than females and have enlarged hind legs, which are used to hold females during copulation–fertilisation, unlike in the eriophyoids, is internal. Arrhenotokous parthenogenesis is probably common (i.e. males produced from unfertilised haploid eggs and females from fertilised diploid eggs). Males, unlike females, do not use the hind legs in walking; they are held up above the body and, besides holding on to the female during mating, are used to carry pupae and females above their backs to new feeding sites. The lifecycle, as far as is known, has more stages than in most eriophyoids. Eggs, laid singly by the female, are ovoid, white and large, sometimes half as big as the female. They hatch into white, six-legged larvae, the fourth pair of legs not yet developed. When full-grown, perhaps after several instars, larvae moult into relatively immobile pupae, sometimes with two stages, and transformation to the adult takes place within the pupal skin. The adult emerges through a dorsal split, and its skin hardens and darkens.

BOX 4.3. Biology of thrips (Information from Ananthakrishnan, 1979; Raman & Ananthakrishnan, in Ananthakrishnan, 1984; Ananthakrishnan, in Shorthouse & Rohfritsch, 1992; Crespi, 1992; Mound, in Williams, 1994; Crespi et al., 1997; Mound & Morris, in Raman et al., 2005)

Thrips belong to the Thysanoptera, an insect order that is mainly tropical or subtropical. More than 5,000 species worldwide in nine families have been described, though the true number is probably twice this. Eight families belong to the suborder Terebrantia and a single large family, the Phlaeothripidae, makes up the suborder Tubulifera (Table 4.2a). In the Terebrantia, the female has a saw-like ovipositor which allows eggs to be inserted into leaves. In the Tubulifera, eggs are glued on to the surface of plants by the tubular tip of the abdomen; there is no ovipositor. Gall induction is uncommon in thrips–about 500 galling species are known (probably < 10 per cent of all species are gallers, though) and nearly all belong to the subfamily Phlaeothripinae. Within the Phlaeothripinae, Mound (in Williams, 1994) identifies three major ‘lineages’, the Liothrips-, Haplothrips- and Phlaeothrips-lineages (Table 4.2b), whose members specialise, respectively, on green leaves, flowers and pollen, and fungal hyphae living on dead wood and leaf litter. Within these lineages, gall induction occurs sporadically in many genera and has evolved independently many times. Almost all thrips galls occur in the tropics and southern hemisphere, particularly in India, Indonesia, the Pacific islands and Australia. Ten per cent of all galls known in India are caused by thrips. South American and southern African faunas are poor, perhaps because they are not well known. The better-known gall faunas of North America and of Europe, though, are definitely meagre.

Thrips feed on fungi and detritus, and some are predatory, but most are herbivores, restricted to particular species or genera of plants, feeding on flowers, pollen, leaves, buds, fruits or young twigs. Their host plants range from ferns to flowering plants. Plants in about eight families are common hosts (including figs Moraceae, peppers Piperaceae, acacias Mimosaceae and eucalypts Myrtaceae) and most galls occur in these families too. There is some evidence, e.g. in Australian acacias, that thrips and their host plants have evolved and diversified together. Some species, especially non-gall-causing Thripinae, are agricultural pests, some acting as vectors of plant diseases.

Thrips are small insects (mostly 0.5–12 mm long) with elongate bodies and two pairs of narrow, strap-shaped wings fringed with long hairs; typical adults are winged, though some species include morphs (forms) in which the wings are reduced in size or are lost. They are not strong fliers but can be dispersed considerable distances passively in the wind. Their mouthparts are asymmetrical–unusual in an insect group. They have only one mandible (the right one does not develop beyond the embryo) plus a pair of maxillary stylets which join to form a food canal. The mandible punches a hole in a plant cell, the stylets are inserted, saliva is injected in and the contents sucked out.

TABLE 4.2. Classification of the Thysanoptera, with an indication of feeding preferences and an estimation of the proportion of gall causers worldwide (information from Mound, in Williams, 1994; Mound & Morris, in Raman et al., 2005)

(a) Families and subfamilies within the two suborders.

(b) The three lineages within the Phlaeothripinae.

The stylets are longer in many Tubulifera compared to Terebrantia, allowing deeper penetration into plant tissues, but are rarely long enough to reach the vascular tissues. Most thrips feed on individual plant or fungal cells, or pollen grains, sucking them dry one by one. Species that feed on fungal spores have wider stylets enabling whole spores to be swallowed.

The typical thrips lifecycle starts with an egg, then two larval instars followed by a prepupa and one (Terebrantia) or two (Tubulifera) pupae before the adult stage. The prepupa and pupae are non-feeding and almost immobile stages in which considerable reorganisation of the body tissues occurs. They fulfil a similar function to the pupal stage of endopterygote insects but have evolved independently. Thrips are haplodiploid insects: females are diploid, with two sets of chromosomes in each cell and develop from fertilised eggs, and males are haploid, developing from unfertilised eggs containing only one set of chromosomes from the mother (so are produced by arrhenotokous parthenogenesis, as in eriophyoid mites and many Hymenoptera). Females, generally, are at least three times as common as males and live longer. In a few species, males are very rare and females are produced from unfertilised diploid eggs, i.e. by thelytokous parthenogenesis. In galling species, mating often occurs in the old gall before the females disperse to new leaves; they lay their eggs on the new leaves, and each female can lay between 10 and 70 eggs. Most thrips produce several generations a year, perhaps 10–12 where temperatures are favourable all year and the host plant produces new leaves continually, to 2–3 where the climate is more seasonal. Development time from egg to adult varies from about 30 days to 40–45 days, depending on temperature, and often the generations overlap, with adults, eggs, larvae and pupae all present in a colony at the same time.

In some fungus-feeding and gall-causing Tubulifera, females and males are polymorphic, i.e. a species exists in several forms. The female founding a colony, the fundatrix, is always fully winged and usually has enlarged forelegs armed with teeth. She fights other females for the best patch of fungus or gall site, and for ownership of the gall in the early stages of its development. Fighting can occur whether a gall is initiated by one or by several females. Her offspring, both males and females, have either full-sized (macropterous) or reduced (micropterous) wings and some micropterous morphs of both sexes have enlarged forelegs and become soldiers. They fight intruders: predators, parasitoids and, particularly, cleptoparasitic (also spelt kleptoparasitic) thrips, which attempt to invade a gall and take it over by killing the original owners. Combatants fight to the death, one eventually puncturing the skin of the other. Once the fundatrix has produced some soldiers, she can leave defence to them and concentrate on laying eggs. Soldiers tend to be sterile or sub-fertile. If they do succeed in mating and laying eggs, few survive because the fundatrices normally produce so many offspring that there is no space in the gall for more. Soldier morphs have evolved in those gall-causing thrips with the following characteristics:

(i) They suffer high rates of cleptoparasitism.

(ii) Intraspecific fighting is common (i.e. fundatrices of the same species fight for possession of the gall).

(iii) Individuals pupate in the gall and emerging adults remain for some time within it.

Evolution of soldiers indicates that the gall is a valuable resource worth fighting for, both as a predictable and reliable food source and as a protected brood chamber in which to rear the young.

In thrips, as in eriophyoid mites, it is feeding by the adult female rather than the larva that initiates the gall. Adults and larvae feed by puncturing individual cells and sucking out their contents. Their stylets are short–they can only reach the epidermis and one or two layers of mesophyll cells. The galls develop typical nutritive cells (described in Chapter 1 and Box 1.3), which occur as scattered patches or distinct layers. In some galls, vascular tissue proliferates, especially phloem sieve tubes that link with the veins of the leaf and provide the growing population with additional nutrients. Many thrips galls contain one or more thrips species in addition to the gall-causing species. These may be inquilines that invade the gall and feed on the gall tissue, sometimes modifying the shape and structure of the gall though unable to initiate galls themselves (as in some cynipid wasps). Or they may go further and kill or drive away the original population, taking over the gall for themselves (these are cleptoparasites, specialised inquilines that, as in Steneotarsonemus nitidus above, always kill or push out their hosts). At one time it was thought that several thrips species might initiate the gall, called a ‘company’ gall, but this turned out to be the result of lack of knowledge of the role of each species in the gall. When fully grown, the gall contains all stages of the host thrips, from eggs to adults, probably from several generations and perhaps from several founding females (fundatrices), and a mature gall may contain hundreds or several thousands of individuals. The size of the gall varies accordingly and often is affected by the age of the leaf too–galls initiated in very young leaves and buds reach a larger final size than in older leaves, and may survive on the tree for longer.

PLANT BUGS (HEMIPTERA)

Gall-causing is sporadic in the Hemiptera, although in some families it is common. Background information on biology and classification is given in Box 4.4 (psylloids), Box 4.5 (aphids) and Box 4.6 (scale insects). Gall-causing psylloids and scale insects (Psylloidea and Coccoidea) cause pit galls on leaves and stems and they and gall aphids (Aphidoidea) also cause leaf rolls and folds and pouches. A few species cause elaborate bud and bud-like galls (e.g. the scales Gallacoccus on Shorea, a Malaysian tree; see Chapter 6) or phyllanthies (e.g. the psyllid Livia juncorum on Juncus; see Chapter 2). Unrelated plant bugs can cause similar galls. Many species, if their populations are large, cause some distortion and curling particularly of young leaves and shoots but without associated thickening, and this damage is not considered to be galling. Other species build shelters called ‘lerps’, constructed by a single nymph from hardened honeydew (e.g. psylloids of the family Spondyliaspidae, Box 4.4). The structure and shape of a lerp is characteristic for a species–it may be in the form of a single or bivalved shell or be tube-shaped. They are most abundant in Australia, particularly on Eucalyptus (Myrtaceae) and, like galls, can provide protection against desiccation and parasitism, probably more efficiently than simple leaf curling. A problem common to all plant bugs, especially those in enclosed galls, is disposal of waste honeydew, which can be copious and can contaminate the bug and its feeding site. The solutions to this problem are ingenious. Some species are attended by ants, which effectively remove the honeydew. Some secrete wax, which prevents honeydew adhering to the body and coats and isolates droplets of it. In others, various mechanisms in and around the anus enable the bug to squirt the honeydew some distance away from its body. Some species dispose of it in their lerps or other protective covers, and a few regurgitate it back into the plant and so none is produced.

Damage to host plants by galling hemipterans is usually slight (wounding, removal of sap and, perhaps, toxic effects of saliva) unless they transmit diseases. In gall aphids that alternate between two host plants, damage to hosts is much more likely to be caused by the free-living stages on the secondary herbaceous host plants than by the galling stages on the primary hosts, which are trees and shrubs. The damage caused by the grape phylloxera Daktulosphaira vitifoliae is exceptionally severe. This is an unusual aphid that has threatened production of wine in many parts of the world (see Chapter 13). Like other gall aphids, though, it is the form feeding on the roots (the equivalent of the secondary host) that destroys the plants (see Chapter 5).

BOX 4.4. Biology of psylloids (Information from Hodkinson, in Ananthakrishnan, 1984; Meyer, 1987; Burckhardt, in Raman et al., 2005)

Psylloids, or jumping plant lice, belong to the superfamily Psylloidea (Hemiptera: Sternorrhyncha). There are about 3,000 species worldwide in six families with about 350 gall causers. The Triozidae, Calophyidae and Phacopteronidae contain most gallers, and most of their galls are pits, folds, rolls or pouches in the leaves (Table 4.3).

Phyllanthies and root galls are unusual. Most galling species are specific to a single host genus or species, usually a dicotyledon. Exceptions are Livia juncorum and its relatives that specialise on rushes (Juncaceae) or sedges (Cyperaceae), both monocot genera, and a few non-British species that feed on palms (also monocots) and conifers. The psylloid fauna of the tropics, particularly in South America and southeast Asia, is much more diverse than in temperate climates and gall causers are commoner there too.

TABLE 4.3. Relative abundance of galls of different types caused by psylloid families (modified after Hodkinson, in Ananthakrishnan, 1984; Burckhardt, in Raman et al., 2005) (+++ common, ++ frequent, + rare,–gall not formed)

There are two types of psylloid nymph:

(i) The more robust, less flattened, and more mobile insect, with longer legs and antennae, typical of Psyllidae, Carsidaridae and Phacopteronidae.

(ii) The more delicate, very flat, less mobile nymphs, with short legs and antennae, found in Triozidae, Calophyidae and Homotomidae.

The flattened type (ii) nymphs are typical of those causing pit galls, where the nymph is sedentary for most of its life. Although there are exceptions, the more robust and mobile nymphs of type (i) are free-living or live in large leaf rolls and folds containing many individuals.

Mouthparts and feeding are typical of the Hemiptera: the mandibles and maxillae are elongated into fine stylets, protected by the modified labium. The stylets, typically, are inserted deep into the plant, moving between individual cells until they reach and penetrate the vascular tissues. Most psylloids are phloem feeders, piercing individual sieve tubes or phloem parenchyma cells and sucking up the soluble contents. All species inject enzymes that start the process of digestion outside the body of the psyllid. A few species have short stylets, perhaps in the nymphs only, and suck the contents of mesophyll cells of the leaf. During feeding, a salivary sheath usually forms around the stylets, enabling them to tap into the same sieve tube should they be withdrawn temporarily.

The typical lifecycle starts with an egg, then five nymphal instars, and adult males and females in equal proportions. Reproduction is normally sexual, though in some populations of a few species males are absent. In temperate climates, there is often just one generation a year with the final nymphal instar or the adult becoming quiescent and overwintering, or the egg overwinters. Some temperate climate species have three or four generations in summer but must shut down during winter. Tropical species, though, may breed continuously, perhaps with ten generations in a year.

In all Hemiptera, as in most gall-causing insects, galls are initiated by first instar nymphs (i.e. larvae) and growth of the gall will cease if they stop feeding. As in gall mites and thrips, hemipterans feed by piercing cells and sucking juices from them, and affect the plant by injecting saliva into the wound, but they do not usually kill the cells. Most aphids, psylloids and scales have long stylets, enabling them to reach the vascular tissues deep in the plant and to tap the sugary sap therein. In some Hemiptera, though, the stylets are shorter, unable to reach the phloem, and nutritive tissue develops as in the galls of other groups.

The lifecycles of most psylloids and scales are simple and sexual, starting with an egg and involving only one host plant. A few species, including some gall causers, produce live young and, in some, males are unknown or rare (thelytokous parthenogenesis: females producing more females without mating). Aphids are more complex with regular alternation between parthenogenetic (thelytokous) generations in the summer and sexual reproduction in an autumn generation (except in the pistacia aphids, see Box 4.5, in which the parthenogenetic generations occur in the winter and the sexual in spring). Some aphids are ‘heteroecious’ too, i.e. they alternate between two host plants. The less advanced families, the Adelgidae and Phylloxeridae, lay eggs while the more recent Aphididae are viviparous. Viviparity speeds up the rate of reproduction and huge numbers of offspring can be produced in a very short time; adult females are expectant grandmothers, depositing daughters with female embryos already developing inside them. These complexities are increased by polymorphism, the existence of a number of morphs or specialised forms of the insect (see Box 4.5). Some morphs carry out housekeeping duties and others are soldiers that defend the gall–soldiers also occur in thrips (see Box 4.3).

BOX 4.5. Biology of aphids (Information from Heie, 1980; Wool, in Ananthakrishnan, 1984, 2004, in Raman et al., 2005; Moran, 1992; Blackman & Eastop, 1994; Foster & Northcott, in Williams, 1994; Havill & Foottit, 2007)

Aphids belong to the very large superfamily Aphidoidea (Hemiptera: Sternorrhyncha) whose origins are ancient–fossil aphids are known from the Permian (225–280 million years ago). Classification of the group is still in dispute. Heie (1980) separates the Adelgidae and Phylloxeridae from the remaining aphid families, putting them into the superfamily Phylloxeroidea, while Blackman & Eastop (1994) include these two as the oldest families of the Aphidoidea, and demote most of Heie’s families to subfamilies within the Aphididae (Table 4.4).

There are about 4400 species worldwide–a conservative estimate; Chakrabarti (in Raman & Gupta, 2007) increases this to 4700 species for the Aphididae alone. Most species are specific to a single host plant species, or to two unrelated host plants between which the lifecycle alternates. Only a minority of the whole superfamily (probably less than 15 per cent) causes galls and most of these occur in the older families Adelgidae and Phylloxeridae, and the two most primtive subfamilies of the Aphididae, the Eriosomatinae and the Hormaphidinae. Few galls are caused by species in the more recent subfamilies and, in many of these, the distortion involves crumpling and curling of young leaves and shoots, thickening them only slightly, and this damage may not be true gall formation.

TABLE 4.4. Classification (superfamilies, families, subfamilies and tribes) of aphids according to Heie (1980) and Blackman & Eastop (1994), with estimates of numbers worldwide (from Havill & Foottit, 2007, for the Adelgidae) and commonness of galling in each group, with the primary host plants indicated (modified from Blackman & Eastop, 1994; Wool, 2004; Miller, 2005) (++ common; + rare [some of the second category may not be true galls])

Unlike psylloids and scale insects, aphids are predominantly a northern temperate group with few species in the tropics and subtropics. There are whole families of tropical plants that are almost immune from attack by aphids. It seems that the typical aphid lifecycle, involving two host species and parthenogenetic generations alternating with a sexual egg-producing generation resistant to low temperatures (see below), is an adaptation to seasonal weather patterns and cold winters, and is not well suited to tropical climates. The tropics may have prevented aphids from reaching southern temperate regions, which also have seasonal climates and cold winters; their indigenous aphid faunas are very small.

Forty per cent of aphid species occurs on trees for part or all of the lifecycle, specialising on the more ancient families. For example, in Britain, they specialise on conifers (Pinaceae), elms (Ulmaceae), beeches and oaks (Fagaceae), birches (Betulaceae) and poplars (Salicaceae), and elsewhere also on laurels (Lauraceae), Styrax (Styracaceae), Hamamelis (Hamamelidaceae), Caryx (Juglandaceae) and Pistacia (Anacardiaceae). Of the remaining 60 per cent, many belong to the more recent subfamily Aphidinae, which has diversified to a large extent on more recent herbaceous plants (e.g. sedges, Cyperaceae; rushes, Juncaceae; grasses, Poaceae; and composites, Asteraceae). It seems likely that the Adelgidae, Phylloxeridae and Aphididae first appeared before the evolution of herbaceous plants and that gall formation in aphids is an ancient feature too.

Aphids are small and soft-bodied and feed, as in most Hemiptera, by sucking the sap of plants through elongated stylets. Most species tap the phloem sieve tubes or phloem parenchyma cells, and a few with short stylets feed on cortex and mesophyll cells nearer the surface of the stem or leaf. The stylets are very long in species living on rough bark where they have to reach the bottom of deep crevices and penetrate thick layers of cork (e.g. the eriosomatine Prociphilus species on conifers).

The main characteristic of aphids, which distinguishes them from other Hemiptera, is their complex lifecycle that involves alternation of sexual and asexual (parthenogenetic) phases. Sex in aphids is odd. Both males and females are diploid (each with cells with two sets of chromosomes) except that males have one sex chromosome less than females–she is XX and he is XO. Males are produced parthenogenetically from a diploid egg in which a specialised meiosis has affected just the sex chromosomes, resulting in a single X. A diploid that has not gone through this reduction division becomes a female, also produced parthenogenetically. Males and sexual females are usually produced in late summer or autumn, triggered by decreasing temperatures and day length (it is the increasing period of darkness that is critical). Mothers continue to produce females at this time too; they always have fewer sons than daughters, in the ratio of 1:3 or less. In the Fordini, sexual reproduction occurs in the spring when daylength is increasing (but probably is triggered in the previous autumn when the parents of the males and females, the sexuparae, form). Males and the sexual females produce sperm and eggs that are haploid, each with one set of chromosomes and with one X chromosome, and sperm developing without an X chromosome degenerate. So, sexual reproduction involving fertilisation of egg by sperm always produces females, which have received an X chromosome from each parent.

FIG 39. The generalised lifecycle of a holocyclic aphid species involving two host plants (modified after Heie, 1980). Part of the lifecycle is on the primary (1º) host and part on the secondary (2º) host.

The basic aphid lifecycle is the ‘holocycle’ (Fig. 39). This starts with a fertilised egg that overwinters, hatches in spring into a nymph that develops into the adult fundatrix, always female and usually apterous (without wings). This individual is the foundress of all the parthenogenetic, viviparous generations that appear during the summer–females producing more females without mating (i.e. thelytokous parthenogenesis) or laying eggs. Usually in autumn, males and females (the sexuales) appear. They mate and the female lays a fertilised egg that overwinters, and develops into the fundatrix of next year’s generations. Each sexual female produces very few haploid eggs, a trend that is taken to extremes in the Eriosomatinae, which lay just one egg half the size of the female’s body. The holocyle involving cyclical parthenogenesis (‘heterogony’) is a basic, primitive feature of aphid biology. Egg-laying is a primitive feature too and, in the Adelgidae and Phylloxeridae, both sexual and parthenogenetic females are oviparous. In the Aphididae, parthenogenetic females are viviparous, producing live young, due to the production of hormones that suppress the development of eggs. Many of these females are apterous as well as being viviparous. Both adaptations speed up development so that production of females can proceed very rapidly, taking advantage of plentiful food and favourable temperatures. Aphids with the most complex lifecycles alternate between two host species (they are ‘heteroecious’). The sexual, egg-laying generation occurs on the primary host, a tree or shrub, and the parthenogenetic generations on the secondary host, a totally unrelated and often herbaceous species. Thus, two periods of migration with winged adults are necessary to complete the lifecycle, which might take one year (as in the Aphidinae, Hormaphidinae and most Eriosomatinae) or two years (in most Adelgidae and most Eriosomatinae: Fordini). Most non-galling aphids, i.e. the majority, and a few gall causers (probably most Phylloxeridae), are ‘monoecious’: the sexual and parthenogenetic generations occur on one host species. This reduces the need to migrate and so reduces mortality–many aphids are blown about haphazardly and fail to land on a suitable host plant.

FIG 40. The lifecycle of an anholocyclic aphid species (after Heie, 1980).

In some aphids, the sexual part of the lifecycle is lost so that all generations consist of parthenogenetic females, i.e. they are ‘anholocyclic’ (Fig. 40). In these, adults or nymphs overwinter. Most anholocyclic species are not gall causers; Adelges abietis, which causes pineapple galls on spruce Picea abies, is a notable exception (see Chapter 6). Some species are entirely anholocyclic with no sexual individuals and live on the secondary host. Others are holocyclic in cold temperate regions and anholocyclic in warmer parts of their range. A few species keep their options open, producing sexual individuals at the same time as parthenogenetic ones.

Besides these complexities of lifecycles, aphids are renowned for their polymorphism, i.e. multiplicity of forms (or morphs). Several female morphs have already been noted:

The fundatrix–wingless, with high fecundity.

Summer parthenogenetic females–oviparous or viviparous, winged or wingless.

Autumn sexual females–oviparous, with low fecundity, produce haploid ova, wingless.

Additional morphs occur in some species and males can be polymorphic too:

Soldier females–defend and clean the colony in some gall-causing species, may be sterile.

Intruder females–enter the gall of another population (of the same or different species) in order to take it over.

Males–dwarf or full-sized, winged or wingless.

Production of different morphs is due to hormones, controlled by the genetics of the species, and by environmental influences. This polymorphism has led to the evolution of the variety of lifecycles; see Moran (1992) for further discussion of this topic.

BOX 4.6. Biology of scale insects (Information from Beardsley, in Ananthakrishnan, 1984; Meyer, 1987; Gullan & Kosztarab, 1997; Cook & Gullan, 2004; Tagaki, in Raman & Gupta, 2007; ScaleNet website)

Scale insects belong to the Coccoidea, the most varied superfamily of the Hemiptera: Sternorrhyncha and the largest–there are about 7350 species worldwide. Although classification of the group is in a state of flux, it seems that about ten families out of a total of about 28 contain gall-causing species (Table 4.5 shows a conservative classification; some families, e.g. the Eriococcidae, are now divided into several new families).

TABLE 4.5. Families of Coccoidea, with examples of gall-causing genera (using a conservative classification, based on Beardsley, in Ananthakrishnan, 1984). Some families have more recently been split into several new families) (–indicates no galling species)

The gall-inducing habit occurs sporadically in unrelated families and only a few families, e.g. the Asterolecaniidae, Eriococcidae and the Beesoniidae, contain a high proportion of galling species. Most Asterolecaniidae cause simple pit galls, while most galls of the Beesoniidae are complex: elaborate globular or cylindrical twig galls or bud-like structures, up to 4 cm long, with their openings sometimes temporarily closed. Males and females may cause different galls, too. Most species are highly host-specific. Diversity of coccoids is high in tropical and subtropical climates especially in the southern hemisphere, e.g. in Brazil, southern Africa and, particularly, Australia, where scales have diversified richly on eucalypts. Coccoid galls are rare in Britain, where only Asterolecaniidae galls occur.

Scale insects are small (< 5 mm long) and, superficially, the structure of all of them is similar so that they can be difficult to identify. Adult males and females are totally different in appearance. The males are delicate, with one pair of wings, no functional mouthparts, and live for a very short time. Females are neotenic, i.e. they look like the nymphs, are scale-like and without wings (‘neoteny’ = precocious development so that juveniles become sexually mature). They are tougher than the males, often protected by waxy or scaly coverings, and survive for several months, usually remaining immobile in one spot on the plant. The first instar nymph, called a crawler, has longer legs than older nymphs and is relatively active; it can disperse some distance by walking or being blown by the wind. Some species interact closely with ants, the crawlers clinging to new queens as they disperse to found new colonies. In the gall-causing genus Cystococcus (Eriococcidae) on Eucalyptus, the female crawlers hitch a lift on the backs of their winged brothers as they leave the parental gall. In some Beesoniidae, the first instar male nymphs are polymorphic: some are normal crawlers that develop into fertile males, while others have enlarged legs with long sharp claws and act as soldiers, defending the colony. These soldier males probably do not become sexually mature and most do not survive beyond the first instar.

Mouthparts and feeding in females and in nymphs are typical of the Hemiptera. Mandibles and maxillae are elongated into fine stylets, which penetrate between cells to reach the phloem of the vascular tissues (see Box 4.4). Stylets of species feeding on thick bark or roots, where the phloem is deep beneath the surface, tend to be very long, much longer than the length of the body; in gall causers on leaves, they are usually shorter.

The lifecycle of most gall-causing and free-living scale insects starts as an egg, apart from Asterodiaspis and some species of Diaspididae which are ovoviviparous (i.e. the egg hatches just before being laid so that it is the first instar that is deposited) and some viviparous species (e.g. Apiomorpha and Gallacoccus). Clutches of eggs are protected from predators and from inclement weather by wax layers or by the scale-like body of the mother. The crawlers hatch and escape from the nest and disperse to new sites; this may not be very far, so leading to aggregations of individuals. Male coccoids, typically, have four instars while females have only two; development can be shorter in females as they have no wings. Males tend to appear first; they fly to the females, often mating before the females have emerged from their galls. The female then lays a clutch of eggs, often still within her gall, and crawlers hatch and disperse to start the next generation. Reproduction in most species is sexual, with diploid females and haploid males–caused by the loss of one set of parental chromosomes rather than by the absence of fertilisation. Some species, such as Asterodiaspis species on oak (Quercus) in Britain, are parthenogenetic (i.e. thelytokous parthenogenesis), and males are unknown. Gullan & Kosztarab (1997) give references to descriptions of several other types of parthenogenesis; sexual arrangements in many scale insects are odd.

Psylloids and scale insects are predominantly tropical and warm temperate groups and their gall diversity in these areas is high, especially in the southern hemisphere. Aphids are cool temperate insects, especially abundant in the northern hemisphere, and their complex lifecycles may have evolved as an adaptation to seasonal climates with cold winters (see Box 4.5).

SAWFLIES (HYMENOPTERA: SYMPHYTA)

Gall-causing in sawflies is unusual (see Box 4.7 for biology and classification) and most galling species are similar to their free-living relatives–with similar lifecycles, with sexual reproduction and egg-laying females, and with caterpillar-like larvae. Unusually for gall insects it is the ovipositing female that initiates the gall. Before depositing an egg, she stings the leaf (and most sawfly galls develop in leaves) often several times in a particular pattern, and this results in a leaf fold or roll or a small swelling. This provides the larva with a shelter and a supply of food, and is prepared before the larva hatches–although in the more advanced Eupontania species, feeding larvae are required before the galls reach their full size and shape. These shelters form a range of structures, some of them true galls but some probably not. Blennocampa phyllocolpa causes simple rolls on roses and some Phyllocolpa folds and rolls are not thickened. These may not be true galls. But the folds and rolls of other Phyllocolpa species are distinctly thickened and, like the beans and balls of Pontania and Eupontania and the swollen stems and buds of Euura, they are more or less constant in size and shape and are definite galls. The young larvae of Phyllocolpa feed inside the folds, though not always on the thickened tissue; they may leave them often before they are full-grown, and continue to feed outside on ungalled parts of the same leaf or on different leaves.

Apart from Blennocampa, all of these genera are nematine sawflies (belonging to the subfamily Nematinae of the family Tenthredinidae; Blennocampa belongs to the subfamily Blennocampinae) and they all specialise on willows and poplars (Salicaceae). Willows Salix are particularly rich in species in northern Europe and this is reflected in the variety especially of Pontania and Eupontania galls, with each willow species tending to have its own sawfly species, and perhaps more than one. Sometimes it is easier to recognise a nematine sawfly from its gall and host plant than from its adult (nematine adults are notoriously difficult to identify). Pontania, Eupontania and Euura galls are closed and are included in Chapters 8 and 9; the leaf rolls of Phyllocolpa are included in this chapter.

BOX 4.7. Biology of sawflies (Information from Wagner & Raffa, 1993; David Sheppard, pers. com., 2003)

Sawflies belong to the Symphyta, the more primitive suborder of the Hymenoptera, whose adults have no ‘waist’ between thorax and abdomen (unlike the larger suborder, the Apocrita, which includes the ants, bees and wasps). The Symphyta contains six superfamilies with 14 families (Table 4.6) and only two of the families contain gall causers. The saw-like ovipositor of the female is characteristic, more robust than the needle-shaped implement or the sting of the Apocrita.

Sawflies occur in every continent, except Antarctica, and are most diverse and numerous in high and low latitudes, particularly in the arctic and north temperate regions and the humid tropics and subtropics. The fauna is poor on isolated islands and in Australia (although there the Pergidae has diversified into species found nowhere else), probably because adult powers of dispersal are poor–they are weak fliers and are short-lived. In Britain, northern Europe and northern North America, the subfamily Nematinae (family Tenthredinidae) is diverse and includes the genera Phyllocolpa, Pontania, Eupontania and Euura, particularly associated with galls on willows Salix.

Most sawfly larvae are plant-feeding caterpillars, similar to the larvae of Lepidoptera, and the majority feed externally on leaves. The rest (about 15 per cent) feed inside plants and most of these are mono-or oligophagous (i.e. they feed on just one or on a few related species)–about 3 per cent are stem and twig borers, 6 per cent are leaf miners and 6 per cent are gall causers. The adults mainly feed on pollen, nectar and honeydew.

TABLE 4.6. Number of genera and species in sawfly families (worldwide) with notes on the food of their larvae (from Smith, in Wagner & Raffa, 1993; David Sheppard, pers. com., 2003; Roininen et al., in Raman et al., 2005)

The typical sawfly lifecycle involves an egg, five or six larval instars (one less in the male), a prepupa, pupa and adult. The prepupa develops from the final larval instar (there is no moult between the two); it does not feed and is usually different in appearance and colour from the larva. It finds a suitable site in which to pupate, usually in the soil or in plant litter, and makes a cocoon or earthen cell; it overwinters inside, pupates in the spring and the adult emerges in spring or early summer. Most species have one generation a year; a few have two or more and species with wood-boring larvae, e.g. the Siricidae, may require four years to become full-grown. Most adults live only a few days, during which time they mate and the females insert their eggs one at a time into slits in leaves or stems of the larval food plant, cut by the saw-like ovipositor. The larvae hatch a week or two later and feed on leaves, buds or stems, sometimes in leaf rolls, mines or galls, or boring into wood, and feeding either singly or in groups.

Males and females usually occur in more-or-less equal numbers and most reproduction is sexual. Sawflies are haplodiploid like other Hymenoptera, females being diploid and produced from fertilised eggs and males haploid, developing from unfertilised eggs (arrhenotokous parthenogenesis). In a few species, males are unknown or rare and, in these, other types of parthenogenesis occur.

THE GALL MIDGES (DIPTERA: CECIDOMYIIDAE)

For classification of the Cecidomyiidae and background information on its biology, see Box 4.8. Gall midges cause a greater variety of galls than any other group. They occur on all types of plant, but reach their maximum diversity on flowering plants. They gall buds, flowers, fruits, seeds and stems, and occasionally roots, and are most diverse in leaves. The galls are leaf folds and rolls (this chapter), or pits, blisters and pouches on leaves and stems (Chapter 5), rosettes and artichokes in buds (Chapter 6), or closed swellings in leaf veins, petioles and stems (Chapters 8 and 9). Gall midge pouch galls are probably the most variable of any group, ranging from simple invaginations similar to those of eriophyoid mites to elaborate projections with a complex structure, incorporating several layers of tissues including a woody layer. Most contain nutritive tissue in patches or in a layer lining the gall chamber. The ‘ambrosia galls’ are exceptional, with a fungus involved in the development of the gall as well as the plant and the larva. Ambrosia galls are closed swellings in stems, buds or fruits, with a cavity lined with the mycelium of a fungus, and this may provide food for the larvae (Chapters 8 and 9). Ambrosia galls may be primitive, with the larvae maintaining the fungus-feeding habits of ancestral gall midges (see Box 4.8), or they may have secondarily become associated with fungi. This is debatable; gall midges in ambrosia galls are not known for sure to feed on the fungi.

BOX 4.8. Biology of gall midges (Information from Skuhravá et al., in Ananthakrishnan, 1984; Redfern, 1975; Gagné, 1989, 2004; Harris, in Williams, 1994)

Gall midges belong to the Cecidomyiidae, one of the largest families of Diptera, the two-winged flies. About 5500 species are known worldwide, but many more species await discovery. They occur all over the world, in tropical and temperate climates, though the faunas of Africa and Australia are less well known than those of the rest of the world. The family name comes from the Latin cecidium = gall, and emphasises the importance of gall formation in the group.

FIG 41. A cecidomyiid larva, ventral view, with sternal spatula (arrowed) (from Redfern et al., 2002, with permission).

Adult gall midges are small and fragile, usually less than 5 mm long, with long legs, reduced wing venation and distinctive antennae (like a string of beads). The larvae are small maggots (Fig. 41), usually no more than 3 mm long when fully grown, and are white, yellow, pink, orange or red in colour. Most have a sternal spatula on the ventral surface of the thorax, a structure diagnostic of the family (though lost in some species).

TABLE 4.7. Feeding habits of the four subfamilies of the Cecidomyiidae, with approximate percentages of the total numbers of species worldwide (also approximate) for each subfamily (Information from Skuhravá et al., in Ananthakrishnan, 1984; Gagné, 1989, 2004; Harris, in Williams, 1994)

The family probably originated in the Mesozoic era–the oldest fossil is a catotrichine gall midge from late Jurassic or early Cretaceous deposits, more than 150 million years old. The earliest cecidomyiids probably fed on fungal hyphae, like their close relatives, the fungus gnats (Mycetophilidae and Sciaridae). The three more primitive subfamilies, the Catotrichinae, Lestremiinae and Porricondylinae (Table 4.7), are still primarily fungus feeders. The largest subfamily is the Cecidomyiinae, which is mainly phytophagous and evolved rapidly with the flowering plants during the Tertiary (this period started about 65 million years ago) and, by 30 million years ago, its gall fauna is thought to have been similar to that of today. Galling has evolved many times in the subfamily and the 70 per cent of species causing galls is higher than in any other insect or mite group, apart from the gall wasps (Cynipidae, Chapter 9). The variety of galls caused by gall midges is greater than in any other group. The tribes within the subfamily and their feeding habits are listed in Table 4.8; tribes with gall-causing species tend to be the largest. Several tribes include ‘ambrosia galls’ with fungal symbionts on which the larvae may or may not feed; the gall midges cause the galls perhaps with the help of the fungi.

All feeding in gall midges occurs in the larval stage; the adults do not feed. The larvae feed on fluids, on plant sap or the contents of fungal hyphae or animal prey, and ingest it by suction. Originally feeding on fungi in the soil or in leaf litter and rotting wood, adaptation to feeding on living plants may have been via cambial sap flowing from wounds in tree trunks, then on soft tissues in leaf and flower buds, finally leading to gall formation in buds and leaves perhaps aided by fungal infection. Predation evolved later, perhaps arising first on the immobile gall causers; common prey items are aphids, psylloids, scale insects and other gall midge larvae.

Because the larvae are hidden within the plant and the head and mouthparts are so small, it is difficult to observe feeding directly. They do have functional mandibles, but it is uncertain whether or not gall-causing species slice into or pierce plant cells. Few wounds have been observed in the walls of the cells, unlike in the galls of eriophyoid mites. It is possible, in some species at least, that minute slits are made that repair themselves almost immediately, and predatory species may pierce the skin of their prey. In most gall-causing species, the plant seems to be induced to produce sap by the action of the larva. Its head can be retracted into the thorax to form a suction cup that is applied tightly to the plant surface; it then secretes saliva on to the plant cells, which causes the cell membrane to become permeable, prevents the cell wall thickening and removes the cuticle. The saliva also contains digestive enzymes that break down cell contents outside the body of the larva. Then muscular pumps in the thorax suck in the liquefied food. In contrast to the galls of cynipid larvae (see Chapter 9, Box 9.1), the nutritive cells in cecidomyiid galls are not destroyed. Nutrients flow continuously while the larva is feeding, from nutritive cells that are supplied by diffusion from the vascular tissues. Extra-intestinal digestion is a feature of all gall midge larvae including the earliest fungal feeders and was clearly a useful feature that allowed the evolution of phytophagy and the subsequent great diversification of the Cecidomyiidae. Like most gall causers, most gall midges are mono-or oligophagous, feeding on only one species or genus of host plants. Polyphagous species tend not to be gall causers or, if they are, their galls are very simple. The plant families with most cecidomyiid galls are the Asteraceae, Apiaceae, Rosaceae and Salicaceae. Less than 1 per cent feed on ferns and conifers and probably these species have evolved from ancestors on flowering plants.

TABLE 4.8. Foods and habits of the supertribes and tribes within the subfamily Cecidomyiinae, with examples of genera in Britain (after Gagné, 1989, 2004)

The generalised lifecycle of cecidomyiids is simple: an egg, usually three larval instars, a pupa and an adult male and female. All reproduction is sexual (although there are usually more females than males), except in a few unusual non-galling species that can reproduce parthenogenetically in the larval stage. One generation a year is common. The adults emerge in late spring or early summer, the males a few hours before the females. The males fly upwind attracted by a pheromone emitted by the newly emerged females, they mate and the females lay eggs almost immediately. Males die after mating and females soon after laying their eggs; they live only a few hours or a couple of days and do not feed. In phytophagous species, the eggs are laid on or near the spot where the larvae will feed and they hatch in a week or two. The larval stage is the longest and, usually, the full-grown larva overwinters, either in the plant or in the soil, often spinning a cocoon first. It pupates usually in the spring, this stage lasting a few weeks, and the adults emerge ready to mate and with their full complement of eggs already developed. Some species have two or more generations a year and, in a few species, each generation takes two or three years. In some species, different individuals spend varying periods in the soil as full-grown larvae, with adults of one generation emerging each year for up to 12 years.

Like most galls, gall midge galls start in young undifferentiated tissues, such as young leaves and the meristems of buds, and all are initiated by newly hatched larvae. Eggs are laid on or near a bud or a new leaf, they hatch and the hatchling crawls to a suitable spot; it wriggles round in a circle for a short time and starts to feed, sucking from a small patch of cells. Within a few hours the cells nearby have lost their normal characteristics and have reverted to a juvenile state. Beneath the larva they become nutritive (see Chapter 1; but not in ambrosia galls) and those either side start dividing to produce the growth–walls of a pit, or enveloping lips that enclose the larva in a pouch, or the multiple small leaves of a rosette. The gall is usually recognisable within a day or two of the larva starting to feed. In most gall midge galls, an aperture is part of the structure of the gall, although this may close temporarily or, occasionally, seal over permanently. The full-grown larva or emerging adult normally leaves through this hole; in sealed galls, though, a new aperture must develop. Pupation occurs either in the soil or in the gall, and the gall may remain on the plant or fall to the ground.

Compared to aphids, cecidomyiids have simple lifecycles: a male and female and sexual reproduction, one or a few larvae per gall and one or a few generations per year (occasionally, a generation takes two or three or more years). There are four types of lifecycle:

(1) Larvae develop rapidly through their three instars and are full-grown by the start of summer. They drop out of their galls to the soil, where they spin cocoons and pass through the summer, autumn and winter, pupating within them the following March or April, and emerging in late April or May to start the next generation. Some species of this type may have several quick generations during the summer, each pupating in the soil and only the last generation overwintering as fully fed larvae. The galls produced by this type tend to be simple pits, pouches or leaf folds and rolls.

(2) A similar rapid development of larvae as in type (1) but the full-grown larvae remain in the galls until the autumn, when galls and larvae fall to the ground. The larvae overwinter in the gall, pupate the following spring, and adults emerge in April or May. Some of the more complex pouch galls belong to this type.

(3) Larvae, especially the first instar, grow less rapidly over three or four months but are full-grown before the winter. They overwinter in the gall, which remains on the plant, pupate inside in the spring and emerge in May or June. Some of these galls are complex, closed swellings that must be prepared for the emergence of the adult (which cannot bite its way out). The larva may excavate an aperture that it seals temporarily with silk or the pupa may be armed with tough horns with which it bores its way out.

(4) Development of the first instar is prolonged and lasts through ten or more months, including over the winter. Then instars two and three develop rapidly, pupation occurs in the gall and adults emerge in late spring or early summer to start the new generation. These galls include rosettes and some closed swellings. This type can lead to a lifecycle of two years if the third instar is prolonged as well as the first, each surviving over subsequent winters. Some species that cause artichoke galls belong to this category.

THE GALLS

The remaining part of this chapter describes the simpler open galls: erinea, caused by eriophyoid mites, and leaf folds and rolls, caused by gall mites, psylloids, aphids, thrips, sawflies and gall midges. Examples of each are included, arranged from the less to the more complex.

Erinea

Erinea are patches of glandular hairs, sometimes called filzgalls, and induced uniquely by eriophyoid mites; no insect, it seems, has evolved the ability to induce hairs that are nutritive. They are amongst the simplest of arthropod galls. They are usually found on undersides of leaves, occasionally on their upper sides or on stems, buds or fruits; gall mites do not feed on roots. The type of hair, elongate or club-shaped, unicellular or multicellular, is characteristic of the gall and of the particular mite species, and is quite unlike the normal hairs of the plant. Some erinea develop on the flat of the blade, not apparent from the other side of the leaf, while others cause discolouration and a more or less marked doming of the leaf, with the felt of hairs in the concavity beneath (Fig. 42, left). Others run along the leaf veins, picking them out as white lines (Fig. 42, right). In the past, the hairy patches were thought to be fungal and were classified as such by mycologists (e.g. the genus ‘Erineum’). Erinea should not be confused with the hairs characteristic of the plant, which may cover the underside of the leaf with a felt-like layer, or form patches in vein angles, or grow in lines along the veins. These are tomenta or domatia. They consist of non-glandular hairs and often shelter predatory mites or fungus-feeding species, and may do the plant a service by reducing its parasitic fauna. The hairs themselves, though, do not provide food and gall-causing eriophyoids are rarely found amongst them.

FIG 42. (above left) Marked doming of the leaf of walnut Juglans regia caused by the erineum of Aceria erinea (Michael Chinery); (above right) silver lines of erinea of Aceria nervisequa on the upper surface of beech Fagus sylvatica leaves. (Tom Higginbottom)

Erinea are open galls and probably provide less protection than folds, rolls and pouches, although the close density of the hairs and usual position on the underside of leaves creates the equable climate and high humidity that the mites require. Once an erineum is formed, its mites usually remain within it, leaving only when the hairs collapse and the leaves dry out at the end of summer. Erinea caused by a species tend to vary considerably in size, depending on the size of the mite population within; fast-growing populations may induce large patches of hairs, sometimes covering most of a leaf surface and causing the blade to buckle.

Details of the erinea are known for few species. Most involve enlargement (hypertrophy) of selected cells only, without an increase in cell number (hyperplasy). The hairs develop from enlarged and elongated epidermal cells, with characteristics similar to nutritive cells of other galls (Chapter 1), and develop in response to feeding by the mites. Doming and swelling of the leaf is caused by enlargement of cells of the upper epidermis and mesophyll, or may be due to enlargement of the air spaces in the leaf. Erinea range from simple to more complex (Fig. 38, 1–8). In the erineum of Aculus hippocastani on horsechestnut (Aesculus hippocastanum, 3), the nutritive hairs are multicellular. On grapevine Vitis vinifera, the bulging of the leaf blade above the erineum of Colomerus vitis (7) involves enlargement of cells, and thickened, multicellular projections extend into the erineum of Aceria erinea on walnut (Juglans regia, 8; Fig. 42, left). This is a route, depicted in Fig. 38, that suggests how mite pouch galls may have developed from erinea (e.g. Eriophyes viburni on wayfaring-tree Viburnum lantana, 18).

An erineal hair develops from an epidermal cell, at first like any plant hair. Its nucleus and nucleolus enlarge, organelles in the cell multiply and the cell enlarges, and divides in galls with multicellular erineal hairs. This stage is very brief in a normal hair; in a nutritive hair, the stage lasts longer due to the feeding of the mite. The hairs develop in response to feeding, at first by the fundatrix when she becomes active in spring (she is the matriarch of all generations in the gall), and later by her offspring as they develop during the summer. The fundatrix pierces an individual epidermal cell and induces neighbouring cells, perhaps several cells distant, to become meristematic and develop into nutritive hairs. Why some epidermal cells and not others form nutritive hairs is not understood. The hairs are thin-walled and without a cuticle, so are easily pierced by the short stylets of the fundatrix and her offspring. Feeding punctures the cell wall and causes scars, at the tip of hairs when they are very young and not fully elongated, and later near the base. The enlarged tips of the hairs form a protective roof over the feeding mites, maintaining high humidity within the erineum (most gall mites, particularly the juveniles, are very sensitive to desiccation). Once a feeding scar has formed, composed of thick callus, the mite can no longer feed from that spot as the cell wall becomes too thick and tough for its chelicerae to pierce. It moves elsewhere on the same hair or pierces another. Individual hairs provide food only for a short time; soon after they reach their full size, their walls thicken with cellulose and later with lignin, and tannins are deposited in the cell vacuole. Mites can no longer feed from them, and they become brown, shrivel and die and eventually drop off. Most erinea contain a mixture of young hairs on which the mites feed and ageing hairs; the proportion of old hairs increases through the summer, causing the erineum to become increasingly brown or rust-coloured.

Although erinea of different species may look similar to the naked eye, their hairs differ in detail. Eriophyes leiosoma on lime Tilia, E. distinguendus on bird cherry Prunus padus and Acalitus brevitarsus on alder Alnus glutinosa provide examples (Fig. 38: 1, 4, 5). In E. leiosoma, the hairs are cylindrical with rounded tips and are flexible, but otherwise are very like normal hairs on the young leaves, hairs that normally fall off soon after the leaves burst from the bud scales. In most erinea the hairs are unlike any hairs produced by the host, although they begin growing in the same way. In E. distinguendus, the hairs develop as rounded papillae that lengthen into a squat club-shape, with a rounded head on a short stalk. In A. brevitarsus, the hairs are club-shaped with globular heads that later divide into several lobes. The stalks of these hairs contain several nuclei, up to ten in each hair, and may be polyploid, i.e. they contain more than two sets of chromosomes (normal cells of the plant are diploid).

The lifecycles of Acalitus brevitarsus and of Colomerus vitis, which causes erinea on grapevine, have been described and other species may be similar. As temperatures increase and buds swell in spring, the overwintered fundatrix (the deutogyne, see Box 4.1) starts to feed on the very young leaves before they have burst completely from the bud scales. She may explore the still-folded leaf, her wanderings marked by a trail of hairs induced by her feeding, before settling down in one place on the underside. She lays eggs, perhaps one a day, attached to epidermal cells and protected from drying out by the erineum hairs. The eggs hatch in a week or two, and the larvae start to feed so enlarging the erineum and increasing its density of nutritive hairs. Juvenile mites do not usually leave the erineum of their birth; they are delicate, not very mobile and are prone to desiccation. In favourable summer temperatures, development to adult after hatching takes 10–14 days (in Colomerus vitis). The new females (protogynes, Box 4.1) are fertilised by sperm from male spermatophores; they lay eggs, and several generations develop during May and June. So the size of the erineum increases and new ones develop as adult females disperse to new young leaves. In late summer or early autumn, most nutritive hairs have browned and shrivelled. The only individuals that will survive the winter are the fertilised females (deutogynes), which disperse to overwintering sites under bud scales and become the fundatrices of new erinea the following year.

Leaf fold and roll galls

Folds and rolls tend to provide more protection from the elements and from predators than erinea. They are induced in the youngest leaves as they burst from the bud, often not yet unfolded, unrolled or expanded–often, this juvenile state is retained in the gall. Some of these galls are narrow folds or rolls of the leaf margin, while others are broader, affecting much of the leaf blade. All involve enlargement of cells of one surface of the leaf relative to the other so that the leaf remains folded or rolled. The most elaborate approach the complexity of pouches, or affected leaves may be bunched together into rosettes, and a few become temporarily or permanently closed. Fold and roll galls are caused by eriophyoid mites and a variety of insects. In psylloids, aphids and gall midges, the newly hatched nymph or larva initiates the gall when it starts to feed, while in mites and thrips, the adult female starts the formation of the gall and her young maintain and enlarge it. In Phyllocolpa sawflies, and probably in other leaf-folding sawflies too, the ovipositing female causes the leaf fold or roll; it will form its characteristic shape even if an egg is not laid.

FIG 43. Rolls of leaflets of dog rose Rosa canina caused by Blennocampa phyllocolpa; usually several adjacent leaflets are affected. (Michael Chinery)

The simplest folds and rolls are not very different from moth caterpillar rolls held in place by silk, or the distorted leaves of young shoots caused by large colonies of aphids in spring. Psylloids, sawflies and aphids cause these simplest of galls and live and feed in a manner similar to their free-living relatives; no differentiation of tissues occurs and some are hardly thickened. Slightly more complex are folds and rolls with rudimentary development of nutritive tissue but with little other tissue modification, and these are caused by some gall mites, thrips and gall midges. The most complex have a well-developed nutritive tissue of more than one layer of cells and modifications of other cells, sometimes with a layer of sclerenchyma (woody) cells, and are caused by some species of all the galling groups mentioned, except sawflies.

The rose leaf-rolling sawfly Blennocampa phyllocolpa causes simple downward rolls of young, fully expanded leaves of wild roses Rosa (Fig. 43). The rolls are not appreciably thickened and this species may not be a true gall causer (even though it is included in most books on gall identification). The Phyllocolpa rolls and folds on willows Salix are usually slightly thickened and probably are true galls, though simple ones (Fig. 44) may be borderline. The gall on arroyo willow Salix lasiolepis in Arizona, USA, and the Phyllocolpa species that causes it have been studied more intensively than most, and the lifecycle and effects on the host plant seem to be typical of the genus. In spring, the female repeatedly stings the underside of leaves with her ovipositor. She works down the leaf from the tip to the base 1–2 mm from the margin and, at the base, inserts one egg into the lower epidermis. Almost immediately the affected leaf margin folds downwards, due to enlargement of the cells of the upper epidermis and mesophyll. The stimulus for the folding is probably due to a substance injected by the ovipositor; the leaf will fold whether or not an egg is laid. A week or two later, the egg hatches and the larva feeds on leaf tissue inside the fold, growing and moulting through five or six instars. In its final instar, it leaves the fold and eats the unaffected leaf in a manner similar to free-living sawfly larvae. There is normally just one larva per leaf fold and a single leaf provides all the food that it needs. When fully fed, it drops to the soil and spins a cocoon in which it overwinters; it pupates in the spring and the adult emerges as the new buds of willow are swelling. The adults mate and the lifecycle starts again. In most Phyllocolpa species there is one generation a year.

FIG 44. Folds of the leaf margin of willows (Salix spp.) caused by Phyllocolpa species (from Redfern et al., 2010, with permission).

Like causers of more elaborate galls, Phyllocolpa species are selective in their choice of food. Besides being restricted to one or a few closely related host species, particular individual trees and some leaves within trees are more attractive than others and are less resistant to attack. In Salix, shorter leaves tend to be more resistant and females prefer to lay eggs in longer leaves on long shoots. These shoots are faster growing, provide more food for a larva and produce fewer defences against the herbivore. On S. lasiolepis, leaf fold galls are commonest half way down the shoot, not at the tip. The number of galls will be higher on these ‘best’ leaves although, when the population of females is high, some will be forced to lay eggs in less desirable leaves or on more resistant host plants where subsequent survival of the larvae will be reduced.

FIG 45. Leaf roll galls of Trioza alacris on bay laurel Laurus nobilis. (Michael Chinery)

The thickened leaf rolls and curls of some psylloids and aphids are, perhaps, more deserving of the description ‘gall’ than the Phyllocolpa folds, even though there is no multiplication of cells or differentiation of tissues. Trioza alacris causes a roll on bay laurel (Laurus nobilis, Fig. 45) and the bright yellow or red rolls on apple (Malus, Fig. 46) are caused by the rosy apple aphid Dysaphis plantaginea. In T. alacris, the female feeds in early summer on the underside of young, expanded leaves and lays several eggs, each inserted between two cells of the lower epidermis. These hatch into waxy nymphs that start feeding, inducing the edge of the leaf to roll downwards; after three days the gall is full-sized, an enclosed tube involving at least two complete turns of the leaf margin (Fig. 47). The galls of D. plantaginea develop quickly too, appearing 24 hours after first instar nymphs have started feeding on the undersides of young apple leaves. The aphids feed on phloem cells, injecting substances that cause the rolls; these substances are transported in the phloem to affect leaf growth some distance away and may have been previously derived from the plant by the aphids. Other apple aphids, non-galling species, tap the phloem cells while feeding on the outside of the gall and grow larger and develop faster than when feeding on normal leaves. It seems that the sap in the gall is richer and remains suitable for longer than in ungalled parts of the leaf. The adults of T. alacris appear in the summer and escape when the coils of the gall loosen, and there may be two or more generations a year. T. alacris was introduced into southern England from Mediterranean Europe with its food plant; it is common on garden bay trees and is spreading steadily northwards.

FIG 46. Leaf roll galls of Dysaphis plantaginea on apple Malus. (Michael Chinery)

A few galls caused by Diptera are simple leaf folds or rolls. The larva of Chirosia grossicauda (Diptera: Anthomyiidae) causes a down-rolling of the tip of a bracken pinnule (Pteridium aquilinum, Fig. 48) and has an effect similar to a leaf miner. An egg is laid on a pinnulet (the smallest segment of the frond) near to its tip. The larva hatches and feeds initially on the lower epidermis and mesophyll next to the central vein of the pinnule. This causes the upper epidermal cells of the central vein to continue to grow while the lower ones do not, causing the pinnule to curl downwards. At first the larva mines within the plant tissue; later, the lower epidermis ruptures and the larva behaves more like a free-living larva than a gall causer.

FIG 47. Sections through the roll gall of Trioza alacris on bay laurel Laurus nobilis (a) one day, and (b) three days after oviposition (after Meyer & Maresquelle, 1983). An egg (arrowed) can be seen in (a) and in (b), a nymph (black) is feeding on the innermost coil.

FIG 48. Gall of Chirosia grossicauda on bracken Pteridium aquilinum. (Tom Higginbottom)

FIG 49. Roll gall of Epitrimerus trilobus on elder Sambucus nigra. (Tom Higginbottom)

Usually each roll contains one larva; if there are two, the roll is proportionally larger. Pupation occurs in the roll from late June onwards, and pupae may drop to the ground and overwinter there, until the adults emerge in the spring.

A second group of leaf fold and roll galls is slightly more complex in that they contain nutritive tissue. Leaves are attacked when very young, still with their margins enrolled, and the galling process prevents them from unrolling. Four examples are included, caused by gall mites and gall midges on elder, hawthorn, apple and pear trees.

Gall mites induce tight rolls in leaf margins of elder Sambucus nigra and hawthorn Crataegus monogyna. In April, new young elder leaves appear with the margins of leaflets naturally tightly enrolled upwards. Feeding by female Epitrimerus trilobus maintains the rolls; they are either short or can extend all round the leaflet (Fig. 49), and the blade may become crumpled and its width reduced if infestation is severe. The mites attack the new leaves for as long as they appear in spring. The roll involves 1 to 1½ turns of the leaf margin with several mites enclosed (Fig. 50). The anatomy of this gall is simple, with no multiplication of cells, and the palisade and spongy mesophyll layers of the normal leaf can still be recognised. The cells of the upper epidermis are reduced in size with scattered cells dead and collapsed after being sucked dry by the mites. Cells of the palisade mesophyll are smaller than usual too, and these two layers form the nutritive tissue, which develops in patches where the mites are feeding. Outwardly similar is the tight upward roll caused by the gall mite Phyllocoptes goniothorax on hawthorn (Fig. 51). Its structure, though, is quite different–inside the roll is an erineum of club-shaped hairs on which the mites feed. This mite causes a normal erineum also, on the under surface of the leaf; the marginal roll is a modification of this, probably giving the mites more protection, and is much commoner. As in most fold and roll galls, modification of tissues is greatest in the centre of the gall and declines as it merges with the ungalled leaf, and is maintained only if the mites remain alive and active inside.

FIG 50. Sections through the marginal roll gall of Epitrimerus trilobus on elder Sambucus nigra (modified after Westphal, 1977a). (a) A diagrammatic comparison of the natural and galled marginal rolls at early and later stages of development; normal leaf on left, galled leaf on right, with mites.

(b) A plan of the galled part of the leaf. Nutritive tissue (stippled); gall mite (cross-hatched); necrosed cells sucked dry by feeding mites (black spots); leaf veins (open circles).

FIG 51. Roll galls of Phyllocoptes goniothorax on hawthorn Crataegus monogyna. (Michael Chinery)

Looser, upward marginal rolls are caused by gall midges on leaves of apple and pear trees: Dasineura mali on crab apple Malus sylvestris, cultivated apple M. domestica and their hybrid, and Dasineura pyri on cultivated pear Pyrus communis. These rolls are wider than mite rolls, usually more than 3 mm wide and sometimes reaching the midrib. Both gall midges gall the newly developing leaves and sometimes are common enough to adversely affect growth of the trees. The gall midges’ lifecycles are similar with several broods a year. Females lay red eggs between the folds of young leaves as they are bursting from the bud, depositing up to 35 eggs per leaf. The larvae hatch in a few days and begin feeding not far from where they were laid, preventing the leaf from unfurling so that a tube forms around them. At first, the rolls may be bright red (on leaves in the sun); later, after the larvae are full-grown, they blacken, become dry and brittle, and affected leaves drop early. Larval growth takes two to five weeks in the summer generations, depending on temperature. Full-grown larvae drop to the soil, spin cocoons and pupate inside, and the adults emerge 10–14 days later. The females must disperse some distance to find new leaves, which become rarer as the season progresses. The final generation each year, and sometimes a proportion of previous generations, overwinter as larvae in their cocoons in the soil, pupating in spring and emerging as the new buds begin to swell in late April or May. The adults and larvae of both species look very similar, but are clearly distinct as neither can survive on the other’s host. Both are European species that have been introduced accidentally into North America and New Zealand where they are serious pests in apple and pear orchards.

Most leaf fold and roll galls are structurally more complex than those so far described, and involve both multiplication and enlargement of cells. They have well-developed nutritive tissue or enhanced vascular tissue, and sometimes include layers of protective woody cells. A variety of eriophyoid mites, psylloids, aphids, thrips and gall midges provides examples.

The eriophyoid mite, Phytoptus tetratrichus, causes a narrow, thickened roll of the leaf margin of lime (Tilia, Fig. 52). The rolls may be down-or upward and short or long, sometimes affecting the whole margin. They are the same colour as the rest of the blade at first, later becoming pale yellow-brown and with a ridged surface. Tilia leaves are simply folded in the bud, without rolled margins, and so, unlike Epitrimerus trilobus and Phyllocoptes goniothorax, this mite must provoke a roll in the leaf margin as it expands. Its roll is more highly organised, too, with marked differential growth between the outer and inner surfaces (Fig. 53). Two or three layers of cells develop into nutritive tissue, the epidermis on the inside of the roll and one or two layers of the mesophyll beneath. These cells collapse and die after the mites have pierced them and sucked them dry. The outside of the roll consists of loosely packed rounded cells with large air spaces, and the outer epidermis forms projections, the ridges on the outer surface of the gall. Nutritive tissue only develops on the innermost surface of the roll. Further out, and as the gall matures, the inner surface produces tough, lignified hairs that block the entrance to the roll. These are similar to the hairs in the aperture of pouch galls (see Chapter 5) and probably help to prevent the entry of predators and other unwelcome visitors.

FIG 52. Leaf roll galls of Phytoptus tetratrichus on Tilia (from Redfern et al., 2002, with permission).

FIG 53. Section through the leaf roll of Phytoptus tetratrichus on Tilia species (modified after Westphal, 1977a). Nutritive tissue (stippled); leaf veins (open circles); tough hairs block the entrance; ridges on outside of roll made up of enlarged epidermal cells and air spaces.

The woolly aphid Eriosoma ulmi (Hemiptera: Aphididae: Eriosomatinae) causes a much larger and looser, thickened, downward roll gall on elms (Ulmus, Fig. 54), involving all of one side of the leaf blade (Malpighi in 1687 appropriately likened this to the colon, i.e. the large intestine). The gall is the same colour as the leaf at first, later becoming paler and yellowish green, and the aphids inside are waxy, dark green when young, bluish grey or brown when older. Most of the cells in the leaf enlarge, particularly those of the upper epidermis, causing the thickening and downward curl, and differentiation between the palisade and spongy mesophyll is lost–all cells become rounded and the air spaces between them disappear. There is little hyperplasy (cell multiplication) except in the leaf veins, which enlarge producing new xylem vessels. No nutritive tissue develops; the aphids suck sap directly from the enhanced vascular tissues. Because the position of the upper and lower epidermal layers is reversed, each develops characteristics of the other–stomata usually on the under surface develop in the upper epidermis, though they have abnormal guard cells, and these probably do not function properly. Elms are the primary hosts of E. ulmi, the fundatrices causing the galls in newly expanding leaves in early summer (see Box 4.5 for a general description of aphid lifecycles). The aphids multiply and produce winged individuals in June and July, and these migrate to the secondary hosts, black-and redcurrants (Ribes nigrum and R. rubrum), and feed on the roots without causing galls. In September to November, the sexuparae fly back to the bark of elms, produce the sexuales, which mate and lay eggs that overwinter, and the fundatrices hatch in spring ready to start the next generation.

FIG 54. Leaf roll gall of Eriosoma ulmi on a leaf of wych elm Ulmus glabra. (Michael Chinery)

An upward roll of the leaf margin of buckthorn Rhamnus catharticus is caused by a psyllid Trichochermes walkeri. The leaf blade is markedly thickened, with many enlarged veins, and curls upwards due to the increase in size of the cells of the lower epidermis. T. walkeri has one generation a year. Small orange eggs are laid in autumn, deposited on buckthorn twigs close to the dormant buds, where they overwinter. They hatch in May and the first instar nymphs feed near the margin of the new leaves, and the margin rolls around each to enclose it (usually there is just one nymph per gall). Only the first instar nymph can induce the gall. As the leaf expands so the gall enlarges and thickens. It grows fastest in June, considerably later than the ungalled leaves, and is full sized by July. This extended period of leaf growth ensures that suitable food is available throughout the nymphal period (there are five instars), so compensating for the reduction in food quality common in deciduous trees and shrubs during the summer. Adults emerge from the galls from late July to mid-September, a week or two earlier if the summer weather is unusually warm, and mate and lay the overwintering eggs.

The gall midge Dasineura odoratae causes thick, pale-coloured, hairy, upward leaf rolls on sweet violet (Viola odorata, Fig. 55). These contain one or many larvae that are white when young, yellow-orange when older. Females lay eggs on the upper surface of the young leaves when they are still rolled, having just burst from the bud. In these rolls, humidity is relatively high, necessary for the eggs, which dry up if it drops below 80 per cent. Feeding by the hatchlings prevents the leaves unrolling and stimulates development of a layer of nutritive cells, several cells thick, on the inside of the roll. The distinction between palisade and spongy mesophyll disappears, too. Vascular tissue increases and the overall number of cell layers is three times that of the normal leaf.

FIG 55. Leaf roll galls of Dasineura odoratae on sweet violet Viola odorata. (Michael Chinery)

As in all gall midges, only the newly hatched first instar larva is capable of inducing a gall. If the eggs fail to hatch, the leaf opens normally. If the larva is removed to another rolled leaf after it has started to feed, it can no longer induce a gall and it cannot roll a leaf that has opened. D. odoratae lays clutches of eggs (up to 60) and the resulting gall varies in size depending on the number of surviving larvae (usually a dozen or so). The larvae grow quickly through three instars. There are several generations throughout the summer–new leaves are produced continuously and are available for galling over an extended period. Three or four generations are common in England and France, more in the south and fewer further north. Full-grown larvae spin silken cocoons in the gall and pupate inside; the pupal stage lasts 10–14 days in the summer and six or seven months over the winter. The adults live for a very short time: they all emerge together, mate immediately and the females lay their eggs, all fully developed at emergence.

The size of the galls of D. odoratae varies and depends on several factors. The number of larvae per gall has the greatest effect, even though the size of full-grown larvae and pupae (and therefore the egg load, the fecundity, of the female) decreases as numbers increase. There is a general tendency for the fecundity of females to decline over the summer generations, too. The growth of the plant also affects the size of the galls. Vigorously growing plants tend to support more galls but, on weaker plants, the galls are often larger suggesting that vigorous plants can partially ‘control’ the effects of the gall midge. Galled plants are generally slower growing than ungalled plants and their leaves are thicker; often they produce fewer leaves, too. The effect on the plant can be severe, with successive heavy infestation throughout the growing season leading to additional galls developing on the flowers and failure to produce viable seed, and sometimes this results in the death of the plant. After the adults have emerged, the galled leaves wither and decay, sooner than healthy leaves. Fungi easily infect them because there is no waxy cuticle to protect the upper leaf surface. Fungal spores may be present in the young gall too, but are prevented from germinating by the galling larvae.

A final group of insects that causes leaf fold and roll galls is the Thysanoptera, though none of these galls is British. Some are simple marginal folds. Others are more complex rolls that affect the whole leaf and contain distinct layers of nutritive and woody strengthening tissue. All are initiated by newly mated females in new young leaves bursting from the bud–the young leaves are still folded and not yet with fully differentiated tissue layers. As in other groups, synchrony between dispersal of the thrips and bud burst is crucial. One or a few females start to feed on a leaf, along the midrib or margin or on the blade, and the more females there are, the greater the effect on the leaf. The initial injury induces the epidermis and the superficial mesophyll to multiply and form nutritive cells, forming patches or a layer a few cells thick. Outer cells of the leaf enlarge, causing a fold or roll around the feeding thrips or preventing the young leaf from unfolding. Cells are packed tight throughout the mesophyll and there are no air spaces or structure typical of normal palisade and spongy layers. Extra vascular elements, particularly phloem, often develop and link with the veins of the leaf. Tannins are common in the gall tissues and a layer of sclerenchyma (woody) tissue may strengthen the gall as it matures. In Fig. 56, the development and increasingly complex structure of three thrips galls is illustrated. The simple leaf fold of Arrhenothrips ramakrishnae becomes more complex if secondary galls develop on the main gall (Fig. 56a), especially if secondary galls develop on both sides of the leaf. The sclerenchyma, tannins and phenols of the rolls of Lygothrips jambuvasi (Fig. 56b) are unpalatable to caterpillars and other grazing insects and help to protect the gall and its thrips. The roll of Crotonothrips memecylonicus (Fig. 56c) forms a compact tube with a narrow longitudinal slit where it meets the leaf blade. Vascular strands develop in the rolls and link up with the midrib vein, and the inrolled halves of the leaf blade may fuse together, creating a closed gall with inner rolls partially dividing the thrips population. This can become quite complex, with vascular elements running across the join. In intervals between feeding, the females lay eggs within the developing gall, scattered over the surface or in discrete clusters or rows. The larvae hatch and feed, so causing further growth of the gall. Most thrips species have several generations in the one gall, all feeding continuously until it becomes full sized.

FIG 56. Sections through the developing leaf fold and roll galls of (a) Arrhenothrips ramakrishnae on Mimusops elengi;

(b) Lygothrips jambuvasi on Anogeissus latifolia;

(c) Crotonothrips memecylonicus on Memecylon lushingtonii (modified after Ananthakrishnan & Raman, 1989). Leaf midrib (open oval); nutritive tissue (stippled); sclerenchyma, tannins and phenols (black). In (c), X indicates that tissues may fuse across these gaps. In (a) and (c), secondary chambers have developed on the primary gall chamber.

CHAPTER 5

Pits, Blisters and Pouches

THE PITS, BLISTERS AND pouches in this, the second of the three chapters on open galls, form a series of increasing complexity. Like erinea, folds and rolls (Chapter 4), most pits, blisters and pouches are formed on new young leaves. They develop as invaginations of the leaf surface, some very slight forming a dimple or small basin (pit) or a swelling within the thickness of the leaf (blister), and others more obvious, like the finger of a glove or a bag-like distortion of the whole leaf (pouch). Many pouches are small, containing just one gall causer, though often making up for this by being very numerous. Others are remarkably large, several centimetres long and may contain thousands of individuals. Fig. 57 summarises the gall types included here (see also Fig. 38, Chapter 4, for examples of mite blisters and pouches). Most pits, blisters and pouches develop nutritive tissues or enhanced vascular strands, and the most complex pouch galls also include layers of woody cells and may become permanently closed. As in all insect and mite galls, continuous feeding by the gall causers is necessary for full development, by the larva or by the adult female and her offspring, and this results in establishment of a nutrient sink, diverting the plant’s resources from its own development to the gall (Chapter 1).

Pit galls in leaves or stems are caused by psylloids and scale insects (Hemiptera: Psylloidea and Coccoidea) and a few gall midges (Diptera: Cecidomyiidae). Background information for these groups is in Chapter 4 and Boxes 4.4, 4.6 and 4.8. Blisters are uncommon; they develop in leaves and are caused by gall mites (Chapter 4 and Box 4.1) and by gall midges. Pouch galls are commoner than either pits or blisters and make up the bulk of this chapter. Most pouches develop on leaves with a few in stems, and are caused by many organisms–gall mites, thrips, psylloids, scales, aphids and gall midges (for background information on thrips and aphids, see Chapter 4 and Boxes 4.3 and 4.5). In thrips, psylloids and aphids, pouch gall structure is fairly uniform although they vary greatly in size. In gall mites, scales and gall midges, pouch structure varies from very simple to a complexity rivalling that of gall wasp pea and marble galls (see Chapter 9).

FIG 57. Diagrammatic sections of pit (a), blister (b) and pouch (c–h) galls caused by plant bugs (Hemiptera), thrips (Thysanoptera) and gall midges (Cecidomyiidae) (modified after Dreger-Jauffret & Rohfritsch, in Shorthouse & Rohfritsch, 1992). The black shape represents the larva or nymph. See Fig. 38 (9 & 10) for mite blisters and Fig. 38 (11, 16, 17 & 25) for examples of mite pouches.

PIT GALLS

These small, basin-shaped depressions in leaves or stems are all simple galls, with little development of the specialised tissues characteristic of other galls, but they do show a gradation in complexity. The dimple caused by the gall midge Drisina glutinosa on sycamore Acer pseudoplatanus is one of the simplest. It is a shallow depression in the underside of the leaf, marked by a small pimple 1 mm across on the upper side, surrounded by a yellowish discoloured patch. A white or pale yellow larva lives in the depression bathed in sap exuded by the cells of the lower epidermis, sap which is probably derived from the vascular tissues. The dimple of Dasineura pustulans in leaves of meadowsweet (Filipendula ulmaria; Fig. 58) is slightly more complex, and includes a layer of nutritive tissue. Enlargement of the cells of the upper epidermis causes the upward doming of the leaf and occurs immediately the newly hatched larva starts to feed. Comparing sections of the leaf through the gall and the ungalled blade nearby reveals other changes. In the gall, the distinct palisade and spongy mesophyll layers disappear and the air spaces of the spongy layer become filled with large rounded cells. The lower epidermis, against which the larva lies, loses its waxy cuticle and forms into typical nutritive tissue (Chapter 1).

FIG 58. Dimple galls of Dasineura pustulans on meadowsweet Filipendula ulmaria. (Keith Harris)

Similar pit galls are caused by psylloids belonging to the genus Trioza, with a flattened nymph that fills the hollow on the underside of the leaf. In some species, the bulge in the leaf is caused by simple enlargement of cells of the upper side, as in the Dasineura pustulans gall. In others, cell multiplication is involved too, producing a thickened pit that tightly encloses the nymph, a close fit ensured by the short hairs that fringe its body. These simple basins represent growth of a gall in which development has stopped at an early stage. In southeast Asia and Australia where psylloid galls reach their greatest diversity, growth of the margins of the pit becomes progressively greater through a series of galls that culminates in pouch galls (see below).

The most familiar pit galls in Britain are caused by the scale insects Asterodiaspis (= Asterolecanium) in the bark of young twigs of oaks, Quercus robur and Q. petraea (Fig. 59). They are common in England and Wales as far north as Denbigh, Cheshire and Norfolk. Originally there was thought to be only one species, A. quercicola, later accepted as a synonym of A. variolosa. Microscopic examination of adult females and first instar nymphs, however, indicates that these two species are distinct and that there is a third, A. minus (Boratynski, 1961, provides keys to separate the females and young nymphs). Native to Europe, A. variolosa (and probably the other two species) has been introduced all over the world wherever the host oaks have been imported, and has spread to some related native oaks (e.g. the chestnut oak Quercus montana in New England, USA). On chestnut oak, the scale can be a major pest, even killing young trees 3 m or so in height. Such extreme damage is rare where the species are native; perhaps predators (e.g. tits, Parus) and parasitoids are abundant enough to keep their numbers down, or perhaps European trees are more resistant. The galls, habits and lifecycles of the three species are very similar. Males are unknown;* all species are parthenogenetic (thelytoky) and the galls often cluster together on the same twig.

FIG 59. Galls of Asterodiaspis variolosa on oak (from Redfern et al., 2002, with permission).

The galls are crater-like depressions in the bark, about 2 mm across, each containing a nymph or adult female (Fig. 60 shows development of the gall diagrammatically). The nymphs are flat, oval in outline and a glassy reddish brown; they become deeper and more circular in the adult stage, and yellowish green or brown with a waxy fringe around the margin. The gall is initiated in June by first instar nymphs (the crawlers) after they disperse from their mother’s old gall. They are active with long legs, and settle on new young twigs–the bark must be smooth and not yet deeply furrowed nor too thick and tough so that the stylets can penetrate it. Only the young crawler is capable of inserting its stylets; if it is removed after it has started to feed, it cannot insert it again. The twig changes as soon as the crawler starts to feed. The stylets penetrate through the bark to the collenchyma (a layer of supporting cells outside the vascular tissue of the stem) where they suck out the contents of each cell, which then dies. Surrounding collenchyma cells immediately start to divide and enlarge, increasing around the body of the nymph, and the gall, slightly soft and spongy at first, becomes apparent within 24 hours. The stylets of this scale do not reach the vascular tissue; it is the collenchyma layer that forms the bulk of the gall. Enzymes in the insect’s saliva convert starch and proteins in the cells to soluble sugars, peptides, etc., on which the scale feeds, and stimulate division and growth of the cells. The saliva of the crawler is certainly involved in gall formation but, as in other galls, the precise mechanism is not understood (see Chapter 1). When the scale is full-grown and stops feeding, the gall becomes dry, tannins are deposited in the cells and wound tissue develops around the gall, eventually producing a dead patch. If infestation is high and these dead patches encircle the twig, it will die and break off.

FIG 60. Development of the pit of Asterodiaspis variolosa (diagrammatic; modified after Parr, 1940). The mouthparts are long stylets that penetrate the bark and stem beneath and, as the scale grows, it sinks into a pit surrounded by stem tissue.

Asterodiaspis variolosa normally has one generation a year. The females are ovoviviparous, i.e. the nymphs hatch from eggs retained inside the mother’s body and are born alive. They remain for a few hours in their mother’s gall, under her protective scale, all that remains of her body after she has produced her brood. Then they become active, push out through a flap at her rear end and disperse to new twigs. Within a few days, they must find a spot to settle in, to insert their stylets and to start to feed. If the weather is cold, wet and windy during this period, many crawlers die. After they have settled, they grow and moult twice–into the second instar in about mid-July, a legless stage which lasts three or four weeks, and then into the adult female, also without legs. Crawlers born during August can induce galls but most fail to reach the adult stage before winter and so die. The adult female feeds through the autumn and her eggs develop. She overwinters in the gall, the eggs mature next spring and most young are born in early summer, after which she dies.

Gall-causing psylloids and scale insects are uncommon in Britain compared to the tropics, South America, southeastern Asia and Australia. In Taiwan, pit galls are common, with some species developing into more complex enclosed galls (Fig. 61). Like most Trioza, the species on Cinnamomum osmophloeum has an annual lifecycle. A simple pit appears in May initiated by the first instar as it feeds, tapping the phloem tissues beneath the epidermis. By early June the nymph is almost completely enclosed by growth of the margins of the pit, resulting in an oval shape ‘like a mung bean’. Hairs form within the opening and thicken as the gall develops, interlocking so that the insect is imprisoned (July). Meanwhile, the nymph grows; the second instar appears in June and, by December, it is in its fifth instar and fully grown. In January, the gall opens when the interlocking thickened hairs shrink and separate, and the adult winged insect escapes.

FIG 61. Summary of the lifecycle of Trioza sp. and development of its gall on Cinnamomum osmophloeum (Lauraceae) in Taiwan (after Tung et al., in Csóka et al., 1998). The insect is shown as a black shape. (For more explanation, see text.)

This example emphasises the difficulty of categorising galls. The Trioza gall starts as a simple open pit, similar to that of other psylloids and the coccoid Asterodiaspis. It becomes enclosed and bean-shaped (see other bean galls in Chapter 9) and opens to release the enclosed insect in a fashion similar to the pineapple galls caused by the spruce aphids (Adelges species, see Chapter 6). Some specialised scale insects (Coccoidea: Beesoniidae) in southeast Asia and Australia produce elaborate galls similar to those of Adelges and these are described in Chapter 6. See below for some bizarre pouch galls, too.

BLISTERS AND PUSTULES

These are simple galls formed within the thickness of the leaf. There is little difference between the two, apart from blisters being thinner-walled and more delicate than pustules. Eriophyoid mites and gall midges cause most of these galls, with the mite galls rougher and more scab-like. They are small, 1–3 mm across, sometimes surrounded by a discoloured zone and are raised on one or both sides of the leaf, with a hole on the upper or underside. The mite galls, especially, can be very numerous and sometimes coalesce, covering and distorting the whole leaf. Most blisters and pustules lack specialised nutritive tissue.

Mite blisters and pustules are caused by Aceria ulmi on small-leaved elms (Ulmus), and a complex of closely related species attacks pear Pyrus and whitebeam, rowan, etc. (Sorbus, Fig. 62)–Eriophyes pyri on pear and rowan and Eriophyes arianus on whitebeam. Pyrus and Sorbus are closely related genera in the family Rosaceae and an old name for the mite on whitebeam, which emphasises this relationship, is Phytoptus pirisorbi (later, the species was transferred to the genus Eriophyes). Although the mites in this species-complex look very similar, they cannot survive on each other’s host plants. Gall midges also cause blisters. Examples of the simplest are Monarthropalpus flavus on box (Buxus sempervirens, Fig. 63) and Physemocecis hartigi on lime Tilia, while Cystiphora sonchi on sow-thistles (e.g. Sonchus arvensis) and Physemocecis ulmi on elms Ulmus are slightly more complex. All gall midge blisters contain just one larva.

FIG 62. Galls of Eriophyes arianus (above left) on whitebeam Sorbus aria (Tom Higginbottom); and E. pyri (above right) on rowan S. aucuparia. (Michael Chinery)

These mite and midge galls are different in their structure. In mite pustules, the main cause of the gall is enlargement of the air spaces within the leaf, with minor hypertrophy and hyperplasy. The A. ulmi gall is less modified than that of E. arianus, with only slight enlargement of epidermal cells and with the palisade and spongy mesophyll layers of the leaf still recognisable. In the E. arianus gall, air spaces are enormous and the leaf structure is disrupted, with no distinction between the mesophyll layers. Cells of the upper epidermis are distinctly enlarged and apertures appear in this layer. Some cell multiplication occurs in the mesophyll and vascular tissues, and many of the stomata in the lower epidermis are malformed and non-functional. In gall midge blisters, cell enlargement is the main cause of the swelling, with hyperplasy important in more complex blisters that include nutritive tissue. In all blister and pustule galls, modification of leaf structure is greatest near the centre of the gall and grades into normal leaf tissue around its periphery.

In the mite galls, the fundatrix (who founds all the summer generations) initiates the galls by feeding on new leaves in spring, and small blisters are apparent in early May. She pierces individual epidermal cells causing several blisters that at first have no aperture and contain no mites. In the gall of E. pyri, the pear leaf blister mite (the same species occurs on rowan), a few epidermal cells near the centre of each blister enlarge and die, so producing a hole. Several mites insert their stylets into the hole but cannot enter as it is too small. As they feed, more cells die so enlarging the hole, and eventually a single fundatrix enters. She lays eggs inside and her offspring feed at the edge of the blister, enlarging it. When adult, the females leave through the aperture and initiate more galls in new leaves, with two or more generations occurring through the summer. Development is temperature-dependent, with more generations where summers are warm. The lifecycle of E. pyri is known in more detail than most because it is a pest of pear trees, especially in countries to which it has been accidentally imported (e.g. the USA and South Africa). Fertilised females from the final generation overwinter under bud scales or amongst the hairs of dormant buds and become next year’s fundatrices. E. pyri is an unusual species in that it has two forms which cause two types of gall, apparently dependent on temperature. In native areas of the mite in Europe, the blister gall form is common. In warmer climates, for example in South Africa and California, a bud gall form is more common. This form is active in winter and spring, producing several generations, and has a quiescent phase during summer. Mites infest buds in autumn and early winter causing them to brown and dry out; they may produce a few distorted leaves or flowers, or none. If fruit does form, the pears are small, misshapen, badly russetted and scabby, and drop early. These mites lay eggs within the buds and multiply during the winter. When healthy buds open in spring, adult females transfer to them, become quiescent during high summer, and migrate in autumn to the fruit buds where they start the cycle again. It seems that no blister galls develop in areas with hot dry summers and mild winters.

FIG 63. Blister galls of Monarthropalpus flavus on box Buxus sempervirens. (Keith Harris)

The gall midge Monarthropalpus flavus causes a simple blister gall on box (Buxus sempervirens; Fig. 63). The female has an unusual hooked ovipositor. In spring, she uses this to pierce the lower epidermis of a new leaf and moves it to and fro through a semicircle, creating a cavity in the spongy mesophyll, destroying cells and slicing through the vascular tissue of a few veins. The path of the ovipositor later becomes the aperture, always off-centre on the underside of the blister. She lays an egg in the cavity. The larva hatches and probably feeds on cell sap exuding from the severed xylem and phloem. The plant responds to this wounding by producing large callus cells around the larval chamber, modified mesophyll cells that cause the swelling in the leaf, and the larva probably feeds on these, too. There is no evidence of hyperplasy (multiplication) of cells. The gall of Physemocecis hartigi on lime is always associated with a leaf vein. This gall is a little more complex in that the cells around most of the larval chamber become lignified, forming a thin woody zone. This may help to protect the larva from attack by parasitoids. More elaborate blisters are those of Cystiphora sonchi (Fig. 64) and Physemocecis ulmi (Fig. 65) that contain typical nutritive tissue. The larval chamber develops above the lower epidermis and two or three layers of spongy mesophyll cells become transformed into nutritive cells.

FIG 64. Blister galls of Cystiphora sonchi on sowthistle Sonchus arvensis. (Michael Chinery)

FIG 65. Blister galls of Physemocecis ulmi on wych elm Ulmus glabra. (Keith Harris)

POUCHES

Pouch galls form by invagination of the leaf to form a projection like the finger of a glove or a bag-like distortion of the whole leaf. Usually it is the underside of the leaf that lines the inside of the gall and, throughout its development, it normally retains an opening to the outside (though not in all aphid and thrips pouches). A few pouches develop on stems but it is on leaves that they are most diverse. Although they are caused by a wide variety of organisms, the pouches of different groups often look superficially similar–gall midge and gall mite pouches or those of psylloids and aphids are sometimes similar. Most pouches develop rapidly and include a layer of nutritive tissue, and vascular strands develop in the gall walls and link with the veins of the leaf. Not in aphid pouches, though; in these, vascular elements multiply and usually no nutritive tissue develops because the long aphid stylets tap the vascular tissue directly. In thrips, psylloids and aphids, the structure of the pouches is fairly uniform though they vary greatly in size, while gall mite, scale and gall midge pouches vary in structure. Examples are described for each group.

Gall mite pouches

All eriophyoid pouch galls are initiated in the spring by the fundatrices, fertilised females that have overwintered in cracks and crevices in the bark or under the scales of dormant buds. A fundatrix wanders over a new leaf usually on its underside, and feeds from individual cells, in a manner similar to erineum and fold and roll mites (Chapter 4), and these cells collapse and die. The leaf around each dead cell domes upwards and the pouches develop rapidly, but the females do not enter them yet. Nutritive tissue forms on the inside of the pouch from the lower epidermis and one or more layers of mesophyll cells, and any difference between the palisade and spongy layers disappears. Pouches can develop the other way up but are less common. When the pouch is partly grown, the female returns and lays eggs inside. The larvae hatch and feed, sucking cells dry; these cells die and stimulate the development of more nutritive tissue, which becomes more abundant the larger the number of mites in the pouch. The mites multiply, often through several generations over the summer, and an increasing number of nutritive cells collapses, causing scar patches on the inner gall surface. If the mites abandon the gall, it soon reverts to the normal structure of leaf with palisade and spongy mesophyll layers and stomata in the lower epidermis. Continuous feeding by the mites prevents this. In late summer or autumn, the new fertilised females leave the pouches to find overwintering sites and the galls decay with the leaves.

Hairs are characteristic of mite pouch galls and develop from epidermal cells usually around the aperture. These hairs are slender and elongate, with single nuclei, thick walls and large vacuoles, and may become lignified; they are quite unlike nutritive cells or erineum hairs. They block the aperture and probably are protective, helping to prevent water loss and the entry of enemies. They do not develop where the mites are feeding. In young galls in which no eggs have been laid or which have been abandoned, these hairs develop all over the inner surface and the pouch aborts without reaching its full size. On a well-colonised leaf, there may be galls of a range of sizes, normal healthy ones and smaller ones that have never received eggs or in which the mites have died.

FIG 66. Pouch galls of Aceria macrorhyncha on a leaf of sycamore Acer pseudoplatanus. (Michael Chinery)

FIG 67. Nail galls of Eriophyes tiliae on large-leaved lime Tilia platyphyllos. (Michael Chinery)

FIG 68. Pouch galls of Aceria campestricola on English elm Ulmus procera. (Michael Chinery)

FIG 69. Sections through the pouch galls of (a) Aceria macrorhyncha on sycamore Acer pseudoplatanus, mature gall on left and abandoned gall on right; (b) Aceria campestricola on English elm Ulmus procera, full-grown gall (modified after Meyer & Maresquelle, 1983). Nutritive tissue (stippled); gall mites (black spots); woody shell (black layer); leaf vein (open circle).

Figs 66–8 illustrate three common species of mite pouch galls. They demonstrate an increase in complexity–summarised in Fig. 69, which shows the structure of the least and most complex. Each fundatrix of Aceria macrorhyncha in late April initiates about 20 pouches on a sycamore leaf, when the palisade and spongy mesophyll layers are already distinct, although this structure is soon lost inside the pouch. The nutritive tissue of this pouch forms a layer two or three cells thick but the cells are not greatly modified. Mature pouches grow up to 6 mm high and have soft, fleshy walls. Eriophyes tiliae causes slightly more complex pouches on large-leaved and common lime. Each fundatrix causes just one gall, near a vein, in leaves that have not yet developed distinct palisade and spongy mesophyll. The cells of the nutritive tissue are larger than in the A. macrorhyncha pouch and contain several large polyploid nuclei–there is nearly three times as much nuclear material in the nutritive layer as in the normal leaf tissue, with these cells containing two to four times as many sets of chromosomes. Lime nail galls are considerably tougher than sycamore pouches, with thicker walls, more supporting tissue (collenchyma) and with vascular strands enclosed in lignin sheaths. The mites multiply in the pouches throughout the summer reaching a peak, in E. tiliae, of 100–200 per gall and, as they feed, nutritive cells collapse forming patches of scar tissue that increase in size into the autumn. By September and October when the mites disperse to overwintering sites, the nutritive layer has disappeared. As in the sycamore pouch, hairs protect the aperture of the lime nail galls and spread throughout the pouch if the gall is abandoned.

FIG 70. Galls of Eriophyes dichrostachia on Dichrostachys glomerata with sections through pairs of galled leaflets (after Meyer & Maresquelle, 1983).

One of the most complex of mite pouch galls is caused by Aceria campestricola on small-leaved elms (Fig. 68). This is a small, hard pouch with thick walls, rounded above with a conical projection below, and is often very numerous on a leaf. Like E. tiliae, each fundatrix causes just one gall and attacks a very young leaf before the palisade and spongy mesophyll layers develop. This gall is an unusual mite pouch gall because, about three weeks after initiation, a woody layer appears between the nutritive layer and the outer gall, forming a hard shell two to four cells thick (Fig. 69b). These cells have thick lignified walls, punctured with wide gaps, and contain living cytoplasm and nuclei and can interchange materials with neighbouring cells. As in the other mite pouches, the characteristic layers develop only if mites are feeding actively and multiplying. In galls with few mites, nutritive tissue develops where they feed but the rest of the inner epidermis may develop hairs and the woody shell becomes fragmented, and tissue layers characteristic of normal leaves may appear. This occurs in abandoned galls, too.

A final example illustrates a bizarre pouch gall formed by the joining of outgrowths from two adjacent leaflets that become superimposed, caused by Eriophyes dichrostachia on the African tree Dichrostachys glomerata (Fig. 70). These cup-like growths fuse together to form an enclosed cavity, and several galls may develop in sequence, soldering together a row of leaflets. Very little is known about development of this gall or the lifecycle of its mite; presumably the gall opens at some stage to allow the mites to escape to infest other leaflets. Pouches formed from two adjacent leaflets are also known in gall midges (see below).

Thrips pouches

Some thrips pouch galls are simple and non-woody, similar to simple eriophyoid pouches, and a female (the fundatrix) can induce many on a leaf. A female, e.g. of Liothrips, punctures a cell of the lower epidermis of a young, undifferentiated leaf and, within a day, a transparent spot appears on the upper surface above the puncture. The spot turns red and grows into a tubular gall 6–8 mm high in less than a week. Females of other species may also cause galls on the same leaf, and it can become covered with them and twisted and distorted as a result. The females enter the growing horns and lay eggs over several days; the eggs hatch and the larvae grow rapidly, and at any one time a gall may contain eggs, larvae, pupae and adults. In some species, the horns reach a length of 30 mm, and they may be simple tubes or become branched and convoluted. The wall of the gall is fleshy and lined with patches of nutritive cells interspersed with dead, shrivelled cells that the thrips have sucked dry. The wall increases in thickness as the horn develops and contains phloem elements linking with the veins of the leaf, and supplying the feeding adults and larvae with nutrients via the nutritive cells.

One of the largest thrips galls is a pouch 8–10 cm across, caused by Austrothrips cochinchinensis, which develops in an axillary bud of Calycopteris floribundus (Combretaceae) in India. When mature it is an irregular, hollow bag with a wrinkled, convoluted surface and a hairy opening at the top, and contains up to 5,000 thrips. It starts to develop when several females infest an expanding bud 5–10 mm long. They feed on the meristem, soon killing it, and then turn to the inner surfaces of the young leaves. These thicken due to multiplication of cells and opposite pairs of leaves fuse together, forming a chamber that enlarges as the leaves in the bud grow (Fig. 71). A large population of thrips inside causes the walls to infold into the chamber, separating the thrips into partially isolated sub-chambers.

The open nature of most thrips galls allows easy dispersal. Newly mated females leave when they are a few days old to start new galls elsewhere and some males probably migrate to other galls of the same species in order to find virgin females. Larvae are mobile enough to leave the galls but rarely do so; they are more susceptible to desiccation than the adults and most do not survive outside. Some thrips produce soldier morphs with enlarged and armed forelegs (see Box 4.3), which defend the gall against intruders, such as other thrips attempting to take over the gall or predators and parasitoids looking for prey. The fundatrix has weapons too and she fights other females for the best gall sites (similar behaviour occurs in gall aphids, see below).

FIG 71. Sections through a normal bud and the developing pouch gall of Austrothrips cochinchinensis on Calycopteris floribundus. Vertical sections: (a) normal bud, (b–d) galled bud; horizontal sections: (e) normal bud, (f–h) galled bud (after Ananthakrishnan & Raman, 1989). Meristem (black); nutritive tissue (stippled); leaf midribs (open circles); arrow in (f) indicates fused leaves.

Plant bug pouches

A few psylloids (Psylloidea) and scale insects (Coccoidea) cause pouch galls, although not in Britain, but their lifecycles and gall structure are not well known. Pachypsylla species (Psylloidea: Psyllidae) gall the North American hackberry (Celtis species, Fig. 72) and pouches of scales are most diverse on Australian Eucalyptus species (Fig. 73). Like aphids, these insects suck fluids through piercing stylets. Their stylets often are quite short; they pierce cells of nutritive tissue that develops beneath the insect but they cannot reach the vascular tissue of the leaf. In the Phylloxeridae (Aphidoidea, Box 4.5), the stylets may be long or short and their galls may develop nutritive cells. True aphids (the Aphididae), on the other hand, have long stylets that can reach the phloem. The vascular strands multiply but most of their pouch galls do not develop typical nutritive cells. Tetraneura ulmi on elms (Ulmus; Fig. 74) seems to be an exception, with a thick layer of nutritive tissue lining the chamber; it is also well supplied with vascular strands. Most species in the aphid subfamily Eriosomatinae cause pouch galls and specialise on poplars, elms and pistacia trees.

FIG 72. Galls of Pachypsylla celtidismamma on hackberry Celtis occidentalis (after Meyer, 1987). (a) Underside of a leaf with five galls. (b) Enlargement of one pouch gall. (c, d & e) Sections through the developing gall showing the position of the psyllid (black) with its stylets piercing the nutritive tissues beneath it; a layer of sclerenchyma develops below the nutritive tissue (neither of these tissues is shown).

FIG 73. Galls of scales (Coccoidea) on Eucalyptus species in Australia (after Gullan et al., in Raman et al., 2005). (a) Apiomorpha munita: a group of tubular male galls on a globular female gall. (b) Maskellia globosa: twig with knobbly woody female galls and tubular male galls on leaves; a female gall is shown in section and two male leaf galls are enlarged.

FIG 74. Mature gall of Tetraneura ulmi on wych elm Ulmus glabra with two openings enabling the aphids to escape. (Michael Chinery)

The grape phylloxera, Daktulosphaira vitifoliae (Phylloxeridae; previously known as Viteus vitifoliae or Phylloxera vastatrix) is notorious due to the damage it caused to European vines and the wine industry in the second half of the nineteenth century (see Chapter 13). It spends all of its life on the one host plant and is unusual in causing two types of galls, one on the leaves and the other on the roots. Its lifecycle is complex and typical of aphids (Box 4.5). In spring, fundatrices hatch from overwintered eggs laid in crevices in the bark of two-year-old stems. Each fundatrix settles on the upper side of a young leaf near a vein, pierces nearby cells with her short stylets and starts to feed. The effect on the leaf is rapid. Cells around her multiply and enlarge, enclosing her in a pouch with a hairy aperture in the upper surface (Fig. 75), and producing a thickened cone on the underside. If she settles on the underside of the leaf, the gall may start to form but fails to enclose the female; she stops feeding and dies, and a small, distorted gall results. Fundatrix galls can also develop in leaf petioles, stems and tendrils. Vascular tissues develop in the wall of a stem gall, supplying cells of the gall with nutrients, but the insect does not feed directly from the phloem.

After two or three weeks and three moults, the fundatrix becomes adult and, without mating, lays 300 or more yellowish eggs in the gall (phylloxerids are not viviparous unlike true aphids). These hatch into crawlers, all females, and either crawl out of the aperture to disperse and form their own galls on nearby new leaves, or remain in the maternal gall, which can become very crowded. When adult they lay more eggs, producing more females and inducing more leaf galls, through three or four generations in spring and summer. Although the last generation on the leaves only produces about 100 eggs per female, the potential for increase is phenomenal: if all survived, one fundatrix could produce 5,000 million females on the vine shoots, all genetically identical to her. Although only a small proportion survives, the rate of spread of the insect in a vineyard can be rapid. As the season progresses, an increasing proportion of crawlers migrates down the vines to the roots. They settle on young rootlets and cause simple swellings, caused by enlargement of cells and by limited multiplication of the phloem, swellings that normally do not enclose the insects. By September, the galled leaves are senescing and most individuals are underground. More generations occur on the roots particularly if the autumn is mild, all still female and genetically identical to the fundatrix. Eventually, eggs are laid on the roots that hatch into sexuparae and develop into winged females. These climb out of the soil and disperse to other vine shoots, and lay one egg each, either a larger female egg or a smaller male egg. Females and males (the sexuales) develop from these and mate, and each female lays one winter egg in a bark crevice, so completing the lifecycle. The root aphids are much less fecund than the leaf forms but, even so, reproduction in one year could, theoretically, produce 5 tons of insects from a single vineyard! Fortunately, most of them do not survive.

FIG 75. Galls of the grape phylloxera Daktulosphaira vitifoliae on vine Vitis vinifera: upperside of a heavily infested leaf. (Michael Chinery)

FIG 76. Galls of Pemphigus species on black poplar Populus nigra and its hybrids in Britain. (clockwise from top right): Pemphigus populinigrae (= P. filaginis), P. bursarius and P. spyrothecae. (a & b, Michael Chinery; c, Tom Higginbottom)

Daktulosphaira vitifoliae is a North American insect, native to an area east of the Rockies and southwards perhaps as far as Venezuela. It was introduced into Europe in the 1860s where it caused enormous damage to European vines (see Chapter 13 for the story of the great wine blight; American Vitis species are more resistant). The full lifecycle is common in the USA but, in southern Europe where winters are mild, it is often shortened to the root generations alone, with nymphs or adults overwintering, and so omits the sexual and leaf gall stages. It is the root forms that cause most damage to the plants, eventually destroying susceptible varieties of Vitis.

Most of the galls caused by true aphids (Aphidoidea: Aphididae) belong to the subfamily Eriosomatinae, which is subdivided into the tribes Eriosomatini, Pemphigini and Fordini (see Box 4.5). They cause pouch galls of a variety of shapes and sizes, from under 1 cm to more than 20 cm in length, and specialise on Ulmus (Eriosomatini), Populus (Pemphigini) and Pistacia (Fordini). Their lifecycle follows the basic aphid pattern, differing from the grape phylloxera in that most reproduction is viviparous, and usually with alternation between a primary host (usually a tree, specific to each aphid species) and the roots of secondary hosts (usually herbaceous species, and most aphids can feed on a variety of these). Galls develop in young, rapidly growing leaves or leaf buds only on the primary host. Reproduction is nearly all parthenogenetic (thelytoky, see Box 4.5) and occurs at a phenomenal rate. A brief sexual stage, involving tiny sexuales that do not feed, occurs on the primary host and produces fertilised eggs (only one per female) and these hatch into the fundatrices. Selected examples are described that illustrate the variety of galls in the subfamily (Figs 76 and 77).

Gall formation is probably similar in all species. It has been described in detail in an American species, Pemphigus betae on narrowleaf cottonwood (Populus angustifolia). The fundatrix nymph settles alongside the midrib of an immature leaf, inserts her stylets and induces a depression about 1 mm deep. Over the next two days, she continually probes back and forth along the midrib over a distance of about 5 mm, and cells either side of this track multiply and enlarge so forming a pouch that closes over her, sealing her inside. The linear pouch gall on the underside is recognisable within four days. The pattern of probing helps to determine the shape of the gall and the site of the future aperture–in Pemphigus spyrothecae on Populus nigra, the probing pattern produces the spiral twisting of the petiole (Fig. 76). Inside, phloem elements multiply in the gall wall. The fundatrix plugs into a phloem sieve tube, grows into an adult and produces large numbers of young, and these feed in her gall so enlarging it. The lifecycles of two pemphigine species are shown in Fig. 78. Pemphigus bursarius (its gall is shown in Fig. 76) usually has a typical holocycle with two host plants, while P. spyrothecae is a ‘monoecious holocyclic’ species, i.e. all stages occur on the primary host and there is no summer migrant stage. But P. bursarius is sometimes anholocyclic; in parts of its geographical range, it lives permanently on the roots of its secondary hosts and the sexual and gall-causing stages are eliminated from its lifecycle. In both Pemphigus species, the gall apertures open before the winged stages are ready to leave, and this allows for ‘housekeeping’. Like most aphids, pemphigines produce honeydew that could contaminate and even drown the aphids in a closed gall. They avoid this by producing filaments of wax, which line the gall cavity and coat the droplets of honeydew. These form ‘marbles’ that prevent the honeydew from wetting the aphids and the gall cavity. The aphids can easily roll the marbles around and they push them out of the aperture of the gall when it opens. Wax glands are essential to the survival of aphids living in galls, and all gall aphids are waxy. Many free-living species are waxy too, although the most recently evolved species have lost their wax plates. Some free-living species are attended by ants, which ‘milk’ them for their honeydew so solving the problem, but this solution is not available to the galling species. In P. bursarius, the gall opens in June although some aphids may not leave until September. The P. spyrothecae gall opens later, the coils loosening in August, with the aphids departing over the next two or three months.

FIG 77. Galls of Pistacia aphids (Pemphiginae: Fordini); all are formed in leaflets except for (e) which is initiated in an axillary bud. (a) Aploneura lentisci on Pistacia lentiscus (Michael Chinery); (b) Forda marginata on P. palaestina (Michael Chinery); (c) Forda formicaria on P. palaestina (Michael Chinery); (d) Geoica wertheimae (= G. utricularia) on P. palaestina (after Meyer, 1987); (e) Slavum wertheimae on P. atlantica (after Aloni et al., 1989); (f) Baizongia pistaciae on P. palaestina, a giant gall formed from a single leaflet (Michael Chinery).

Pistacia trees, native around the Mediterranean from Spain and Morocco to Iran, are the primary hosts of the Fordini, and these cause a variety of pouch galls (Fig. 77). In most species, the fundatrix settles on the upperside of the leaf but, because the invagination is deep, the gall appears on the underside. Most of the pouches develop in newly expanding leaflets and some are large and bizarrely shaped, turning red in the sun and containing thousands of aphids. The Israeli fauna has been particularly well studied; there are about 16 species, and all are specific to particular Pistacia species. The pouches are the result of enlargement and multiplication of cells, as in the Pemphigus galls, and contain numerous vascular strands in the gall wall linking to the veins of the leaf and stem.

FIG. 78. Lifecycles of gall-causing pemphigids. (a) Pemphigus bursarius; (b) Pemphigus spyrothecae (modified after Foster & Northcott, in Williams, 1994). The central circles depict months of the year. The whole lifecycle is diploid except for production of haploid gametes, the sperm and ova. Red arrows indicate ‘gives birth to’, green arrows indicate ‘grows up into’. TP, thelytokous parthenogenesis; AP, arrhenotokous parthenogenesis; n, haploid; 2 n, diploid.

Like Pemphigus, the lifecycles of most Fordini are holocyclic, requiring two unrelated host plants, and the secondary hosts are usually grasses (Poaceae). Unlike Pemphigus, the lifecycle generally takes two years to complete (Fig. 79, compare Fig. 78a). The gall on Pistacia is initiated in spring and grows quickly but does not open to release the first winged forms until autumn. These migrants fly to the secondary hosts and feed on the roots without causing galls; they multiply through several generations as long as the weather is mild and produce winged sexuparae. These fly back to the primary host in the following spring and produce the sexual males and females. Mating occurs and the females produce one fertilised egg each. These pass through a second winter, protected by the shrivelled body of the female, in a crevice in the bark, before hatching in the spring (the third spring) into the fundatrix. So, compared to the poplar aphids, development is slow, but multiplication can be phenomenal, e.g. the large gall of Baizongia pistaciae (Fig. 77f) may contain 10,000 or more individuals.

FIG 79. Lifecycle of Baizongia pistaciae (modified after Blackman & Eastop, 1994).

Some Pistacia aphids, like Pemphigus spyrothecae on poplar (Fig. 78b), have lost the secondary host so that all stages occur on one Pistacia species. Slavum wertheimae (Fig. 77e) is one such species and is also unusual in having a one-year lifecycle. Some Fordini have a geographical range wider than the distribution of Pistacia trees and, outside the range of Pistacia, they occur only on the secondary hosts, with no gall or sexual stage (as in some forms of P. bursarius). Forda formicaria and F. marginata, for example, are found in northern Europe, North America and New Zealand where Pistacia does not grow, and are anholocyclic on roots of grasses.

The fundatrices of several eriosomatid species are competitive and territorial. In spring, they must search for suitable young leaves to gall. The timing between the hatching of the overwintered eggs and the arrival of fundatrices on suitable leaves is tight–the ‘window’ of synchrony may be only about three days, so restricting the number of leaves available to be galled. If two or more fundatrices of Pemphigus betae arrive on the same leaf at the same time, they will compete for the best place at the base of the leaf. Not all leaves are equally attractive, too. The ‘best’ leaves are those that grow rapidly, are destined to become the largest, that attract the greatest share of nutrients (i.e. become the strongest nutrient sink), and that contain the lowest concentration of chemical defences (but these desirable leaves may be only 1.6 per cent of the total available). Small leaves with galls often drop early, before the aphids have left, and this can mean death of the whole clone. The fundatrix in the most favourable position will actively defend it against all comers, kicking and shoving with her hind legs, and contests can last two days and may be to the death. Even after starting a gall, the fundatrix can still be displaced while the gall is open. Although the largest nymph usually wins, there is a cost. While she is defending she is not feeding, so the gall grows less rapidly, becomes a less strong nutrient sink and, when adult, she will produce fewer offspring. In P. betae, even with the field to herself, a fundatrix produces six times as many offspring on the largest leaves compared to the smallest, and if small leaves are the only ones available, 80 per cent of females may die attempting to initiate a gall. Some species defend the gall later in development, too. In P. spyrothecae (Fig. 78b), the offspring of the fundatrix become soldiers in their first instar, their weapons being their stylets and hind legs. They patrol the aperture of the gall when it opens, actively defending it and driving away intruders, including predators. They also carry out housekeeping jobs, pushing out wax-coated honeydew droplets and any cast skins or dead aphids. They lose this behaviour when they moult into the second instar. Some of their offspring also become soldiers in the first instar and these may be unable to grow up into winged sexuparae.

A few species cause two types of gall on the primary host. In Forda formicaria on Pistacia palaestina, the fundatrix nymph induces a small pouch gall less than 5 mm long at the tip of the mid vein of a leaflet. Inside, she matures and produces her first offspring. In their first instar, they disperse to other leaflets and each causes a crescent-shaped final gall in the leaflet margin (Fig. 77c), which enlarges as numbers increase through the summer. The same thing happens in Thecabius affinis in Britain; the fundatrix gall is a small fold on the edge of a leaf of black poplar and her offspring’s galls are large pouches either side of the midrib (Fig. 80).

A problem arises for the aphids in galls that become completely sealed, e.g. Baizongia, Geoica and Slavum–they need to escape and do not have suitable tools with which to bite or bore their way out. The galls of Geoica wertheimae (Fig. 77d) and B. pistaciae (Fig. 77f) crack open from September to November, allowing the autumn migrants to escape. In the ‘cauliflower’ gall of Slavum wertheimae (Fig. 77e), winged aphids develop in the tips of the branches, and these break open at the end of the summer. In Tetraneura ulmi (see Fig. 74), the pouch develops as an invagination of the leaf with a hairy aperture on the underside, which seals over completely before the gall is full-grown. In June or July, a split appears in the gall wall above the stalk. It starts in the outer epidermis and the mesophyll cells enlarge, rupturing the inner epidermis, and forming an aperture with thick lips that flare outwards, allowing the aphids to escape.

FIG 80. Fundatrix (top) and final galls of Thecabius affinis on leaves of Populus nigra (from Redfern et al., 2002, with permission).

Gall midge pouches

The following descriptions illustrate a range of gall midge pouches (Fig. 81) with simpler to more complex structure. The ‘lighthouse gall’ of Rondaniola bursaria on ground-ivy Glechoma hederacea, a hairy pouch up to 4 mm tall, is simpler than most (Figs 81a and 82), with fleshy walls not much thicker than the normal leaf and a thickened ring at the base. Hairs develop from epidermal cells lining the aperture; they probably help to protect the interior from extremes of climate and from predators. In August or September, the larva is full-grown and the gall detaches itself from the leaf, leaving a neat circular hole. The split occurs above the thickened base and is due to maceration of the middle lamella, the layer between cells that glues them together. The split develops only if the larva inside is full-grown and active; if it dies, the gall remains attached to the leaf. The larva overwinters in the detached gall, pupates in the spring and the adult emerges to infest newly expanded leaves the following year.

The pouch gall of Dasineura urticae is thick and irregular in shape, 3–8 mm across, and develops on leaves, petioles, flower stalks or stems of stinging nettle Urtica dioica, sometimes several merging together (Fig. 81b). Adults emerge from pupae in the soil; they mate and the females lay eggs on the upper surface (usually) of young expanding leaves or in grooves of the stem nearby. The eggs hatch two to four days later and, on the leaf, one or more hatchlings moves into the trough above the midrib or a side vein usually near the base of the leaf. Adjacent cells multiply causing lips to grow around the larva, enclosing several in one pouch if they are feeding close together, and a slit-like, hairy aperture develops usually on the upper side (Fig. 83a). In the full-grown gall, nutritive tissue lines the cavity, a thick layer of cells rich in starch lies outside this and vascular elements develop further out, linking to the veins of the leaf. The cortex, which forms the bulk of the gall, consists of large vacuolated parenchyma cells. This pouch gall is more complex than that of R. bursaria in that it has a layer of starch storage tissue, able to supply carbohydrates to the nutritive layer and thence to the larva. The structure of the gall is much the same whether it forms in a leaf, a stem or in flowers. In the flowers and depending exactly where the gall forms, the floral parts may be totally inhibited. If the embryo is not destroyed, though, and if pollination can occur, some seed may be produced.

FIG 81. Full-grown pouch galls of Cecidomyiidae: (a) Rondaniola bursaria on ground-ivy Glechoma hederacea; (b) Dasineura urticae on stinging nettle Urtica dioica; (c) Dasineura ulmaria on meadowsweet Filipendula ulmaria; (d) Geocrypta galii on lady’s bedstraw Galium verum; (e) Mikiola fagi female galls, the male galls are slimmer, on beech Fagus sylvatica; and (f) Hartigiola annulipes hairy galls, smooth galls are also common, both on beech; (g) Didymomyia tiliaceae on lime Tilia sp. (Michael Chinery)

D. urticae has two generations a year. Adults emerge from the soil at the end of May or early June and their offspring cause the first generation of galls. Four or five weeks later, the larvae are full-grown and drop out of the galls. They pupate in the soil (there is no cocoon) and emerge as adults in July to August. Nettles produce new leaves continuously throughout the summer. The second generation of galls develops in these, or in the stems and flowers, and the larvae are fully fed in late August or September, when they leave the galls and enter the soil to overwinter. They pupate in early May, and adults emerge to start the next generation in late May or June.

FIG 82. Sections through the pouch gall of Rondaniola bursaria: (a) 5–8 days old; (b) the mature gall (modified after Meyer & Maresquelle, 1983). Nutritive tissue (stippled); larva (black shape); leaf vein (open circle).

FIG 83. Sections through developing and mature galls of Dasineura urticae on a leaf of stinging nettle Urtica dioica: on the midrib (a–d) and on the leaf blade (e–g); the tissue layers are shown only in the full-grown galls, (d) and (g) (after Meyer & Maresquelle, 1983). Larva (black); nutritive tissue (stippled); starchy layer (hatched); midrib (open oval).

The galls of Dasineura ulmaria, Geocrypta galii, Mikiola fagi, Hartigiola annulipes and Didymomyia tiliaceae are increasingly complex (see Fig. 81 for their galls). They all start to develop in the same way. The newly hatched larva settles on a young leaf or stem, starts to feed and nutritive tissue develops beneath it and, around it, cells divide and grow to envelop it. In D. ulmaria, M. fagi and H. annulipes, the eggs are laid on the underside of the leaf and the pouch has an opening on the underside. In D. tiliaceae, the larva starts feeding on the upper side and the gall develops the other way up, with its aperture at the top. And in G. galii, the larva settles in a groove of the stem and the gall develops around it. Fig. 84 illustrates the final structure of these pouches. In D. ulmaria, the lips extend on the underside of the leaf to form a pale yellow, hairy, conical projection, with a small patch of nutritive tissue at the head of the chamber. In G. galii, growth of the lips causes a rounded gall and continual movement of the young larva inside results in nutritive tissue lining the whole chamber. In H. annulipes, early development of nutritive tissue at the tip of the pouch (equivalent to the base of the pouch in D. ulmaria) is soon modified into sclerenchyma, and new nutritive tissue develops in the sides of the chamber, induced by the larva circling horizontally through 360 degrees while it feeds. As this larva feeds, it hammers the cells with its head without rupturing them. In the complex gall of D. tiliaceae, a new cambium layer develops outside the woody layer (Fig. 84e) and this layer causes a split to develop around the woody inner gall so that it eventually falls to the ground (see below). In the H. annulipes and D. tiliaceae galls, the nutritive cells induced by the younger larvae are consumed by the third instar larva, which grows very rapidly. Although nutrients continue to be supplied from the vascular strands and from stored starch, the larva sucks out the contents of successive layers of nutritive cells so that nutrients have to cross several layers of dead and empty cells. In the gall of D. tiliaceae, too, nutrients from the vascular strands need to diffuse to the larva across the barrier of the sclerenchyma layer (Fig. 84e; this also occurs in the D.

FIG 84. Sections through full-grown cecidomyiid pouch galls. (a) Fleshy gall of Dasineura ulmaria on meadowsweet Filipendula ulmaria. (b) Stem gall of Geocrypta galii on hedge bedstraw Galium mollugo. (c) Woody gall of Mikiola fagi on beech Fagus sylvatica. (d) Mature five-month-old hairy gall of Hartigiola annulipes also on beech (galls of this species are often hairless). (e) Mature woody four-week-old gall of Didymomyia tiliaceae on lime Tilia sp. before abscission of the inner gall (modified after Meyer & Maresquelle, 1983; Rohfritsch, in Shorthouse & Rohfritsch, 1992). Larva (black); nutritive tissue (stippled); starch layer (hatched); sclerenchyma or woody layer (cross-hatched); leaf vein (open circle); vascular bundles (dashed lines).

FIG 85. Development of the pouch gall of Hartigiola annulipes over four months (modified after Rohfritsch, in Shorthouse & Rohfritsch, 1992). A section of the mature five-month-old gall is shown in Fig. 84d. For more explanation, see text. Larva (black); nutritive tissue (stippled); woody tissue (cross-hatched); vascular tissue (open circles and dashed lines).

ulmaria gall, Fig. 84a). As in some pouch galls of mites (see above), these woody cells are alive and active and retain connections with their neighbours via perforations in their thick lignin cell walls. Thus, nutrients can diffuse across this layer unimpeded.

Most gall midge pouch galls develop rapidly in only a few weeks. The gall of H. annulipes is unusual in that it requires five months to develop to maturity, from April to September, and it is worth describing its development in detail. The long development has led to muddles over the species’ name; until Rohfritsch described its development (in 1971), the immature galls were thought to belong to a different species. Early development is similar to other cecidomyiid pouches (Fig. 85). Twenty-four hours after the hatchling has started to feed, the leaf has domed upwards with a small patch of cells modified into nutritive tissue. After two or three days, these initial nutritive cells have lignified, and lips begin to grow downwards around the larva, and new nutritive tissue forms around the sides of the chamber. By one week old, the lips have almost encircled the larva and are becoming woody. Then, development slows down. After two months, the upper epidermis becomes detached from the palisade mesophyll and the leaf thickens around the central chamber, and a space forms between the upper epidermis and the palisade layer. At four months, the pouch is extending upwards and ruptures the upper epidermis; hairs develop from the palisade layer, which is now modified into a new epidermis. Then the gall grows rapidly, and reaches its full height of about 6 mm (Fig. 84d) within a further month, and the enclosed larva becomes full-grown.

Like the Rondaniola bursaria gall described above, the galls of Mikiola fagi, Hartigiola annulipes and Didymomyia tiliaceae are dehiscent, falling to the ground before the leaves are shed in the autumn. The galls split from the leaf along a predetermined line of weakness, a layer of thin-walled parenchyma cells. In Mikiola fagi, this line occurs between two woody layers at the base of the gall (see Fig. 84c), and these separate when the middle lamella (the layer that glues cells together) dissolves. In D. tiliaceae, dehiscence is a little more complex. The layer of parenchyma cells occurs outside the woody inner gall (Fig. 84e) and breaks down so loosening the inner gall. The outer cells of this parenchyma zone then increase in size and number, exerting a pressure against the inner gall and so pushing it out. A hole is left behind in the leaf, which soon gets covered over.

Apart from Dasineura urticae, these gall midges have one generation a year in Britain and their lifecycles are similar. Mikiola fagi is typical (although its gall is rare now in Britain and it may even have died out here). Adults emerge from last year’s galls in April or early May, mate and lay red eggs singly or in batches of 20 or more on the scales of beech buds. The larvae hatch, push between the opening leaf scales to the underside of the emerging leaf, and start feeding next to a vein, usually alongside the midrib. In mid-or late May, the galls are visible though less than 2 mm tall, with smooth, soft, green walls, and each contains one larva (rarely, two survive to pupate in one gall). The galls grow to about 1 cm tall, harden and often turn red or purple. The first instar larva is red, the second pink, and the third is white when fully fed in September. Then it spins a papery membrane across the aperture and the gall falls to the ground. If the larva has died during development or has been parasitised, the membrane is not spun and the gall remains on the leaf, falling with it in the autumn, or remaining on the tree until spring. Most larvae pupate in the gall on the ground in the spring, developing into bright red pupae; a few pupate early and the pupa overwinters. Adults emerge in spring and produce new larvae to coincide with bursting of the next generation of beech leaves.

FIG 86. Sections through cylinder–piston galls of Contarinia ramachandrani (upper) and C. manii (lower), both on Acacia ferruginea (after Meyer & Maresquelle, 1983; Harris, 2010, has recently described the species). The upper leaflet forms a piston that fits inside a cylinder formed from the lower leaflet, enclosing the larva (black) in a chamber.

The most bizarre cecidomyiid pouches are the ‘cylinder–piston’ galls that grow on Acacia leaves in India. They are formed from two adjacent leaflets, one forming a plug or piston that fits inside a cylinder formed by the other, leaving a larval chamber at the bottom (Fig. 86). The female gall midge lays eggs between young leaflets when they are still touching one another in the bud; the eggshell is sticky and keeps the two surfaces together as the leaf expands. The larva hatches, starts to feed and nutritive tissue forms above and below it, in both leaflets. At the same time, the epidermal cells either side of the larva enlarge and form papillae that fit together, holding the two leaflets in contact. The gall grows rapidly. At first the piston and cylinder fit tightly together with a tiny larval cavity at the base; later, the cavity enlarges as the larva grows. The galls of these two species are rare, apparently found only on Acacia ferruginea in southern India, and the host plant itself is an endangered species. Sometimes these galls occur in rows down a leaf, involving in turn first the upper then the underside of a series of leaflets. This requires strict coordination between developing pairs of leaflets, and is also found in elaborate mite galls (see Fig. 70 above). Even though gall mites and gall midges are completely unrelated arthropods, both groups induce the same type of reaction in the leaflets.

CHAPTER 6

Big Buds, Cigars, Pineapples and Rosettes

THIS, THE THIRD AND FINAL chapter on open galls, describes growths that develop in buds. They start in young, undifferentiated meristem tissue whose cells can, potentially, develop in any direction, and involve the new leaves that the meristem produces. They develop either in terminal buds at the tip of a shoot, or further down the stem in a bud in a leaf axil (very unusually a new bud is induced in a novel site, e.g. on a leaf vein). All of these galls consist of young modified leaves that are bunched together because the stem fails to elongate, with one or several insect larvae or mites living on the meristem or between the young leaves. The meristem is nearly always eventually killed, preventing growth from that point in future years. These insects and mites escape from the gall through crevices and apertures in the structure of the gall; none of them is capable of boring or biting its way out. In the pineapple galls and an unusual psylloid gall (described below), the insects are sealed inside temporarily and must rely on changes in the plant tissue to allow them to escape.

Shoot-tip galls vary considerably in their complexity (Fig. 87). The least complex are the ‘big buds’ of eriophyoid mites and the cabbage-like growths of psylloids. Cigar galls in the common reed Phragmites are induced by a small group of chloropid flies (Chloropidae is not a family that is normally gall-causing). Two families of Hemiptera, the adelgids and the coccoid family Beesoniidae, cause pineapple galls, and rosettes and artichokes are caused by gall midges. For background information on these mites and most of the insects, see Chapter 4 and the boxes therein. All of these galls contain nutritive tissue and, because they develop in buds that are sites of active growth whether galled or not, they have a rich vascular supply, although this is often enhanced in the galls. With their enclosed animals, the galls draw nutrients from elsewhere in the plant. The effect of this nutrient sink is probably quite weak in the big buds, simple cabbage galls and in some rosettes, and stronger in the pineapple galls and the more elaborate rosettes and artichokes.

FIG 87. Diagrammatic sections through shoot-tip galls. (a) An ungalled bud; (b) an eriophyoid big bud; (c) the cabbage gall of Spanioneura buxi (the whole gall, not a section); (d) the cigar gall of Lipara lucens (the whole gall and a section); (e) an adelgid pineapple gall; (f) a gall midge rosette. (a, e, f, after Dreger-Jauffret & Shorthouse, in Shorthouse & Rohfritsch, 1992; b, after Westphal, in Shorthouse & Rohfritsch, 1992; c, d after Redfern et al., 2002.) Larva or nymphs are represented as black shapes.

BIG BUDS

Big buds are enlarged and distorted leaf and flower buds caused, uniquely, by eriophyoid mites. Although different species of big bud mites are specific to particular host plants, the structure of their galls is remarkably similar. Three common species are Phytoptus avellanae on hazel Corylus avellana, Cecidophyopsis psilaspis on yew Taxus baccata and Cecidophyopsis ribis, the blackcurrant Ribes nigrum gall mite (Fig. 88). All of these may be numerous enough to damage their host plants especially when these are young. C. ribis is a notorious pest which, besides restricting growth of blackcurrant bushes and reducing the fruit crop, is the vector of ‘blackcurrant reversion’, which can kill the plants.

Big buds can develop in any bud. They vary considerably in size (those described here may grow up to about 1 cm long or across) depending on the number of mites feeding inside. They start in young meristematic tissue in buds that, if undamaged, would develop into leaves and flowers the following year. These tiny new buds appear on the plants in spring when the mites are dispersing from their old galls and the buds are induced to develop precociously (normally these buds remain dormant for much of their first year). The mites insinuate themselves between the scales of the buds, one or a few per bud, pierce individual cells of the meristem and the new leaf initials and start feeding, in a manner similar in all gall mites (see Chapter 4 and Box 4.1). This results in enlargement and multiplication of cells and formation of typical nutritive tissue (Chapter 1), which develops particularly from the upper epidermis of the tiny young leaves. The nutritive tissue is very similar in these three big buds: the epidermal cells enlarge and divide and groups of cells in the layer of mesophyll beneath also divide, so producing characteristic fleshy wart-like protuberances, with the mites living between them. As the mites feed, they cause suction scars in the walls of the nutritive cells, which eventually are sucked dry and collapse. At first, these necrosed cells are few and scattered amongst the undamaged ones, but as the mite population increases, the collapsed cells increase and form dead patches on the nutritive tissue. While the gall is growing, the meristem produces some new leaf initials but fewer than in a normal bud. The gall also causes a considerable increase in the width of the stem below the bud. In hazel big bud, though not in those on yew and blackcurrant, buds in the axils of the outer leaves may remain viable and grow into spindly side shoots, which may produce deformed leaves the following year.

FIG 88. The big bud galls of (anticlockwise from top left): Phytoptus avellanae on hazel Corylus avellana; Cecidophyopsis psilaspis on yew Taxus baccata; and Cecidophyopsis ribis on blackcurrant Ribes nigrum. ((a) Michael Chinery; (b and c) Tom Higginbottom)

The lifecycle of the three mite species is similar. Adult females (the fundatrices), having mated, disperse from the old galls in April, May or early June, coinciding with development of new shoots of the host plant. In Cecidophyopsis ribis, dispersal occurs when daytime temperature averages 11°C. Besides walking, the mites may be dispersed to new host plants by wind and rain or may hitch a ride on insects, birds or mammals. They may arrive too early, before the new buds have started to develop, and must survive for a while on the surface of the plant, or they may arrive too late, after the best time for infestation. It is a chancy process and mortality of the mites during dispersal is always high. Successful females enter the buds and lay eggs, which are large in proportion to the size of the females. The mites multiply through several generations during summer and autumn and the gall enlarges. They overwinter in the gall and may produce another generation or two in spring if the weather is warm, before adults disperse to start the cycle again, and before the old galls dry out and fall off the tree or bush. At any one time, a big bud may include eggs, juveniles and adults, and a large gall may contain hundreds or thousands of individuals. If the numbers of mites in a bud is small, in C. psilaspis at least, the meristem may survive and shoot the following year. This mite can also live as an inquiline amongst the leaves of the artichoke gall of Taxomyia taxi on yew (whose gall is described below) and may even cause the death of T. taxi larvae if both species are invading a new bud at the same time.

Though vulnerable during the dispersal phase, for most of the year the mites are protected inside the gall and survival rates there are high. Pest species, therefore, can be difficult to control. Adults and larvae of C. ribis transmit blackcurrant reversion (probably a virus disease), which causes leaves to be odd-shaped, reduced in size and sometimes chlorotic (yellowed, with little chlorophyll). Shoot growth is poor due to reduced rates of photosynthesis, infected flowers are sterile so that setting of fruit is reduced and, if the attack is severe enough, the plant dies. The disease spreads rapidly through a crop and fruit production becomes unprofitable if more than 5 per cent of buds are infested. Control of the mites by spraying with an acaricide is effective only when they are dispersing, and this period is difficult to time accurately as it varies with climatic conditions. The mites do not seem to be regulated by predators. A chalcid species, Aprostocetes eriophyes (Hymenoptera: Eulophidae), feeds on the mites in the gall (on C. psilaspis as well as C. ribis and perhaps on other species of big bud mites too) and predatory non-eriophyoid mites are common, but these probably cannot increase in numbers fast enough to control outbreaks of the pest. The best chance of control is the production of varieties of blackcurrants that are resistant to the mite. C. ribis and reversion disease are European in origin. Both have accidentally been imported into North America and New Zealand where they are important pests.

PSYLLID SHOOT TIP GALLS

Only a few psylloids cause galls in apical buds or in shoot tips and most are not well known. Two species are described here: Spanioneura buxi (Fig. 89), a British and European species that causes a simple open cabbage-like gall on box Buxus sempervirens, and a North American psyllid, Pachypsylla celtidisgemma, which causes a much more complex closed bud gall on hackberry Celtis occidentalis (Fig. 90). They illustrate two extremes of psylloid gall complexity.

The box gall is common in Britain. Leaves at the shoot tip are spoon-shaped, curved upwards, broadened and crowded into a more-or-less compact ‘cabbage’. Behind the apex the galling effect becomes progressively weaker with individual leaves concave or with depressions. Several greenish waxy nymphs live on the upper surface of affected leaves. The galls are of minor economic importance in Europe, where young hedging plants may appear unsightly, but it can be a serious pest of ornamental box in gardens in North America where the species has been accidentally introduced. There is one generation a year. Eggs are laid in crevices on box twigs in August and overwinter, hatching in the spring. First instar nymphs are flattened in shape and vary in colour from yellow to deep green or brown. They invade the opening buds and start feeding on the upper surface of the young leaves, causing them to curl upwards. There may be up to ten nymphs per bud, so that the resulting cabbage gall varies in size and has varying amounts of waxy deposits, produced by the insects. The nymphs grow rapidly and adults appear in May. They are about 3 mm long and active, jumping and flying when disturbed and, in summer, are bright green with brown markings; they become yellow-brown in autumn. They mate in late July or August and lay eggs soon afterwards, after which they die.

FIG 89. The ‘cabbage gall’ of Spanioneura buxi on box Buxus sempervirens. (Tom Higginbottom)

FIG 90. Gall of Pachypsylla celtidisgemma on hackberry Celtis occidentalis. (a) The mature gall, entire (with an ungalled bud higher up the right-hand twig); (b) the young gall in section, late July; (c) one chamber of a mature gall in section, early January (modified after Walton, 1960). Insect (black); nutritive tissue (fine stipples); granular plug in opening (coarse stipples); woody layer (cross-hatched).

The structure and development of the hackberry psylloid gall is well known. The gall has one or several chambers (up to about 13) that appear externally as separate bulges, each containing one nymph. The gall is green at first and covered with bud scales; it grows beyond the scales, which persist at its base, and becomes light brown and woody with a smooth polished surface. Adult males and females appear in spring, mate and the female lays eggs (often >100 per female), singly or in groups of two to five, inserted and glued into the vein angles or alongside veins on the underside of new leaves. After four or five days, tiny oval flat nymphs hatch; they are about 0.3 mm long, dusky lemon yellow in colour with red eyes, and the body is fringed with fine hairs. This first instar has strong legs and is active. It walks to the base of its leaf and down the petiole to one of the buds in the axil of the petiole. One or several nymphs crawl between the bud scales and each settles at the base between two leaf initials. There are usually three buds in an axil and all may become galled. The nymph starts to feed after inserting its stylets into the meristematic tissue of the bud–its stylets are more than twice the length of the body and much longer than in later instars. Only the tips of the stylets penetrate the plant tissue; their length allows them to be inserted and withdrawn several times at some distance from the nymph’s body without it moving its position. Pierced cells are sucked dry, adjacent cells shrink and collapse, and those a little further away start to divide and grow, so that each nymph becomes enclosed in an individual crater. The gall grows rapidly around and over the nymph, imprisoning it inside. By late July, the gall is nearly full-sized and the nymph has moulted into its second instar; now it is entirely enclosed in the gall chamber, although the path of the original opening is still apparent (Fig. 90b). The nymphs develop through five instars and the size of each chamber enlarges–faster than the nymph grows so allowing it to move about, feeding in several different positions. The nymphs feed on the nutritive cells of the parenchyma (Fig. 90c), a layer supplied with vascular strands although these are not pierced by the stylets. As feeding progresses, the parenchyma cells collapse, enlarging the chamber, and the thickness of the gall wall decreases. In August the outer layers of the gall become woody; the gall is now full-sized with each nymph in its third instar. The woody layer thickens through the autumn and winter (Fig. 90c) while the nymph develops through its fourth and fifth instars, the final one lasting from November to the following June. As the gall matures, a granular plug develops in the wall of each chamber, the plug formed from sclerenchyma cells separated by large air spaces, and this becomes a zone of weakness through which the fifth instar nymph can escape. By this stage the nymph is large, completely filling its chamber, and has a tough caudal shield on the abdomen armed with strong spines. It braces itself against the wall of its chamber and pumps up the abdomen to push against the plug, and this dislodges it and allows the nymph to crawl out backwards. This escape process takes several hours. Old galls with neat round holes remain on the hackberry trees for several years, no doubt providing convenient hiding places for a community of ‘successori’ (unrelated small animals that shelter in the galls).

CIGAR GALLS, CAUSED BY LIPARA SPECIES

Lipara belongs to the family Chloropidae (Diptera), most of whose species are not gall causers. Lipara species have specialised on grasses, specifically the common reed Phragmites australis, and four of them cause a range of damage from stem boring to complex galls. All species cause the internodes at the top of the shoot to shorten so that the leaf sheaths overlap each other: Lipara similis is a stem borer, while L. pullitarsis, L. rufitarsis and L. lucens form galls of increasing complexity. The stem is thickened to varying degrees caused by the telescoping of the apical internodes, and this results in cigar shapes of increasing thickness. In all, flowering is prevented and the growing point eventually dies (Fig. 91 shows the gall of L. lucens, the most elaborate).

Phragmites australis is a perennial grass. Its shoots die down each year and new ones grow in April, May and June from underground rhizomes. Flowering starts in July and the shoots dry and die back in autumn, so that food for shoot feeders is available for only a few months each year. Adults of all the Lipara species emerge from their overwintering sites in late spring and summer (May to early July), mate, and the females lay eggs one at a time on the surface of new shoots. A week or ten days later, the ivory-white larva hatches and crawls upwards, entering the shoot under the edge of the sheath of the uppermost leaf. The larva gnaws and burrows through several layers of enwrapped leaves to a position just above the apical meristem (the growing point), where it feeds on newly emerging leaves. At this stage, the meristem is not damaged and the stem scarcely thickened, although elongation of the internodes is prevented. Only one larva per shoot can survive, and it initiates the gall when it has reached its position just above the meristem. A dark brown rotting channel above the growing point marks the activity of L. similis, whose shoot never thickens into a gall. In L. pullitarsis, a European species not found in Britain, the young inner leaves above the meristem become distorted and the larva lives enveloped by them, the shoot thickening slightly to form a simple gall. In L. rufitarsis and L. lucens (see Fig. 87d), a distinct chamber develops in the pith below the meristem, which causes an obvious thickening of the shoot. The gall is full-sized by August, nearly twice the diameter of a normal shoot in L. rufitarsis and 2½ times or more in the more robust gall of L. lucens (Fig. 91). The larva is now in its third and final instar, but is not full-grown and is still feeding on the enwrapped leaves above the meristem. Below the meristem the gall is packed with parenchyma pith tissue. The larva now gnaws through the meristem with its tough mouth hooks so killing it, and feeds on the prepared pith, carving out a cavity that enlarges as it feeds. In the L. lucens gall, sclerenchyma cells develop in the chamber walls, increasing in abundance so that, by the time the larva is fully fed in late summer, the gall is hard and woody. In L. rufitarsis, there is no sclerenchyma development and its gall remains relatively soft. The full-grown larva overwinters head upwards at the bottom of the chamber, pupates there in spring, and the adult emerges in early summer by crawling up through the channel in the meristem and enwrapped leaves above, cut earlier by the larva.

FIG 91. Gall of Lipara lucens on common reed Phragmites australis. (Michael Chinery)

In mainland Europe, the three gall causers can be found in the same reed bed and so could compete for the best shoots. They illustrate two lifecycle strategies, which probably reduce the chance of competition:

(i) The adult of L. pullitarsis emerges from its overwintering site in April or early May, a month earlier than the other two species. At this time, the reed shoots are very small and are only just appearing above the soil, and any of them can support L. pullitarsis and its simple gall. This species does better on the more vigorous shoots that eventually grow large and thick (5–8 mm diameter) and produce sufficient new leaves to supply the larva. The drawback to this strategy is that poor weather (cold, frosty, wet) in spring can kill the adults and delay growth of the plants so that the hatchlings do not survive. The females lay numerous eggs scattered over the reed bed so that chances are high that some, at least, will survive.

(ii) L. rufitarsis and L. lucens adopt a more cautious strategy. The adults emerge in May to June when the weather is milder and late frosts and floods are less likely. But the plants are larger now and more resistant to attack, the leaves and stems with a high concentration of silica that is typical of grasses. Galls will only form successfully in young, thin, less robust shoots, less than 3.5 mm in diameter. The females carefully select shoots on which to lay their eggs and lay only one egg per shoot so that the hatchlings, which cannot leave the shoot on which they were laid, do not have to compete for the growing point. They lay fewer eggs than in L. pullitarsis and egg and hatchling mortality is lower. The older larvae feed on the parenchyma in the gall chamber below the meristem, which is more nutritious than young leaves–the parenchyma cells contain less fibre and more proteins and carbohydrates, materials continually supplied by extra vascular strands that end on the surface of the chamber. The cells are easy for the chloropid larvae to feed on, slashing into them with their strong mouth hooks.

PINEAPPLES

Pineapple galls are distinctive growths that develop in young shoots, forming cones of modified leaves or scales that overlap due to telescoping of the shoot axis (Figs 92 and 93). They are caused by two families of plant bugs (Hemiptera), the conifer woolly aphids (Adelgidae) and some unusual scale insects (Beesoniidae). Both groups cause galls on only one host plant genus, the adelgids on spruce (Picea, Pinaceae) and the scales on the tropical tree Shorea curtisii (Dipterocarpaceae). Although the final cone-like shape of these galls is similar and there are parallels in their development, they are initiated in quite different ways: the adelgids attack terminal or axillary buds, while the Gallacoccus scales induce novel bud-like structures with a new meristem on the petiole or midrib of leaves. There are major differences in their lifecycles too. Adelgids, like other aphids, have complex lifecycles with alternation of sexual and parthenogenetic (thelytokous) generations, a preponderance of females and often alternation between two host plants. The usual coccoid lifecycle is simple and sexual, with males common and no thelytokous parthenogenesis. General information on both groups is given in Chapter 4 and Boxes 4.5 and 4.6.

FIG 92. Pineapple galls of Adelges abietis on Norway spruce Picea abies; younger gall on left with chambers not yet opened, older gall on right with opened chambers (Michael Chinery).

FIG 93. Diagrammatic sections through mature galls, from left to right, of Adelges laricis, Gallacoccus anthonyae and G. secundus (after Meyer & Maresquelle, 1983). The dotted outline indicates the final position of the fundatrix; black dots indicate her offspring.

As in other Hemiptera, it is the first instar female nymph, the fundatrix, which initiates the gall. She is tiny (< 0.5 mm long) and has long stylets looped beneath her body. She is relatively active with well-developed legs and needs quickly to find a site to gall. Once settled with her stylets inserted into the plant tissues, the gall develops rapidly and she moves very little or not at all. Despite having settled in one spot, the stylets can probe and explore a considerable area and it seems that the pattern of probing and the effect of substances in the saliva induce the gall to develop and determine its final shape and structure (how this occurs is not understood). She initiates two areas of nutritive tissue, both some distance away from her body, and vascular elements appear and link with the normal vascular bundles of the stem, so ensuring a continual supply of nutrients. Unlike the gall-causing Hemiptera described in Chapters 4 and 5, in which the gall tissue develops around the fundatrix so that she sinks into a pit or becomes surrounded by a leaf fold or pouch, pineapple gall fundatrices remain at or near the edge of the gall. Also unlike true aphids, the fundatrices of Adelges and Gallacoccus do not tap directly into the phloem of the vascular bundles. The first area of nutritive tissue provides food for the fundatrix and establishes the nutrient sink effect of the developing gall. The second area ensures food for her offspring, with chambers for them prepared and ready before they hatch from their eggs. By the time they hatch, the gall is full-sized and the fundatrix has died. In all pineapple galls the offspring of the fundatrix are temporarily imprisoned inside their chambers. They cannot bite or burrow their way out and will be released only when changes in the plant tissue cause the gall to dry out and the chambers to open. This is typical in Hemiptera, occurring also in simple leaf rolls and in pouch galls (see Chapters 4 and 5). Examples of adelgid and eriococcid galls are described in more detail.

Adelgidae, the conifer woolly aphids

There are 12 British species of Adelges and the related Pineus and five of these cause pineapple galls on their primary host, spruce Picea. The development of the galls and the lifecycles of adelgids are known in detail. Most species are heteroecious, alternating to a secondary host in order to complete the lifecycle; this is always another conifer (e.g. Larix, Abies, Pseudotsuga, Pinus) on which galls do not develop. The basic lifecycle is a complex pentamorphic holocycle (i.e. involving five forms or morphs) and takes two years to complete. Most individuals are female and without wings; only one stage involves males, and winged females appear only twice, when flight to the alternate host occurs. Some species are anholocyclic, in which the lifecycle is simplified, with only two morphs, no secondary host, no males (and no sexual stage), and lasts only one year.

Two common species are described: Adelges laricis has the full pentamorphic holocycle, alternating between Norway or sitka spruce Picea abies or P. sitchensis and European larch Larix decidua, and Adelges abietis is anholocyclic, on Norway or sitka spruce only. Their galls are similar (Figs 92 and 93). The A. laricis gall is soft and fleshy, globular, pale yellow or greenish white in colour with a waxy texture, and with its modified needles short, forming points on the surface. It is often found at the tips of shoots with no growth occurring beyond it in future years. The slits to the chambers are usually the same colour as the rest of the gall and open to release the enclosed aphids in June. The A. abietis gall is harder and longer than wide, only slightly paler green than normal shoots and its modified needles are longer than in A. laricis. It tends to be found at the base of shoots, with growth often continuing beyond the gall the next year, and often occurs in clusters of two or three galls. The slits to the chambers are characteristically a deep pink or purple and they open later than in A. laricis, in August or September.

Development of the galls is similar, though with differences of detail. The fundatrix hatches in the autumn and settles on the surface of a dormant bud, one that would produce a shoot next year. A. abietis settles at the base of a bud on a vigorous primary shoot and A. laricis part way up a bud of a weaker side shoot. At this stage, the meristem has produced several leaf initials and is enclosed and protected by brown bracts. The fundatrix inserts her stylets into the base of the bud, probing over a considerable area and initiating two patches of nutritive tissue (Fig. 94, NT1 and NT2), and overwinters, feeding very little; she will remain in this spot until she dies. In early spring, feeding resumes, causing the bases of the new needles to swell, and the gall is obvious before the new shoots have shed their bracts. The path of the stylets differs in the two species. In A. laricis, they pass downwards inside the outer bract and initiate NT1. They then turn upwards through the collenchyma layer (which marks the junction between the previous and current years’ growth) and form NT2, which is extensive and prevents growth of the bud in the following year. In A. abietis, NT1 develops close to the main vascular tissue of the stem and NT2 affects only one side of the shoot, allowing normal growth in future years. By April, the fundatrix is adult, the gall is full-sized and the chambers for her offspring are prepared. She lays a batch of eggs that remain attached to her posterior end, covered with white woolly wax. They hatch into active crawlers (gallicolae), which crawl up the gall to enter the chambers–hatching and opening of the chambers must be synchronised otherwise the crawlers will die on the surface of the gall. The apertures are bordered with hairs that interlock to close them and the gallicolae feed on nutritive tissue lining the chambers. The stylets of the first instars are short, capable of penetrating only one cell layer; they become longer as the nymphs grow and, in the final instars, they reach the vicinity of the new vascular strands. The galls of both species form strong nutrient sinks. In A. laricis, the most rapid growth coincides with the spring flush of leaves in late May, while in A. abietis, the gall and gallicolae sequester nutrients intended for growth of the cones and seeds. When the gallicolae are mature, the chambers open due to shrinkage of the aperture hairs. It is the gallicolae that determine when the chamber apertures open–if nymphs of one species are transferred to the gall of the other species, the timing of opening of the chambers will change too.

FIG 94. Diagrammatic sections through the developing galls of Adelges laricis (on the right) and A. abietis (left); the arrows indicate the path of the stylets of the fundatrix (after Rohfritsch & Anthony, in Shorthouse & Rohfritsch, 1992). (For full explanation, see text.) Fundatrix (black); first (NT₁) and second (NT₂) patches of nutritive tissue; collenchyma tissue (wavy lines); vascular bundles (dashed lines and open circles).

Although their galls are similar and occur on the same host species, the lifecycles of A. laricis and A. abietis are very different (summarised in Fig. 95 and Table 6.1). In A. laricis, there is a brief sexual stage, with tiny sexuales that do not feed. They mate and each female lays a single egg that hatches into the fundatrix; when adult, the fundatrix produces a clutch of eggs that hatch into the gallicolae and inhabit the gall. When adult, the gallicolae fly to a larch tree, lay eggs that hatch into sistens and these settle as first instar nymphs near a dormant bud or in a crevice in the bark, and overwinter. They are black and non-waxy. In April, they lay masses of grey or pale orange eggs that hatch into progrediens and feed on the new larch needles, causing some distortion if numbers are high. The progrediens mature either into wingless adults that remain on the larch tree and produce further generations of progrediens, or into winged adults that become the sexuparae, which migrate back to spruce trees. The progrediens produce copious wax and honeydew and, if infestation is heavy, the larch tree appears bluish in colour. Thus, asexual reproduction occurs in four stages in this species. In A. abietis there is no sex and all reproduction is carried out by the fundatrices and gallicolae. Stage 1, the sexuales, has been lost, suggesting that the potential for genetic variation is less in this species.

FIG 95. (a) Lifecycle of Adelges laricis with five morphs (1–5) and two host plants; (b) the simpler lifecycle of A. abietis with only two morphs (2 and 3) and one host plant, and no sexual stage (for further explanation, see text and Table 6.1; simplified after Carter, 1971; Meyer, 1987).

These adelgid species are native to Europe. A. abietis was introduced accidentally to North America in the early nineteenth century and occurs on several spruce species, including sitka spruce. Individual trees vary in their resistance to the adelgids–some are completely resistant while others are highly susceptible, and sometimes galls are so common that growth of the tree is adversely affected. The size of the galls varies with the vigour of the shoots too. The variation in resistance seems to be genetically rather than environmentally controlled.

Gallacoccus galls on Shorea curtisii

Gallacoccus anthonyae and G. secundus cause complex galls on Shorea curtisii (Dipterocarpaceae) in Malaya. These galls are most unusual–they develop from new meristems induced to form on the petioles and veins of the leaves, sites that never initiate new growth in the normal plant. The resulting galls (Fig. 93) are cone-shaped, 1–2 cm long or more, and consist of overlapping scale-like leaves–three in the G. secundus gall, which develops on the upper side of leaves, and 24 in G. anthonyae, which hangs down from the underside. The G. secundus gall contains just one chamber, while the mature gall of G. anthonyae consists of several chambers either side of the main axis, forming a structure reminiscent of an adelgid gall. The chambers contain the offspring of the fundatrix, which are released when the gall dries out and the chambers open.

The two Gallacoccus galls start in the same way. The first instar fundatrix nymph settles on the under surface of a young petiole or midrib, inserts its stylets, and creates a bulge with a new meristem. As in the spruce adelgid galls, two patches of nutritive tissue develop (Figs. 96a, 97a, NT1 and NT2). The nymph moults at an early stage and it is the second instar that settles on the new meristem, covered by a new leaf initial. In G. anthonyae, the new meristem grows quickly producing many new overlapping leaves and forming a bud (Fig. 96b). The fundatrix nymph sits in the axil of the first leaf, now at the base of the gall, and feeds from nutritive tissue NT2 (NT1 is abandoned on the left). While the bud grows, she becomes adult and mates with a visiting male from a nearby mature gall. Then she moves upwards into the axil of the sixth leaf and initiates a third patch of nutritive tissue (Fig. 96c, NT3; the open shape shows the original position of the fundatrix). She produces a number of male and female offspring (without laying eggs–she is viviparous) and then dies. These nymphs crawl up the gall and settle on the underside of the new leaves (Fig. 96d; the positions of the fundatrix when nymph and adult are indicated by open shapes, and the three patches of nutritive tissue, now abandoned, are shown). Fig 97 shows the development of the G. secundus gall, whose new meristem ceases to function after producing only three leaves. After mating, the adult fundatrix moves into the space between leaves two and three (Fig. 97c), which becomes the main cavity of the gall, with greatly thickened walls. Her offspring, male and female, line the walls of the cavity (Fig. 97d) and she dies (open shape indicates her position when a nymph and the three patches of abandoned nutritive tissue are also indicated).

FIG 96. Development of the gall of Gallacoccus anthonyae on Shorea curtisii (Dipterocarpaceae) from initiation (a) to the mature gall (d) (after Meyer, 1987; see text for further explanation). Black shapes indicate nymphs and adult coccoids; open shapes indicate positions of earlier stages; NT₁, NT₂, NT₃ = successive patches of nutritive tissue (indicated by stippling).

FIG 97. Development of the gall of Gallacoccus secundus on Shorea curtisii from initiation to maturity (a–d) (see text for explanation, labels as in Fig. 96; after Meyer, 1987).

Unlike adelgids, the Gallacoccus fundatrix produces both male and female offspring that inhabit the gall, equivalent to the adelgid gallicolae. The subsequent development of the males and females differs. The male nymphs grow up into winged adults, leave the parental gall and fly to a newly developing gall, where they mate with the new adult fundatrix. The females develop more slowly, leaving the parental gall while still in their first instar. They disperse to new sites and initiate new galls that are partially grown (Figs 96b and 97c) before they become wingless adults ready to mate. These galls are tropical and all stages can be found on the trees at the same time, unlike the spruce adelgids that live in a seasonal temperate climate.

Despite forming on the surface of a petiole or vein, the new meristem appears similar to normal meristems, budding off new leaf initials in the same way. It forms when a patch of epidermal and underlying cortex cells de-differentiate and rejuvenate, a process stimulated, it is assumed, by substances in the saliva of the fundatrix nymph, injected as it feeds. Future nutrients are ensured as vascular tissues develop linking the new bud with the vascular bundles of the stem. An actively feeding fundatrix nymph is necessary for the new bud to function. When the fundatrix dies, or if it dies prematurely, the cells of the gall revert to parenchyma and it stops growing.

Mangalorea hopeae (Beesoniidae) on Hopea ponga (Dipterocarpaceae)

I end this section on pineapple galls with a description of another unusual coccoid gall, which was mistaken for the fruit of a tree for nearly three centuries (see also Chapter 14). The gall is a spiny green ball, 1–3 cm across when mature, single (Fig. 98) or occurring in clusters of two, three or more in the axils of leaves. They start in buds; there is no initiation of new meristems as in the Gallacoccus species. The host tree, Hopea ponga, is common along the southwest coast of India, where it is regularly browsed by stock and coppiced for firewood, and is often restricted to small bushes only 50 cm high. The spiny spherical galls are similar in appearance to the fruits of breadfruit (Artocarpus, Moraceae) and durians (Durio, Bombacaceae), plant families that are related to the Dipterocarpaceae to which Hopea belongs. But no spine-like structures occur in the dipterocarps; the spines seem to be a throwback to an ancestral trait, evolved before the Dipterocarpaceae became distinct from related families.

FIG 98. The gall of Mangalorea hopeae on Hopea ponga (Dipterocarpaceae). (a) two single galls in leaf axils; (b) section through a young gall containing a reproducing female (black) in a chamber at the base of the gall (after Jenkins & Mabberley, in Williams, 1994).

The first instar female nymph, the fundatrix, initiates the gall. She inserts her stylets deep into a bud, growth beneath her stops and surrounding tissues grow so that she sinks into a hollow that nearly encloses her. The gall has an ovoid central core containing sclerenchyma and vascular tissue linking to the vascular bundles of the stem, and an outer zone of tough woody spines (Fig. 98b). Normal buds and young stems are hairy and it seems that the spines develop from these single-celled hairs–they become multicellular and lignified with vascular strands and are covered with unicellular hairs. Several fundatrix nymphs may feed together, so inducing clusters of galls on a single bud, sometimes of a range of ages. In the young gall, the spines are closely packed except at their tips which taper to points. The fundatrix feeds from the same spot throughout her life. She soon becomes adult and produces male and female offspring, and gaps appear between the bases of the spines due to the widening of their tips and enlargement of the core–except around the fundatrix where the spines remain closely packed, without gaps, so protecting her. Her male offspring crawl into the gaps between the spines and feed, and grow through three instars. When adult, the males are winged and are released when the core enlarges further, causing the spines to separate. These males have no mouthparts so cannot feed, and they live only a short time. They disperse and mate with first instar females that are sitting on the surface of new stems, having left the parental gall as soon as they were born–these are precocious nymphs, capable of mating before they are adult! The mated female nymphs become the fundatrices of the next crop of galls.

Galls of Adelges, Gallacoccus and Mangalorea hopeae are the most complex to be caused by Hemiptera. Compared to aphids, their mouthparts are relatively short and tend not to be inserted very deeply; they feed on cortical parenchyma rather than phloem and induce a short-distance transport system in the gall that links up with the vascular tissue of the shoot, so ensuring that a continual supply of nutrients is directed to the gall and insects. Adelgids are more closely related to aphids than the coccoids and have similarly complex lifecycles dominated by thelytokous parthenogenesis. In Mangalorea and Gallacoccus, reproduction is sexual and the lifecycle simple, and each fundatrix produces just one batch of offspring. The spiny ball of Mangalorea hopeae is not much like a pineapple gall but, like the adelgid and Gallacoccus galls, it is much more complex than other hemipteran galls.

ROSETTES AND ARTICHOKES

Rosettes and artichokes are clusters of modified leaves in a bud at the shoot tip or in a leaf axil, and are caused by gall midge larvae infesting the meristem and the tiny new leaves that develop from it. All are caused by cecidomyiids. Rosettes and artichokes are very similar and often the terms are interchanged–rosettes tend to be wider than long and artichokes longer than wide (Fig. 99). The artichoke gall of oak (Quercus, Fig. 152b, Chapter 9), caused by the gall wasp Andricus foecundatrix, is not included here. Although superficially similar to gall midge artichokes, it properly belongs in Chapter 9 because it causes a closed gall; the larva, and later the pupa, is enclosed in an inner gall that the adult must bite through to escape. Cecidomyiid larvae and adults do not have mouthparts capable of doing this and, like all the gall causers in this chapter, they push their way out of a gall that is never permanently closed.

FIG 99. Rosette (upper) and artichoke galls (lower) caused by (clockwise from top left) (a) Dasineura crataegi on hawthorn Crataegus monogyna; (b) Rabdophaga rosaria on white willow Salix alba; (c) Spurgia euphorbiae on cypress spurge Euphorbia cyparissias; (d) Two-year gall of Taxomyia taxi on yew Taxus baccata. ((a) & (d), Keith Harris; (b) & (c), Michael Chinery)

The structure of rosettes and artichokes is similar although some are looser and less neat than others. In all, there are more leaves than normal and the stem fails to lengthen so that the new leaves remain bunched together, and the meristem is always eventually killed (unless the larva dies very young). In the simpler galls, several larvae live amongst the inner leaves and on the meristem and no distinct chamber is formed. The more complex, which exert a stronger nutrient sink, enclose a single larva in a gall chamber above the meristem, and the tip of the shoot is distinctly widened. Examples of the simpler ones are Dasineura crataegi (Fig. 99a) on hawthorns Crataegus monogyna and C. laevigata, Spurgia euphorbiae (Fig. 99c) on cypress spurge Euphorbia cyparissias and Rabdophaga heterobia on basket or almond willow Salix triandra, while more complex artichokes are caused by Taxomyia taxi on yew (Taxus baccata, Fig. 99d) and a North American species, Rabdophaga strobiloides on heart-leaved willow Salix eriocephala.

Fig. 100 shows a section through a typical rosette that contains several larvae between thick, fleshy, modified inner leaves (the gall of Spurgia euphorbiae on cypress spurge). The mesophyll thickens and loses its chloroplasts, and there is little differentiation between the palisade and spongy layers. The meristem becomes broad and flat with an irregular surface formed from clusters of nutritive cells. The inner leaves become nutritive; moving outwards from the meristem, the leaves become progressively less modified and longer with only their inner surfaces nutritive. The larvae are yellow or orange when mature and move about actively amongst the inner leaves. The outer gall leaves are green and tougher, more similar to normal leaves in structure although they are shorter and broader.

FIG 100. Diagrammatic section through the rosette of Spurgia euphorbiae on cypress spurge Euphorbia cyparissias (after Meyer & Maresquelle, 1983; larvae (black); stippling shows the extent of the nutritive tissue; scale indicates 1 mm).

Similar rosettes are the button-top galls of Dasineura crataegi on hawthorn (Fig. 99a) and Rabdophaga heterobia on the almond willow. D. crataegi is often common in regularly trimmed hedges. It is unusual in that the underside of its leaves develop small, fleshy, red or green projections. These structures are extensions of the epidermis and mesophyll, though without palisade and spongy layers. Although much larger, they resemble the hairs on normal leaves, and may have no specific function–they may just be a by-product of the galling process. All of these rosettes develop rapidly in summer and the meristem and inner leaves die when the larvae are full-grown. There are two or more generations of galls a year, with the larvae pupating in cocoons in the gall or dropping to the soil. In R. heterobia, the first generation infests the male catkins of the willow, causing a downy swelling at the base or tip rather than a leafy rosette. The artichoke galls of Taxomyia taxi (Fig. 99d) and Rabdophaga strobiloides are more highly structured; they contain only one larva that sits on the meristem in a distinct central chamber, surrounded by modified leaves. As in the simpler rosettes, the gall leaves are modified, less efficient at photosynthesis than normal, with the inner ones without chlorophyll.

The lifecycle of R. strobiloides is probably typical of rosette and artichoke gall midges with one generation a year. The females lay eggs in late May on or near the terminal buds of the willow, the hatchlings enter these buds and feed on the meristem, which modifies into nutritive tissue, and the gall grows and becomes full-sized in August. The full-grown larva overwinters, pupates in a cocoon in the gall in April and the adult emerges in May.

The lifecycle of T. taxi is much more complex. This gall midge is unusual in that it can develop from egg to adult in either one or two years, and these two lifecycles produce quite different galls. Most individuals develop in two years, causing the artichoke gall (Fig. 99d) in the second year. The one-year lifecycle, usually occurring in only 5–10 per cent of a population, induces enlarged buds only about 5 mm tall (Fig. 101). The insects develop through the same stages, an egg, three larval instars, a pupa and adult, but the first instar lasts about 14 months in the two-year galls with an overwintering period, and only two months in the one-year galls. Eggs are laid in late May and June on the leaves of new elongating shoots and hatch about ten days later (up to three weeks later if June is cold and wet). The hatchlings crawl towards the tip of the shoot and enter axillary or terminal buds–the terminal bud is favoured although growth and development of the larva are similar in either. These are tiny dormant buds that, if not infested, would produce the next year’s shoots. The apical meristem becomes flattened and slightly enlarged but otherwise the larva has little effect and production of leaf initials proceeds slowly, as in the normal buds, through June and July. In August, when the galls are two months old, development of the one-and two-year galls diverges. In the buds that will become two-year galls, leaf production speeds up so that by October there are twice as many new leaves as in ungalled buds (Fig. 102). In buds destined to become one-year galls, the meristem erupts in August into a fleshy pad consisting of nutritive tissue, and this event signals the end of leaf production. The pad develops in the two-year galls too, but not until one year later and after many more leaves have been produced. So, the one-year gall remains small with only about 30 leaves, compared to 130 or so in the artichoke gall and 75 in ungalled buds. In normal buds, development stops during the winter when there are about 35 leaf primordia in each bud, while the galled bud (two-year gall) continues to grow, though only slowly if temperatures are cold. As the weather warms in spring, leaf production in the galled and ungalled buds increases again, faster in the galls, and by August these have produced their maximum number. The full-grown artichoke gall contains five types of modified leaves (Table 6.2). The two inner leaf types (D and E) make up the gall chamber around the larva; they form a ‘bud’ similar in size to the one-year gall. At the same time as the gall leaves are multiplying, the normal shoots are elongating, a process that is totally inhibited in the galls.

FIG 101. One-year gall of Taxomyia taxi at tip of shoot; a young fruit lies below it. (Robert Cameron)

FIG 102. Average number of leaves produced by normal buds of yew Taxus baccata and by buds galled by Taxomyia taxi over two years, with date of pad eruption indicated. Pad eruption is also shown in one-year galls for comparison (modified after Chappell & Lovett, 1977).

TABLE 6.2. Characteristics of the five types of leaves in the artichoke galls of Taxomyia taxi on yew Taxus baccata compared to normal yew leaves (+ present,–absent; modified after Chappell & Lovett, 1977)

Fig. 103 summarises the two lifecycles of T. taxi and the changes in their galls. The first instar flattens the meristem and, particularly in the two-year gall, stimulates it to produce more leaf primordia than normal. The first instar grows very little. The eruption of the meristem into a fleshy pad in August, two or fourteen months later, changes everything: it stops producing new leaves, and the rapid development of nutritive cells stimulates the larva to moult into its second instar and to grow very rapidly–in the month after eruption of the pad, its weight increases seven-or eight-fold. In October, it moults into the final instar and grows until prevented by cold weather. It grows again in the spring, when its weight doubles, and pupates in the gall without a cocoon in late April. Adults emerge a month or so later. The pad cells remain functional while the larva is feeding; in April, when it is full-grown, they collapse and die. After the first instar, development is identical in the one-year and two-year lifecycles. The factors influencing the duration of the first instar are completely unknown.

Other gall midges are known to have varying lifecycle times. Two species that cause leaf pouch galls in Japan are usually annual but a few individuals take two years to develop due to an extended first instar. Adults of a single brood of wheat blossom midges emerge from the soil year after year over several years. Variation may be a strategy to avoid or reduce parasitism. But a strategy with a dominant two-year lifecycle is unusual: apart from T. taxi, no other examples are known in gall midges. Although the yew gall midge is capable of developing in one year and would produce twice as many offspring if it did so, mortality caused by parasitism is higher, and the two-year parents on average leave more descendants than the one-year adults, a strategy that clearly benefits the species. Survival and mortality are discussed further in Chapter 10.

FIG 103. Diagrammatic summary of the 2-year and 1-year lifecycles of Taxomyia taxi in their galls on yew Taxus baccata (the black shape indicates the larva on the meristem; nutritive tissue (stippled); modified after Redfern, 1997). The female lays an egg either on a bud or on a leaf of a new shoot.

PART THREE

Closed Chambered Galls

The galls in Chapters 7, 8 and 9 share a common theme in that they contain chambers that are more or less closed from the outside world. The gall causers are enclosed by plant tissue and they escape from the gall by chewing their way out, or pushing out through a weak spot or through a tunnel prepared earlier by the larva, or being released when the gall rots and disintegrates. Except for a few nematode galls, all are caused by insects.

Flower head galls are a distinctive group and form the subject of Chapter 7. They include galls in thistle, knapweed and burdock heads (in the Asteraceae) and in fruits of figs (Moraceae). Two families of insects that have not appeared in previous chapters cause most of these galls: gall flies (Tephritidae, order Diptera) gall the Asteraceae and fig wasps (Agaonidae, order Hymenoptera) gall the figs. Chapter 8 includes galls in stems and roots, caused by a variety of insects and by some nematodes. Nematodes are unusual gallers and they have not appeared earlier. The insects include sawflies and gall midges that occur in earlier chapters, and beetles (Coleoptera), moths (Lepidoptera) and gall wasps (Cynipidae, order Hymenoptera) that are new galling groups. Closed galls in leaves and buds form the subject of Chapter 9; these are caused by sawflies and gall midges, and by gall wasps which make up the bulk of the chapter.

Gall-causing is unusual in nematodes, beetles, moths, sawflies and tephritid flies, although some of their galls are well known and characteristic: for example, the woody flower head galls in thistles (tephritids) and the bean galls in willow leaves (sawflies). In gall midges, fig wasps and gall wasps, gall formation is the rule rather than the exception and, in gall wasps especially, some galls are fantastic and bizarre structures.

As in the galls described in previous chapters, these galls develop in young, undifferentiated tissues with cells that can develop in any direction–or plant tissues are induced to return to a younger state. Gall midge galls develop in response to sucking and feeding by the first instar larva and there is little or no destruction of plant cells. Neither the female nor the egg plays any part in the process. Larval and adult nematodes feed by piercing and sucking, and all stages may be able to cause swellings (though little is known about most nematode galls). In all of the other groups considered in these three chapters, the larvae feed by chewing, destroying plant cells in the process, and growth of the galls depends on feeding by the young larvae. The ovipositing female and the egg play no part in gall development in beetles, moths and tephritid flies, but are responsible for an initial small swelling in gall wasp and sawfly galls and also, probably, in fig wasps.

CHAPTER 7

Flower Head Galls

THE FLOWER HEADS OF this chapter are specifically those of composites (family Asteraceae) and of figs (Moraceae). In these plants, the flowers are numerous small florets clustered together into a compact inflorescence at the tip of a stem or in the axil of a leaf. In the Asteraceae, the inflorescence is supported on an enlarged receptacle and is enclosed and protected by tough spiny bracts–particularly spiny in thistles, knapweeds and burdocks, the main host plants included here. In figs, the inflorescence is turned inside out, with florets enclosed and protected inside the ‘fruit’. Both types of inflorescence provide a rich food supply for insects to exploit; a large number of seeds are clustered together in one place and are provided with a rich vascular supply. Apart from the gall causers, many non-gallers feed inside the heads too.

Gall flies (Tephritidae) specialise on thistles, knapweeds and other Asteraceae and cause galls that are not completely closed. Each larva lives in a chamber with an exit hole or tunnel that is blocked with chewed florets and debris, and the emerging fly must push through this in order to escape. The galls are hidden inside the protective bracts, and it is often not obvious from the outside that the flower head is galled. Tephritid galls in the flower heads range from simple accumulations of callus tissue to complex woody structures with distinct layers of specialised tissues, and examples of this range are included. Tephritid flies also cause woody galls in stems (see Chapter 8). Also included here because it causes a callus gall very like those of the less specialised tephritids, is the gall of a weevil (Rhinocyllus conicus, Coleoptera: Curculionidae) in thistle flower heads–beetles are otherwise out of place in this chapter (most of them cause stem and root galls and they are introduced in Chapter 8).

All fig wasps (Hymenoptera: Agaonidae) are associated with figs and their larvae live inside the fig ‘fruits’. Their galls are completely enclosed in the florets and are not apparent from outside the fig. The adults escape from their galls by chewing their way out through holes cut by their strong jaws. All fig wasps and their host fig trees are tropical and subtropical and none are found in Britain. The fig–fig wasp interaction is unique in insect galls in that both partners depend on the interaction and are benefited by it (rather than just the galler as in most gall interactions). The other well-known example of a galling organism benefiting its host is the Rhizobium–legume association (and Rhizobium is a bacterium; see Chapter 3, for this and a few other bacterial examples). (Mycorrhizal fungi on roots of plants also benefit their hosts and are considered to be gall causers; they are mentioned in Box 2.3, Chapter 2, but are not otherwise included in this book.)

BOX 7.1. Biology of tephritid flies (Information from Freidberg, in Ananthakrishnan, 1984; Freidberg, in Csóka et al., 1998; White, 1988; Headrick & Goeden, 1998; Norrbom et al., 1998; Korneyev et al., in Raman et al., 2005)

Tephritidae is one of the larger families of Diptera with about 4200 species worldwide, described in about 500 genera. Their classification is in a state of flux; Table 7.1 lists the currently accepted subfamilies and summarises the feeding habits of the larvae.

Although the diversity of the Tephritidae is greatest in the tropics, the Tephritinae, which includes all of the gall causers, is mainly subtropical and temperate in its distribution (though this may reflect the lack of knowledge of tropical Tephritinae). Compared to other Diptera, the family is fairly young in evolutionary terms, perhaps arising 25 million years ago in tropical Africa. Gall-causing is a relatively recent phenomenon in this family and arose several times in the Tephritinae. The complexity of galls varies greatly, from simple accumulations of callus to woody galls with a complex structure. The woody gall causers in Britain (Urophora, Myopites, Noeeta) belong to the less advanced ‘Lower Tephritinae’ while Tephritis, causing callus galls, belongs to the ‘Higher Tephritinae’ (see Korneyev et al., in Raman et al., 2005). But the North American Eurosta is also a higher tephritine and it causes a complex woody gall. So the more advanced tephritids do not necessarily cause the most complex galls.

TABLE 7.1. Feeding habits and examples of genera in the subfamilies of Tephritidae (after Freidberg, in Csóka et al., 1998; Headrick & Goeden, 1998; Ian White, pers. com., 2004)

Nearly all tephritids are herbivores, with larvae feeding inside fruits, stems, roots, leaves and flower heads. The larvae are white, yellowish or grey maggots with strong mouth hooks that slash through plant tissue, sucking up the cell contents and sap. Gall-causing larvae are shorter and wider than non-gallers and are less agile, moving little once the gall has developed. Adults are robust flies of moderate size (average wing length 2–8 mm, though some tropical species are larger) and feed on sugary liquids, e.g. nectar and honeydew.

Lifecycles of tephritids are simple: an egg, three larval instars (in Urophora, the first instar is retained inside the eggshell), a pupa that develops inside a puparium (the hardened larval skin), and an adult male or female. There are three lifecycle strategies in the family:

(1) Species with a broad host range, with larvae feeding in fleshy fruits and developing rapidly. They pupate in the soil, the adults survive for a relatively long time, they lay large numbers of eggs (800–3,000 per female), and often there are several generations a year (subfamily Trypetinae).

(2) Specialist fruit feeders, with larvae developing less rapidly, usually with only one or two generations a year. Adults live less long (one to two months) and females are less fecund (each lays 50–400 eggs). Otherwise they are similar to (1) (subfamily Trypetinae).

(3) Species feeding in stems, roots, leaves and flower heads, usually specialising in one or a few related host species. Larvae develop slowly and most species have one or two generations a year, usually pupating inside the host plant, with the fully fed larva or pupa overwintering. Adults are short-lived (less than one month, except for species that overwinter as adults) and fecundity is low (50–150 eggs per female). Includes the leaf-mining Trypetinae and all of the Tephritinae, including the gall causers and most of the British species.

BOX 7.2. Biology of fig wasps (Information from Weiblen, 2002; Kjellberg et al., in Raman et al., 2005)

Figs (Moraceae) are plants mainly of the tropics and subtropics with unusual ‘fruits’. These enclose large numbers (hundreds or thousands) of male and female florets–the fig fruit is an inside-out flower head or inflorescence (a syconium, from the Greek sykos = a fig; Fig. 104). Fig species have an unusual and complicated method of reproduction–pollination is dependent on the females of their particular fig wasp species, just one species or sometimes a few closely related species. Fig syconia are either monoecious or dioecious (strictly, ‘functionally dioecious’), i.e. the monoecious figs produce both pollen and seeds and the dioecious figs produce either pollen or seeds, even though all of the figs contain both male and female florets. Female fig wasps, which carry the pollen from fig to fig, can develop only in galled female florets, and seeds can be produced only by ungalled female florets. So, in dioecious figs, female wasps carrying pollen emerge from fig syconia that are functionally male and seeds mature in syconia without galls, which are functionally female (see the text).

Fig wasps belong to the family Agaonidae in the large superfamily Chalcidoidea (chalcids), a group of Hymenoptera that is mainly parasitic on other insects. Fig wasps are totally dependent as larvae on their figs, where they either feed on the seeds inside ovaries of female florets causing galls in the process, or are parasitoids on the larvae of other fig wasps in the flower head. Unusually for galls, the fig species also is dependent on this association. The lifecycles of both the figs and the fig wasps are so closely adapted and tightly synchronised that each is dependent on its own species. There are over 700 species of figs and probably more fig wasps (not all species are known). This plant–insect interaction provides one of the best examples known of ‘coevolved mutualism’, where each organism benefits the other and is dependent on it (mutualism) and each has adapted to the other over time (coevolution; see Chapter 11). It seems that this mutualism is ancient, having appeared just once in the Cretaceous period about 90 million years ago, when flowering plants were diversifying rapidly. Fig species have diversified with their specific fig wasps since then. In most, new species of both partners have arisen by cospeciation; in some, host switching may be involved, illustrated in Fig. 105. The only other examples of mutualism in galls involve bacteria in root nodules, e.g. Rhizobium and their host legumes (Fabaceae).

FIG 104. (a) Section through the syconium of a monoecious fig; (b) a single female floret. In (b), the stigma, style and swollen ovary are visible; the perianth (petals and sepals) is much reduced.

FIG 105. Possible methods of evolution of new species of figs and fig pollinators, (a) by cospeciation and (b) by host switching (after Weiblen, 2002). Letters A–C indicate fig wasp species, and numbers 1–3 indicate fig species. In (a), the ancestral pollinator fig wasp (A) splits into two descendant species (B and C), causing the ancestral host fig (1) to split into descendant fig species (2 and 3). Evidence suggests that the fig wasps evolve more rapidly than the figs and drive the speciation of the figs. In (b), an individual pollinator fig wasp (A) switches from its host fig (1) to a second fig species (2) that happens to be missing its normal pollinator fig wasp (B; indicated by its terminated line); in time, this errant individual founds a new fig wasp (C) able to pollinate fig (2). Fig (2) may evolve into a new species with fig wasp (C), or may support two fig wasp species (B and C), perhaps inhabiting different parts of its geographical range.

But not all fig wasps are so altruistic. Most are gall causers but some do not pollinate their figs (Table 7.2)–these are cheats. Theirs is a one-sided interaction similar to that of other gall causers, where the insect gains food and shelter and provides nothing in return. Agaonids belonging to the subfamily Sycoryctinae are parasitoids of the gall causers and may be more closely related to the chalcid families Pteromalidae and Torymidae than to other Agaonidae (this may apply to the gall-causing subfamilies Otitesellinae and Sycoecinae too).

The basic lifecycle of fig wasps is straightforward and either lasts a year or there are several generations a year. Eggs are laid by the female into gall florets, larvae feed in the young seed and cause the gall, pupae develop inside the galled ovary, and the males and females emerge with the sex ratio biased towards females. Like other chalcids and the gall wasps (Cynipidae), the larvae are legless grubs with well-developed jaws that destroy cells as they feed, either the nutritive tissue that develops in the gall or their host larvae (the parasitoids).

Tephritid flies are introduced here and background information on the biology and classification of the family is included in Box 7.1. The final section is a general description of fig wasps and their galls, and background to the biology and classification of the Agaonidae is given in Box 7.2.

TEPHRITID GALLS

Flies of the family Tephritidae are often known as fruit flies, picture-winged flies or gall flies, although none of these names is satisfactory. In Britain, Drosophilidae are known as fruit flies, not all tephritids have patterned wings (and picture-winged is used for other families too), and only about 5 per cent of species causes galls–so, tephritids is the common name used here. The family has specialised in the Asteraceae. The larvae feed in the flower heads or stems, and sometimes in the roots, leaf petioles or leaf veins. Flower head galls are characteristic in Britain and Europe although, worldwide, stem galls are commoner. The simplest galls consist of accumulations of callus tissue resulting from the wounding of cells, with the larvae feeding on the callus as well as on the normal plant tissues. Some tephritid larvae cause similar wounding but do not feed on the resulting callus; they are simple herbivores and are not classed as gall causers. Other species enhance their food supply without causing a gall. In North America, Paracantha gentilis feeds on the florets and achenes of native thistles and supplements this by sucking up sap from the receptacle cells: with its mouth hooks, it scores the surface of the receptacle to form cup-like depressions that fill up with sap. Thus, as in many other groups, straightforward herbivory merges indistinctly into gall formation. The more complex galls are woody swellings with several types of tissues, caused by Urophora and Myopites species in Europe and Eurosta and other genera in North America. Apart from Urophora cardui, which forms a woody stem gall, Urophora and Myopites gall the flower heads, while Eurosta and its relatives gall the stems. Examples of the flower head galls are described below, and the stem galls of U. cardui and Eurosta in Chapter 8. Wherever they feed and whether or not they cause galls, all tephritid species are specialists, feeding in one or only a few closely related host species.

Flower heads of Asteraceae provide a rich food supply for tephritid larvae whether or not galls are caused, and the gall causers clearly evolved from simple herbivores feeding on the developing florets, achenes (they contain the seeds) and the receptacle. In Europe, tephritids divide up this resource to thoroughly exploit it and this probably reduces competition between species (Table 7.3). Some species attack the flower heads early, before flowering, feeding on the young florets, developing achenes and receptacle, and often causing galls–in these, cells are able to multiply and enlarge, the vascular tissues can increase, and the gall and its larvae become a nutrient sink. Tephritis and Paroxyna species include non-gall causers and some that induce simple callus galls (see below). Paroxyna larvae are smaller than Tephritis but even a single larva cannot survive in the smallest heads, because food supply is insufficient. Most species of Urophora, Myopites and Noeeta cause complex woody galls, often with several larvae per gall (examples are given below). Their larvae vary in size too (Urophora > Myopites > Noeeta, decreasing in size) and do not gall the smallest category of heads. Tephritid galls in small-headed hosts tend to be in the stems and roots (see Chapter 8). Other species attack later after flowers have appeared, and are restricted to large flower heads with medium-sized and large achenes. They feed on the ripe achenes and receptacle, which no longer draw nutrients from elsewhere in the plant and cannot be induced to form a gall (but see Terellia tussilaginis, below, for an exception). The largest heads may support several larvae but smaller heads only one, and often in these the whole of the receptacle is eaten.

Stem- and root-feeding tephritids, both simple borers and gall causers, probably evolved from flower head feeders. Nearly all of their hosts are Asteraceae with small heads (e.g. Achillea, Solidago, Artemisia) and the larvae may have moved into the stem below the flower head to supplement their food supply. Stem galls are common, e.g. Eurosta solidaginis in goldenrod Solidago in North America and Urophora cardui in creeping thistle Cirsium arvense in Europe (see Chapter 8; this is the only Urophora species in Britain to cause a stem gall). Creeping thistle has the smallest flower heads of any Cirsium species and it is unusual in having separate male and female flower heads. Achenes do not mature in the male heads and so food supply is less there than in female heads. Two non-galling tephritids illustrate this difference. Xyphosia miliaria attacks early when florets and receptacle are growing and receiving nutrients from elsewhere in the plant, and it can develop in both male and female heads. Terellia ruficauda attacks late when heads are fully developed, and can survive only in the female heads. In both species, there is sufficient food in the head for only one larva.

Tephritid adults are sluggish flies that sit for lengthy periods on the food plant, which allows their behaviour to be watched. The males are territorial and mating occurs on the plant. Males usually arrive first and establish a territory by patrolling a number of flower heads or shoots of suitable size for the females to lay their eggs. They are aggressive to other males and to other flies of roughly the same size–there is much waving of wings and rocking of the body, with side-to-side jumps and sometimes collisions, and eventually one male departs. The successful male then attempts to mate with any female that arrives, sometimes of another species. Females are more choosy and are more active. They visit the territories of several males and mate several times during their lives. They do not tolerate males until they have fed for a while so that their eggs mature, and then they select a mate only after a period of courtship. This involves a routine of semaphore-like waving of the wings and the pattern of spots and stripes on the wings, characteristic of the different species, seems to be important in reducing hybrid mating. Copulation takes some time, with the female walking about carrying the male on her back. After mating, the female selects flower heads or buds of a particular size in which to lay her eggs.

The proportion of nutrients diverted from growth of the plant to the developing gall and larvae increases as the galls become more complex, and so woody galls tend to have a greater effect on their hosts than the callus galls. The simpler callus galls are illustrated by Tephritis and Terellia species in burdocks and thistles, and by the weevil Rhinocyllus conicus in large-headed Carduus and Cirsium thistle species, and the complex woody galls by Urophora and Myopites in thistles, knapweeds and fleabanes.

Female Tephritis lay eggs into young, unopened, flower heads that contain undeveloped florets and achenes. These are supported on a receptacle consisting of dense, closely packed cells, well supplied with vascular bundles. Lesser burdock Arctium minus provides an example. The size of the head and its stage of development are critical. Tephritis bardanae requires closed heads of burdock with a diameter of 6–16 mm. Smaller heads provide insufficient food for the larvae and, in larger heads, the ovipositor is not long enough to pierce the bracts; and the female is too large to squeeze between the spines in order to get closer to the bracts to lay eggs. Oviposition occurs in late May and early June when most flower heads are small but, even so, the number of suitable heads on a plant on any one day is never high. T. bardanae inserts a small clutch of eggs into the base of an inner bract (Fig. 106a). Several clutches may be found in a head, the result of laying by two or three females. After laying the eggs, the female smears a pheromone (a chemical that affects the behaviour of other individuals of the same species) across the head with her ovipositor. This deters other females from laying in the same head, and so reduces competition between the subsequent larvae–but the deterrent is not perfect as the chemical soon evaporates or washes off in rain. A maximum of only about seven larvae per head will survive to pupate. Tephritis formosa, laying eggs into the small, non-spiny heads of the perennial sow-thistle Sonchus arvensis, requires closed heads of only 3 mm diameter and inserts her eggs between the inner bracts and the outermost young florets. Sow-thistles produce a sticky white latex if damaged, which would gum up the delicate stylets of the ovipositor; the female avoids this by carefully insinuating the ovipositor between the bracts so that no tissues are pierced. After hatching, the first instar T. bardanae larva mines through the parenchyma tissue of the receptacle and enters a young achene (Fig. 106b). The achene enlarges as soon as the larva starts to feed. The larva grows rapidly, hollowing out its achene, and moulting twice (Fig. 106c). The larva then devours neighbouring soft achenes and receptacle tissue. The receptacle cells respond by multiplying to produce callus, which grows up between the damaged achenes and is supplied by enhanced vascular strands in the receptacle. The larva feeds on the callus and is fully fed by mid-August. The most rapid growth of the larva coincides with rapid growth of the achenes, augmented by the extra receptacle tissue. Time is short; the larva must be fully fed before the head starts to dry out. Then, the callus, damaged florets, frass and other debris harden to form a solid concretion around each larva, filling the whole head (if there are six or seven larvae). In mid-to late August, the skin of the larva darkens to become a black puparium, and the larva pupates inside, and the adults emerge during September and October. They overwinter in dense vegetation and leaf litter, surviving for about nine months. The females, at least, must feed during this time as their ovaries are not mature in autumn–foods available during winter and spring could be sugary sap and honeydew on leaf surfaces, liquefied bird droppings and cuckoo-spit (from froghoppers Philaenus spumarius). The T. bardanae gall is not obvious from the outside of the burdock flower head. In sow-thistles Sonchus and hawkbits Leontodon, the flower heads are smaller and less tough than in burdocks; they are galled by other Tephritis species and are distinctly swollen and sometimes ‘button-shaped’ (Fig. 107). The cells of the outer receptacle and the base of the bracts enlarge and multiply to accommodate the larvae inside.

FIG 106. Development of Tephritis bardanae in flower heads of lesser burdock Arctium minus (after Straw, 1986, quoted in Redfern & Askew, 1998). (a) Egg batch inserted into the base of a bract; (b) first instar larva entering a young achene; (c) young third instar larva inside an achene, having eaten its contents.

Rhinocyllus conicus (Curculionidae) is a weevil native to Europe and western Asia. It is included here because it causes simple callus galls very similar to those of Tephritis bardanae in burdock. R. conicus has an economic importance too, as part of biological control programmes for weedy Carduus and Cirsium thistles in North America and other parts of the world (see Chapter 13). The larvae feed in the flower heads of large-headed species, particularly of nodding thistle (Carduus nutans). Adults are large, robust weevils with a short, stout rostrum (Fig. 108a). In June, the females lay eggs on the outside of the thistle flower head, each one covered with a yellow-brown cap of chewed host plant tissue (Fig. 108b) that prevents the hatchling from drying out and falling off the head. If the cap is incomplete or is removed, the young larva cannot burrow into the head. The larva hatches in about a week and chews a tunnel through the bracts, feeding on the receptacle and young achenes (Fig. 108c) and sometimes burrowing into the flower stalk. It severs the vascular tissue and the burrow fills with callus tissue produced by the wounded cells of the receptacle. The larva constantly turns around in its burrow, consuming the callus as fast as it grows, and grows through July and August. When fully fed in its fourth instar, it constructs an ovoid chamber with walls made of chewed-up plant debris and faeces, and this hardens into a tough black cocoon (Fig. 108d). Inside, a pupa develops and the adult emerges about two weeks later. If there are many larvae in the head, they eat everything inside except the bracts and pappus hairs and the head becomes stuffed with black pupal cocoons. The adults may remain inside their cocoons for several weeks before emerging to overwinter in the ground. In April and May, the adults start feeding on the leaves of the thistle rosettes, and their eggs mature. They mate on the plant and females lay eggs as soon as the thistle heads appear. Unlike Tephritis bardanae and other tephritids, female R. conicus lay eggs on heads of a wide range of sizes, usually unopened but sometimes when in flower. Further south in Europe, there may be two generations a year, adults of the first emerging early enough to lay eggs in late July and August, and these produce more adults in September that overwinter. In Britain, the weevil is restricted to southern England and has only one generation a year.

FIG 107. Galled and ungalled flower heads of perennial sow-thistle Sonchus arvensis galled by Tephritis formosa (from Redfern et al., 2002 with permission).

FIG 108. Rhinocyllus conicus in flower heads of nodding thistle Carduus nutans. (a) The adult; (b) egg caps on the underside of a thistle head (scale = 10 mm); (c) the larva and (d) pupal cocoons inside the head (scale = 10 mm). (Images in (a) and (c) provided by Peter Harris; (b) and (d) after Redfern & Askew, 1998.)

The effect on the plant varies depending on the size of the thistle head and the number of larvae within it. If it is attacked when young and by several larvae, a head may fail to produce any mature achenes and be totally eaten out, with its base noticeably broadened compared to an unattacked head. But in a head with only one or two larvae which was more mature when attacked, some achenes will survive and the gall will not be obvious from the outside.

Returning to the tephritid flies and a second genus: most species of Terellia are not gall causers. They are late-attackers (Table 7.3), and feed on achenes and tunnel through the receptacle but do not induce callus tissue. They pupate in the head, often in a cocoon of silk and floret hairs constructed by the full-grown larva and, if several cocoons are present, the head may be slightly distorted. Terellia tussilaginis is unusual. It feeds inside burdock achenes causing them to enlarge slightly, becoming simple galls. The galled achenes do not create a strong nutrient sink; in burdock, the achenes are particularly large and provide sufficient food for the larvae. T. tussilaginis uses heads that are larger and further developed than those required by Tephritis bardanae and, usually, the two species do not compete for heads. Adults appear on the plants in mid-to late July after most T. bardanae adults have left, and males are less aggressive than T. bardanae males, with territories that are less distinct. Often, several males will try to mate with one female, six or seven stacked one on top of the other on her back! The females lay eggs into heads about 17 mm across, just before they open or when the tips of the purple florets are appearing. Each female lays a small clutch of eggs (Fig. 109a) between the florets or between the inner bracts and the achenes; the achenes are by now nearly full-sized and about 5.5 mm tall. In eight or nine days, the first instar hatches through a split in the side of the egg (Fig. 109b) and crawls to an achene, now full-sized but still soft. It bores into the achene leaving a small hole (Fig. 109d), and eats the seed embryo and then the endosperm (the food store of the seed). Occasionally, a second larva enters the achene and the older one kills it–there is not enough food for two. The larva grows rapidly and moults twice, and the achene becomes slightly more rotund than an ungalled achene. The third instar eats out its achene, then chews a hole near its base, pushes out its head and eats the contents of an adjacent achene–one does not provide enough food. When fully fed, the larva is yellowish white with a dark brown posterior plate (a feature characteristic of many tephritids). It retreats into its galled achene, plugs the hole with chewed-up floret hairs and overwinters, usually in the head that has fallen to the ground. Inside the galled achene next May or early June, the golden-brown puparium forms and, a month or so later, the adult pushes its way out of the hole. T. tussilaginis replaces the seed embryo of the achene, taking nutrients intended for its growth and slightly increasing its nutrient sink effect. It is thus a true gall causer, although a simple one, and no specialised nutritive tissue develops in its gall.

FIG 109. Development of Terellia tussilaginis in a flower head of lesser burdock Arctium minus (after Straw, 1986, quoted in Redfern & Askew, 1998). (a) Eggs; (b) hatching larva; (c) a normal achene entire compared with (d) a galled achene with a small entry hole (arrowed) and in section containing a young larva; (e) a galled achene with a large hole at the base chewed by the larva, and a section containing a full-grown larva.

FIG 110. Diagrammatic sections through an ungalled achene and through Urophora galls of increasing complexity in flower heads of black knapweed Centaurea nigra (a–d) and spear thistle Cirsium vulgare (e) (modified after Korneyev et al., in Raman et al., 2005). (a) A hypothetical non-galling achene feeder (no Urophora with this habit is known). (b) The achene gall of U. quadrifasciata with adjacent ungalled achenes. (c) The flask-shaped ovary–receptacle gall of U. affinis. (d) The multichambered cup-shaped achene–receptacle gall of U. jaceana. (e) The multichambered block-shaped achene–receptacle gall of Urophora stylata. (f) shows the stem gall of Urophora cardui in creeping thistle Cirsium arvense for comparison (see Chapter 8). (Nutritive tissue, black; woody tissue, finely stippled; callus tissue, coarsely stippled.)

Urophora is the largest genus in the tephritine tribe Myopitini (see Box 7.1) and most of its species cause the complex woody galls characteristic of thistle and knapweed flower heads. The best-known species are U. jaceana in black knapweed Centaurea nigra and U. stylata in spear thistle Cirsium vulgare, which gall both the achenes and the receptacle (also U. cardui in creeping thistle Cirsium arvense that galls the stem, see Chapter 8). Two other species cause less complex galls: U. quadrifasciata galls the achenes and U. affinis the receptacle of several knapweed species, including black knapweed (U. affinis is not a British species). They illustrate a series of increasingly complex galls, U. quadrifasciata > U. affinis > U. jaceana > U. stylata (Fig. 110), with the stem gall of U. cardui probably the most highly evolved. Unusually for the genus, U. quadrifasciata’s gall is not woody; it may have evolved from a more simple achene feeder similar to Terellia tussilaginis.

Urophora is a Palaearctic genus, with the gall causers native in Europe and western Asia. Most specialise on just one or a few related host species; the most specialised species, U. cardui, occurs only on creeping thistle. The least specialised, U. quadrifasciata, galls knapweeds in several subgenera and has also been recorded from saw-wort Serratula tinctoria in mainland Europe. It has the widest distribution, occurring as a native in northern Africa and western Asia as well as in Britain and Europe. U. jaceana, which galls two of the hosts attacked by U. quadrifasciata (Centaurea nigra and C. jacea), is native only in western, northwestern and central Europe. Both species and U. affinis have been introduced into Canada for the biological control of the knapweeds (see Chapter 13), accidentally for U. jaceana but intentionally for the other two, and their distribution now is much broader. U. quadrifasciata is opportunistic and active as an adult. In the nine years after it was introduced in 1972, it spread more than 400 km from its original release site in British Columbia and now is widespread in Canada and the USA. U. jaceana must have been introduced into eastern Canada before 1923, yet it is still restricted to Newfoundland and Nova Scotia even though its host black knapweed is widespread in Canada and northeast USA. Although Urophora adults seem to be sluggish flies reluctant to move far, they are perfectly capable of flying several kilometres a year. So other factors must restrict the distribution of U. jaceana in Canada. U. cardui has a restricted range in Europe where it is native (discussed further in Chapter 12).

Urophora adults, like other tephritids, are territorial with elaborate mating behaviour. The male usually reaches the food plant first, attracted by its architecture (cues are the size and arrangement of the shoots and flower heads, the pattern of bracts on the heads and the length and orientation of their spines). Like Tephritis bardanae on burdock, the male establishes a territory containing flower heads of a suitable size for oviposition and so attractive to females, and the courtship behaviour of the males and the choice of the females normally prevents hybrid mating. The specific wing patterns (Fig. 111) are prominently displayed during courtship and females will only accept the correct male. Apart from U. cardui which lays its eggs into leaf buds, Urophora females oviposit into unopened flower heads, selected according to their size. U. quadrifasciata is the least choosy–on black knapweed, heads of 5–10 mm diameter are suitable and are available for about three weeks in June and July. U. jaceana females, on the same plant, require younger heads of only 3–5 mm diameter, which are available for only about 15 days. Although the two species rarely gall the same head, they can survive together as long as the U. jaceana gall is not too large; the larvae do not compete directly for food and space. U. stylata causes a more solid gall than U. jaceana’s (compare Figs. 110d & e) in the larger heads of spear thistle Cirsium vulgare; only those that are unopened and 8–12 mm in diameter are suitable for oviposition. So, the ‘window of opportunity’ for oviposition is narrow. Lack of synchrony between female activity and development of flower heads is one of the main reasons that the number of galled heads at a site varies from year to year. Having chosen a flower head, the female inserts her ovipositor between the bracts and its tip examines the florets inside. If they are not at the correct stage of development, she withdraws it and investigates another head. In suitable heads, U. quadrifasciata places her eggs one at a time between the developing stamens or into individual florets. The heads suitable for U. affinis, U. jaceana and U. stylata are younger and the florets are smaller. U. affinis lays her eggs between the inner bracts or in a small clutch between the florets, and U. jaceana (Fig. 112) and U. stylata deposit theirs into the space between the florets and the overlapping bracts. U. stylata stands on top of the head while laying, while U. jaceana inserts her ovipositor from the side between the bases of the bracts, piercing some florets as it curves upwards to reach the space above the florets. The larger species have more eggs to lay than the smaller (60 eggs per female in U. quadrifasciata, 180 in U. jaceana) and often not all of the eggs are laid–perhaps because suitable heads are in short supply or cold, wet weather reduces the activity of the females.

FIG 111. Adults of Urophora species, showing their distinctive wing patterns: U. stylata, top right; U. stylata, top left; U. solstitialis, lower right; U. cardui, lower left (from Redfern, 1995, with permission).

First instar larvae of U. quadrifasciata hatch in three or four days and burrow down individual florets to the achenes. For the larva to survive, the young seed (ovule) must already be fertilised (so, by the time the eggs hatch, the head is flowering). Fertilisation means that the seed will be connected to the vascular system supplying the head, and this will ensure that the larva inside will have enough food to develop. Although the larva enhances the nutrient sink established by the developing seed, it cannot initiate a sink by itself. In U. affinis, U. jaceana and U. stylata, the larvae hatch before flowering and fertilisation occur, and are able to establish a nutrient sink themselves. In these species, it is the second instar larva that hatches and initiates the gall; the first instar is retained inside the eggshell.

FIG 112. Section of a black knapweed Centaurea nigra head containing a clutch of Urophora jaceana eggs (after Redfern & Askew, 1998).

The U. quadrifasciata gall is relatively simple and involves the achene and its nutrient supply only. Several achenes in a head will be galled but these do not fuse together unlike in the other Urophora species. As the larva burrows down its floret into the achene, the cells inside the seed multiply to form small patches of nutritive tissue lining the larval cavity. It feeds on these and burrows through the base of the achene to reach a vascular bundle, where the parenchyma cells within the bundle are its main food source. This gall does not become woody. After the egg hatches, the gall takes 15–20 days to become full-sized, by which time the larva has eaten out the whole achene except for its outer wall, reduced to a flimsy skin like tissue paper. U. quadrifasciata develops more rapidly than the other species and there are usually two generations a year. Larvae of the first generation pupate about three weeks after eggs were laid and adults emerge two weeks later; these lay eggs into late-developing heads of the same host or into other knapweed species that develop later. More galls form and the full-grown larvae overwinter in the remains of the galled achenes. They pupate in the spring, and adults emerge in May and June ready to gall next year’s flower heads.

The gall of Urophora affinis is woody but with one chamber only unlike those of U. jaceana and U. stylata, although there may be several separate galls in the flower head. U. affinis galls several knapweeds, though fewer species than U. quadrifasciata. In June and July it lays eggs and the hatchlings mine down through the achenes to feed on receptacle tissue. This stimulates a ring of receptacle cells to grow up around each larva, enclosing it almost completely in about ten days and leaving a hole at the top. The base of the gall links up with a vascular bundle in the receptacle, the inner cells of the larval chamber become typical nutritive tissue (described in Chapter 1), and the outer layer becomes hard and woody. Four weeks after the gall started to form, the larva has eaten all of the nutritive tissue and is full-grown, in a galled achene that is considerably larger than the neighbouring achenes. Most larvae overwinter in their galls, pupate in the spring and adults emerge in June and July. In some years, especially in the south of its range, a few adults emerge in late summer and there may be a partial second generation. Fig. 110 compares the gall of U. affinis with those of U. quadrifasciata, U. jaceana and U. stylata; U. jaceana and U. stylata are described next.

FIG 113. Spear thistle Cirsium vulgare head galled by Urophora stylata. (Tom Higginbottom)

The woody, many-chambered galls of U. jaceana and U. stylata are common in black knapweed and spear thistle, respectively. U. jaceana can also live in brown knapweed Centaurea jacea and a few other knapweeds, and U. stylata may occur in other large-headed thistles (e.g. nodding thistle Carduus nutans) especially in mainland Europe in areas where spear thistle is uncommon, but neither can survive in the other’s host plants. U. jaceana’s gall is cup-shaped and similar to that of U. affinis (Fig. 110) except that individual galls fuse into a woody mass. The U. stylata gall forms mainly from the receptacle and is more solid than U. jaceana’s gall, and is described as block-shaped. The galls of both species are often not apparent from the outside–but squeezing the flower head will reveal the hard lump inside. If the heads contain large galls, though, they are often characteristically distorted (Fig. 113). The lifecycles are similar. Eggs are laid in clutches in June and July, usually above the young florets (Fig. 112). They are elongated and curved and the first instar moults inside–it is the second instar that hatches, a week or two later. The larva mines down a floret to the achene which, with the surrounding receptacle tissue, swells to enclose it. Usually several larvae burrow down separate florets simultaneously and gall tissue forms round each. During July and early August, the gall is soft and fleshy with the main patch of nutritive tissue at the base of each chamber. A thin woody layer develops outside this and is incomplete at the base at first. This enables nutrients to pass from vascular bundles in the receptacle to the larva, which is growing rapidly at this time. By mid-August, the individual galls have coalesced into a single large multiple gall with several chambers, and the larva is in its third instar. By September, the gall is hard and woody and contains fully fed larvae (Fig. 114). When full-grown, the larva has a tough, dark-brown posterior plate that blocks the entrance to its chamber and may help to prevent parasitism–the larva always orientates itself head downwards. In autumn or winter, the galls usually fall to the ground. The larvae inside are fat little barrels, rich in calories, and mice and voles hungry during the winter eat many of them. In spring, the larvae reverse their position in the chamber and pupate, and the adults emerge in June and July. The adult U. stylata must push out through the spongy callus that blocks the exit tunnel at the top of the chamber. This callus rots in spring while the galls are on the ground.

FIG 114. Section through the gall of Urophora stylata in the head of spear thistle Cirsium vulgare with a mature multiple gall; it contains a fully fed third instar larva in the left-hand chamber (the other chambers contain parasitoids and caterpillar frass, see Chapter 10) (after Redfern, 1995).

Urophora galls in knapweed and thistle heads illustrate several galling strategies (and the stem gall of U. cardui adds another). Although U. quadrifasciata and U. jaceana exploit the same host plant (Centaurea nigra) and so both species are presented with the same array of resources, their strategies are very different. In most respects, the strategy of U. jaceana is the more specialised of the two. It requires the females to lay whole clutches of eggs in flower heads in just the right state of development, with a premium on those in the most vigorous plants. The benefits of this strategy are:

A strong nutrient sink that provides food for the insect supplied from the plant generally and not just from the galled flower head. The more larvae there are in a gall, the stronger the sink, so that larvae from large galls are much the same size as from small galls.

The large woody gall gives greater protection against inclement weather and provides a refuge from parasitism for larvae in the central chambers (the length of the parasitoids’ ovipositors imposes limits–this is discussed further in Chapter 10).

This strategy is risky if there is any mismatch between development of the flies and their host plants and, for an individual female, many of her progeny may be lost in a single event–‘all her eggs in one basket’–they will all be destroyed if the flower head dies. Development is relatively slow but the resulting larvae are large and well protected.

By contrast, U. quadrifasciata has a more scatter-gun approach. Individual larvae are not so well protected, but the progeny of any female are scattered over more flower heads and plants. U. quadrifasciata is capable of infesting a wider range of host plants than U. jaceana and can infest flower heads over a longer time, and in some areas it can have two generations a year.

The different strategies affect the host plants. U. quadrifasciata destroys the achene it inhabits and causes a few neighbouring ones to abort, and so reduces the number of viable seeds per flower head. Otherwise, it has little effect on the plant. U. affinis causes more florets and their achenes in the galled head to abort, and also causes neighbouring heads on the same plant to be less productive or to fail to develop. The multichambered galls of U. jaceana, and also of U. stylata in thistles (and U. cardui in stems), have the greatest effect, which increases as the size of the gall increases. Besides destroying the individual achene that each larva attacks, all achenes adjacent to the gall fail to produce seeds and a large gall may prevent the head from producing any viable seeds. Nutrients are drawn from neighbouring heads also and their seed production may be reduced to zero too, and a heavy infestation on a small plant may result in no viable seeds at all. But this rarely happens. There are many other factors that also affect the success of the gall flies, reducing their impact on the plant: failure of adults to find suitable host plants and heads of the right size for oviposition, bad weather when they are active, and mortality of adults, larvae and pupae due to predators, parasitoids, etc. Interactions like these combine to reduce the impact of the gall causer on the plant most of the time, so that it is unlikely to destroy its food supply (these aspects are discussed further in Chapter 10).

Myopites species gall plants in the tribe Inuleae of the Asteraceae, e.g. common fleabane (Pulicaria dysenterica) and golden-samphire Inula crithmoides. Their galls in the flower heads are hard lumps formed from lignified achenes and surrounding receptacle, but are less complex than the woody Urophora galls. Two species occur in Britain, M. inulaedyssentericae on common fleabane and M. eximia on golden-samphire (Fig. 115). Neither is common, and both are restricted to coastal sites in southern England. Their lifecycles are similar and broadly the same as the lifecycles of Urophora jaceana and U. stylata, though they run later in the year. There is usually one generation a year, though occasionally a few individuals require two years to complete their development. They overwinter as fully fed larvae and pupate in the gall the following summer, and the adults emerge in August.

FIG 115. Galled heads of common fleabane Pulicaria dysenterica (left with a section) and golden-samphire Inula crithmoides (right with an ungalled receptacle) caused by Myopites inulaedyssentericae and M. eximia respectively (from Redfern et al., 2002, with permission).

The lifecycles of Myopites and Urophora differ in detail. Myopites females need to feed to mature their eggs and their mouthparts are distinctive, extended into a long proboscis that probes summer flowers for nectar. They lay eggs into heads at the start of flowering, in older heads than those required by the Urophora species–in common fleabane at this time, the outer ring of strap-shaped florets is well developed, while the central tubular florets are still immature. They lay eggs into or between the tubular florets, usually near the centre of the head. The first instar hatches in about five days and mines down a floret and through its achene to reach the vascular bundle beneath. By this time the achenes are fertilised and are developing rapidly so that there is already a good vascular network established in the receptacle. The larva makes use of this and enhances it, in a similar way to Urophora quadrifasciata. Two weeks later, the larva moults into the second instar and tunnels vertically into the receptacle, now enlarged and domed, pushing the central florets upwards. The larvae feed on vascular bundles that end on the tunnel walls and on patches of nutritive tissue that line the tunnel walls. The activity of the larva keeps the upper part of the tunnel open–it strengthens the walls by gluing remains of damaged cells on to the sides of the tunnel and this material dries into a hard cement. Later, the adult will use the tunnel as an escape route. Five weeks after oviposition, the M. inulaedyssentericae larvae moult into the third instar and their tunnels extend laterally. They are making for the large vascular bundles that form an outer ring in the receptacle, that supply the bracts and the outer strap-shaped florets and patches of nutritive tissue continue to form. By mid-October, a layer of woody sclerenchyma tissue surrounds each larva’s chamber, penetrated by vascular bundles, and the larvae are fully fed. Although a flower head containing several larvae will feel hard and woody, the chambers tend to remain more or less separate. The larva seals the upper part of its tunnel with several translucent membranes, overwinters and the adult emerges next summer, breaking the membranes as it does so.

Myopites galls are similar to Urophora galls in several ways. The younger larvae exploit nutrients intended for seed growth, first as achene miners, like U. quadrifasciata, and then as receptacle miners. Later, they behave more like U. jaceana and U. stylata by feeding on special nutritive tissue in a gall that becomes woody. The mining stage, regarded as a primitive feature, is relatively long in Myopites, lasting well into the third instar. The flower head is affected less dramatically, too. Each Myopites gall destroys its own achene and a few adjacent ones but, even if the head contains several larvae, most heads produce some healthy achenes.

Woody flower head galls caused by tephritids are characteristic of the Asteraceae in the Palaearctic. In other regions of the world especially in North and South America, although there are many flower head tephritids and structurally similar Asteraceae, very few have yet been found that cause woody galls. An exception is Stamnophora vernoniicola in Africa, a myopitine tephritid that causes a multiple woody gall in the flower heads of Vernonia (a genus of Asteraceae). Stamnophora flies are similar to Myopites but without the elongated proboscis. The gall of S. vernoniicola can be large, up to 3 cm across, and contains many separate chambers (sometimes 50 or 100) that open into a common chamber near the top of the flower head. The larvae live in these chambers and pupate there, and the adults emerge through the common opening. S. vernoniicola also causes stem galls on other Vernonia species, similar to Urophora cardui galls but with chambers opening into a common chamber, as in the flower head galls. Apart from this, little is known of the lifecycle of the gall fly or of the development of the gall.

Urophora, and other tephritids that cause woody galls in flower heads, belong to the ‘Lower Tephritinae’ (see Box 7.1). Gall-causing ‘Higher Tephritinae’ in Britain are less common and most of their galls are simple accumulations of callus. Examples are Tephritis bardanae in burdock and species of Dithryca, Sphenella, Oxyna, Paroxyna and Campiglossa in various Asteraceae, in stems and roots or in flower heads. Eurosta (see Chapter 8) is a North American higher tephritine that does cause a woody gall but none of its relatives are British.

FIG WASPS AND THEIR GALLS

Most fig wasps cause galls and all are associated with fig trees, living inside the fig ‘fruits’ (inflorescences). Their lifecycles and habits, and the type of gall produced, are similar in most species. Most of them cause galls in the ovaries and are essential to the survival of the fig–the fig wasps are the only means by which the flowers enclosed inside the figs can be pollinated. The classic fig–fig wasp interaction is characteristic of the subfamily Agaoninae (Box 7.2, Table 7.2) and most of the following account refers to this group. Some fig wasp species in other subfamilies are normal gall causers that exploit their hosts without pollinating the flowers, or are parasitoids on other fig insects. At present these are classified in the Agaonidae but some of them may turn out to be more closely related to other chalcid families and eventually be removed from the Agaonidae.

Fig wasps are tiny, odd-shaped insects with males and females that look very different. Their shape is adapted to their way of life in the fig. The female has a long, flat head and jaws with appendages armed with teeth; these help her to push through the bracts that line the ostiole of the fig (Fig. 104 in Box 7.2). Sometimes, the antennae are also modified to help her to enter the fig. She often loses her wings as she enters and the ends of the antennae break off. Once inside, most females never get out again and, after laying her eggs, they will die inside–the fig is a ‘tomb blossom’ for females though a nursery for the larvae. The female brings pollen with her collected from the fig inflorescence she developed in and carried in special structures: pockets on the underside of the thorax, or pollen baskets on the front legs, or deep grooves between the abdominal segments. Females with these adaptations are ‘active’ pollinators; as they crawl over the florets, they remove pollen from the pockets and carefully place it on to each stigma. ‘Passive’ pollinators carry pollen scattered generally over the abdomen and they pollinate the florets incidentally as they crawl amongst them. While pollinating the florets, the females lay their eggs. The ovipositor is long and sharp and pierces the styles of particular florets, placing the eggs one at a time, precisely next to the ovule (which would develop into a seed if not attacked; Fig. 116). The eggs do not hatch if laid in the wrong place.

FIG 116. Section through a female fig floret containing a fig wasp egg (modified after Meyer, 1987). The arrow indicates the point of insertion of the ovipositor on the stigma and the dashed line the path of the ovipositor down the style; the egg (black) is laid on the surface of the ovule (stippled).

The egg is laid with a drop of liquid, which stimulates the fertilised ovule to enlarge, initiating the gall. Fertilisation of the ovule encourages development of the endosperm (the food store of the ovule), which becomes the nutritive tissue of the gall on which the larva will feed. So, besides pollinating fig florets that will produce seeds, female gall wasps (though not of all species) also pollinate the gall florets, so ensuring that their offspring have enough food to develop. Once the hatchling starts feeding, it destroys the seed embryo, and then starts on the nutritive cells, which draw nutrients from the vascular bundles that would have supplied the seed. The ovary enlarges as the larva grows, so that galled florets are more swollen than florets containing seeds. Pupation occurs in the galls, and the males emerge–before the females. The males are strange-looking insects; usually they have no wings, no eyes and no ‘feet’ (the tarsal segments of the legs are reduced), and the antennae are small and fold away into grooves on the head. They chew their way out of their galls leaving a hole behind and, still inside the fig, search for galled florets containing females. A male chews into one and inserts his abdomen so that the genitalia at its tip are in the correct position to mate with her before she emerges–his abdomen is flexible and curved and the genitalia telescopic to allow this to happen. He will mate with many females and the females, who remain in their galls for a while, may be mated by several males. The final job for the male (of many species) is to burrow out of the fig, excavating a tunnel through the receptacle with his forelegs; these are large and strong and armed with teeth. This tunnel allows the mated females to escape from the fig, laden with pollen collected from ungalled florets in the head. In some species, several males work together to cut a large exit hole for the females; the males climb out and form a circle around the hole, and guard the females from predatory ants. Females of some species use the fig ostiole to escape. They are strong fliers and disperse widely, searching for a tree of the correct species and with figs at the right stage of development to receive eggs. The females are attracted to the figs by their smell, which is unique to each fig species.

To ensure the survival of both figs and fig wasps, the wasps must not gall all of the female florets–otherwise there would be no seeds and no future fig trees, and both species would die out. There are several strategies that ensure that some florets escape. In some species, some florets are not available to the fig wasp and will produce healthy seeds if pollinated. Egg-laying is restricted by the length of the ovipositor and is successful only in florets with short styles (see Fig. 104b in Box 7.2). So, long-styled florets are left to produce viable seeds, and these ensure the future of the fig plant. In monoecious figs, short-and long-styled florets occur in the same fig inflorescence, which therefore produces both fig wasp females carrying pollen and seeds (and so is functionally both male and female). In dioecious figs, short-styled and long-styled florets occur in different fig inflorescences on different trees. So, figs with short-styled florets are galled and produce females that carry pollen to other figs (and so are functionally male), and figs with long-styled florets, if pollinated, produce seeds (and are functionally female). Females emerging from their figs and loaded with pollen are attracted to both types of figs, which have the same attractive smell, ensuring that the seed florets get pollinated. The wasps enter both types of fig, pollinate the florets and attempt to lay eggs in them, but are successful only in the short-styled figs–and these subsequently produce the next generation of fig wasps. (The story is oversimplified; if short-styled florets are in short supply, perhaps because several females are ovipositing at the same time, the females will lay eggs in long-styled florets, though their success rate there is reduced.) Other fig species rely on another strategy: the fig wasp’s ovipositor is long enough to gall both long-and short-styled florets, but she cannot lay eggs in all of them so some escape–she is limited by shortage of eggs or by shortage of time in which to lay them. In some fig wasp species there are too few females in the fig wasp population to exploit all of the female florets in the time the figs are available; the fig restricts the number of females entering by closing its ostiole. So, one way or another, fig trees produce enough seeds to ensure the future of the fig–fig wasp mutualism.

Apart from the Agaoninae, the Agaonidae includes non-pollinating gall-causing fig wasps and parasitoids (Box 7.2) that contribute to a rich community of insects in the figs (see also Chapter 10). The female gall causers either enter the fig through the ostiole, like the pollinators, or sit on the outside of the fig drilling through the receptacle–their ovipositors are long enough to reach some of the florets inside. The parasitoids (subfamily Sycoryctinae) have extraordinarily long ovipositors that reach far into the fig from the outside. They attack pollinating and non-pollinating gall causers as well as other parasitoids.

Finally, an observation on dessert figs, cultivated for the table. Since at least the start of the seventeenth century, it has been understood that for figs to ripen and to be worth eating, they must undergo ‘caprification’ (the puncturing of fig flowers by fig insects, ensuring their pollination; from the Latin caprificus = a wild fig tree). Caprification used to be encouraged artificially. The fig growers would hang strings of young, wild, springtime figs in the branches of their cultivated fig trees so that, when the females loaded with pollen emerged, they would enter the cultivated figs and pollinate the florets, so allowing them to mature. Nowadays, cultivated figs are of self-pollinating varieties so that caprification is unnecessary. This also means that consuming a fig is unlikely to involve a mouthful of fig wasps!

CHAPTER 8

Galls in Stems and Roots

STEM AND ROOT GALLS, closed from the outside world, are caused by several groups of insects and a few nematodes. Galls in leaf petioles, which have a structure similar to stems, are also included in this chapter, and some of these may extend into the midrib of the leaf. Beetle and moth galls are the simplest, with no nutritive tissue (see Chapter 1), and are described first. Sawfly galls in stems are more organised and have a more constant size and shape, though they are also without true nutritive tissue; they come next. Galls of other groups contain specialised nutritive cells in patches or in definite layers and the most complex include other layers of specialised tissues too. Two nematode galls are described and a gall caused jointly by a nematode and a gall fly, followed by stem galls of gall midges and tephritids. The chapter ends with a short section on stem galls of gall wasps, i.e. species that only gall the stems. Some gall wasps cause galls in two plant organs, e.g. a bud and a root, or a leaf and a stem, and these are included in Chapter 9. Beetles, moths and nematodes have not been encountered in earlier chapters and are introduced here. Sawflies and gall midges are introduced in Chapter 4 and Boxes 4.7 and 4.8, and tephritids in Chapter 7 and Box 7.1; for gall wasps, see Chapter 9 and Box 9.1.

BEETLE GALLS (COLEOPTERA)

Although the Coleoptera is the largest and most diverse order of insects, few beetles cause galls. Most that do are weevils in the two families Curculionidae and Apionidae, both in the large superfamily Curculionoidea, one of the most advanced beetle groups. Galling is very unusual in other beetle families–the longhorn beetle Saperda populnea (Cerambycidae) and the buprestid Sphenoptera jugoslavica (Buprestidae; not a British species) are examples. Beetle galls are simple. Most are accumulations of callus tissue caused by the larvae wounding plant cells. In flower heads, this results in callus galls similar to the simpler tephritid galls (see Rhinocyllus conicus, a curculionid weevil, in Chapter 7). Beetle galls are commoner in roots and stems where they increase the number and size of pith and parenchyma cells, so causing a swelling. Sometimes there is also an increase in vascular tissue but never any real formation of nutritive tissue, although some Ceuthorhynchus and Apion species cause a slight differentiation of tissues. All galling beetles have close relatives that are not gallers; they are simple borers and miners in stems and roots. Particular weevils often favour particular plants. In Britain, Larinus and Rhinocyllus (Curculionidae) specialise in thistle flower heads (Carduus, Cirsium; Asteraceae), and Gymnetron and Rhinusa (Curculionidae) occur in seed heads of speedwells Veronica and toadflax Linaria (both Scrophulariaceae). Apion weevils (Apionidae), now split into several genera, cause stem and root swellings on legumes (Fabaceae) and docks Rumex (Polygonaceae), and Ceutorhynchus (Curculionidae) has specialised on crucifers (Brassicaceae). Most galling beetles and their galls are not well known and most species that have been investigated are those with some economic importance. Rhinocyllus conicus and Sphenoptera jugoslavica are examples, causing galls in thistles and knapweeds respectively, and are used to help in the biological control of these weeds (discussed further in Chapter 13).

Weevil adults and larvae are usually easy to recognise. The adult beetles have a long snout (rostrum) bearing chewing mouthparts at its tip, the antennae are often elbowed and the body is clothed with hairs or scales. Gall-causing larvae are inactive grubs, white or yellowish in colour, without legs, and the body is characteristically curved so that the head almost touches the tail. The head is well developed and usually dark brown and has strong chewing jaws. Both adults and larvae are plant feeders, and often feed on the same host individual. The adults chew holes in the leaves and stems and the larvae burrow inside, the gall developing around them. Gall weevil lifecycles are straightforward and are usually annual: eggs laid on the food plant, several larval instars, pupae sometimes in a cocoon and usually in the plant, and adult males or females. Two galling beetles are described here, one a buprestid and one a weevil. Sphenoptera jugoslavica (Buprestidae) causes a simple root swelling in a knapweed and the weevil Eutrichapion scutellare (Apionidae) is responsible for a slightly more complex gall in stems of gorse.

Sphenoptera jugoslavica is native in eastern Europe and western Asia where it feeds on diffuse knapweed Centaurea diffusa and one or two closely related knapweeds. Neither the beetle nor the host plant is found in Britain or western Europe. There is one generation a year. The adult females emerge from their galls over an extended period during the summer and feed for a while on the new leaves of knapweed rosettes until their eggs mature (rosettes are the flat circles of leaves produced by knapweeds in their first year). Then they mate and between late May and early September they lay small groups of eggs at the base of the petioles of rosette leaves. The eggs must be wedged firmly between the petioles so that they do not dry out; they are not inserted into the plant. The larva hatches in two weeks and gnaws the surface of the petiole, then moults once and mines into the petiole, using the base of an adjacent petiole to push against, and this prevents it from falling off the plant. It continues through the crown of the rosette and downwards into the taproot, and the gall starts to grow around it. A gall can develop any time between July and September and, usually, only one larva per root will survive. At first the larva is a simple miner, feeding on the cortex of the root. When it reaches the vascular tissue in the centre of the root, its mine starts to fill with callus tissue and it feeds on this (Fig. 117). When the larva is half grown, it burrows deep into the root and overwinters. In spring, it returns to the gall and resumes feeding until May or June when it is full-grown. Then it tunnels upwards and makes a cocoon out of compressed frass at the top of the root, and pupates inside. The adult emerges in mid-or late summer. Often the new adult remains inside the cocoon for a week or two, before chewing a hole in it and tunnelling out through the root crown, to feed, mate, and to start the next generation.

FIG 117. Sphenoptera jugloslavica adult (left) and section through the root gall of S. jugoslavica with larva (right) on diffuse knapweed Centaurea diffusa. Note the characteristic shape of the buprestid larva, flattened, with broad thoracic segments and a small dark head. (Peter Harris)

FIG 118. Stem gall of Eutrichapion scutellare on gorse Ulex europaeus (Pauline Ivimey-Cook).

The effect of the gall on the plant can be severe. Because it develops in the main taproot, the gall slows down the growth of the rosette or prevents its growth altogether. Knapweed rosettes must reach a minimum size before they can produce a flowering shoot, and so the root gall can delay or prevent flowering and seed production. In large-galled rosettes, the plant will develop flowers and seeds, but produce fewer than normal. If galled, small rosettes may never flower. The plant has been accidentally introduced into British Columbia, Canada, where it is a serious weed of dry grasslands, and the beetle has been introduced in attempts to control it, with some success (see Chapter 13).

Eutrichapion scutellare causes an ovoid swelling in stems of Ulex, on dwarf furze U. minor as well as on gorse (Fig. 118). The swelling, up to 10 x 6 mm, is usually found in young stems. The gall has a smooth surface, ironing out the grooves of the normal stem, and inside is a thick-walled chamber containing one weevil larva. In the ungalled stem, the vascular tissue forms a complete ring enclosing the pith in the centre. In the gall, the larval chamber is in the centre and is surrounded by a layer of parenchyma cells that the larva feeds on, stimulated to multiply by continual wounding as the larva feeds. Interspersed with these cells are collapsed scar cells that are also the result of wounding. Enclosing the larva and parenchyma layer is a thin protective ring of scar tissue also formed from wounded cells. The vascular tissue outside this is enlarged and forms an incomplete and irregular ring, with wide channels of parenchyma tissue between the groups of vascular bundles. Details of the lifecycle of this weevil are not known.

MOTH GALLS (LEPIDOPTERA)

Few Lepidoptera cause galls and those that do belong to several unrelated families (Miller, in Raman et al., 2005, records 179 species worldwide in 20 families, estimated as less than 0.1 per cent of all Lepidoptera). All are ‘micromoths’ (e.g. Tortricidae, Momphidae, Nepticulidae). The usual plant parts attacked are stems and leaf petioles and, like beetle galls, the galls are simple accumulations of callus tissue. Many galling species have relatives that are stem borers and leaf miners, and the galls are slightly more complex versions of their borings and mines. In Britain, most moth galls are found on trees and shrubs, e.g. oaks, willows, poplars, Scots pine and larch. Apart from Mompha, with three species on willowherbs (Epilobium, Chamerion), most genera contain only one or two gall-causing species and these attack unrelated host plants.

Gall-causing moth larvae are typical caterpillars with an elongate body and prolegs on the abdomen, and a distinct head bearing strong chewing jaws. They are more active than most larvae that live inside plants and move freely inside their galls. Frass often fills the chamber or is pushed out of a hole cut in preparation for the adult to emerge, and silk spun by the caterpillar inside the gall is characteristic. The larva may pupate in the gall, sometimes spinning a cocoon first, or leave when fully grown to pupate in the ground. The emerging moth, unlike beetle adults, cannot bite its way out of the gall. Often the larva prepares an exit hole, spinning silk across it or leaving a thin window of bark. The pupa pushes out through this, breaking through using sharp horns on its thorax, and the pupal skin is left sticking out after the adult has emerged.

Two British species are described briefly although, because so little is known about most moth galls, these may not be characteristic of moth galls in general. Heliozela sericiella (Heliozelidae) galls leaf petioles of oak (Quercus robur and Q. petraea, and other species abroad). The female has a piercing ovipositor and inserts an egg into the petiole near the base of the midrib. Like sawflies (e.g. Pontania species, see Chapter 9, Sawfly Leaf and Bud Galls), the female injects a substance that causes parenchyma cells of the petiole to multiply and enlarge, and this swelling occurs whether or not the egg survives. The H. sericiella swelling is not very distinct. The larva hatches and feeds on this callus tissue in the swelling until it is in its fourth instar and nearly full-grown. It then becomes a leaf miner, tunnelling out of the gall and into the blade near the base of the leaf, and an ovoid blister develops around it. This is not part of its gall but is a swelling caused by the silken cocoon that the larva has spun inside. The larva cuts out the blister, which becomes its case; the case and larva drop to the ground leaving an oval hole at the base of the leaf blade, and the larva pupates inside the case and overwinters, emerging as an adult next spring.

The gall of the pine-resin gall moth, Retinia resinella (Tortricidae), is quite common on twigs of Scots pine (Pinus sylvestris; Fig. 119). Although the twig is slightly swollen, it is the characteristic mass of resin 1–3 cm across that draws attention to the gall. The moth has a two-year lifecycle. Eggs are laid in June, singly at the base of buds. In three weeks, the larva hatches and burrows inside as far as the pith of the stem, causing it to ooze resin. The cambium of the vascular tissue divides actively, producing additional phloem and xylem tissue that cause the swelling on one side of the twig, and the resin encloses the larva on its other side. The larva feeds on phloem and parenchyma cells, producing a groove along the inside of the twig. This is, therefore, a true gall, the larva causing and feeding on enhanced plant tissue, even though most of the ‘gall’ in Fig. 119 is a mass of hardened resin. By autumn, the gall is the size of a pea and the larva overwinters inside. It continues to feed next spring and summer and the resin mass increases to 2–3 cm across. The gall now consists of two chambers, the resin of the second year enclosing the first year gall. By its second autumn, the larva is fully fed. It overwinters in a silken cocoon inside the gall, pupates in the spring and the adult emerges in June.

FIG 119. The resin-gall of Retinia resinella on Scots pine Pinus sylvestris, entire (left) and in section (right) containing a caterpillar (from Redfern et al., 2002, with permission).

SAWFLY STEM GALLS

Several sawflies cause galls in stems, all belonging to the genus Euura and all specialising on willows. Euura is related to the Phyllocolpa, Pontania and Eupontania sawflies, in the same tribe of the subfamily Nematinae (Tenthredinidae; see Chapter 4 and Box 4.7, and Chapter 9). The galls are simple swellings usually in young stems or in the midrib or petiole of leaves, occasionally in buds, and contain one or several larvae in separate chambers. The swellings are simple accumulations of callus in which the normal plant tissue can sometimes still be recognised. There are no specialised nutritive or woody layers. Two species are described here: Euura lasiolepis, a North American sawfly that galls arroyo willow Salix lasiolepis, and the European and British E. amerinae, which specialises on bay willow S. pentandra.

Both species have one generation a year and their lifecycles are similar. The adults emerge from the old galls in late May or early June and live for about a week. They mate on the willow plants and the females lay eggs into rapidly growing new shoots, into the site where the larvae will feed. They prefer vigorous shoots of young plants. Arroyo willow has a shrubby growth habit and forms large clones with numerous vegetative shoots developing each year from underground stems, and can be galled year after year as long as the plant lives. Bay willow, on the other hand, is tree-sized and is suitable for galling only in its first 8–12 years, when it is young and is growing rapidly. If it is regularly pruned or coppiced so that its juvenile life is extended, it can be galled for a longer period. The female E. lasiolepis has an ovipositor that is a long narrow blade, and pierces through the petiole of very young leaves, depositing the egg deep into the young shoot. In E. amerinae, the female pierces the stem several times laying an egg each time, so that a series of swellings develop along the stem or a multichambered gall develops if the swellings merge. The eggs are bathed in a fluid produced by the female, and this fluid causes a swelling before the larvae have hatched. In most Euura, the initial swelling enlarges into an almost full-sized gall, a solid mass of callus cells, before the egg hatches. The larva hatches in 25–30 days and tunnels through these cells, wounding them as it feeds, and sometimes causing more callus to develop and the gall to enlarge. The larvae grow through the summer, passing through five or six instars (in the male and female, respectively). They are full-grown in the autumn and the gall becomes hard and woody. The Euura lasiolepis larva cuts an emergence burrow leaving a thin window at the surface of the gall (perhaps the E. amerinae larva does this also). It then retreats back to its chamber, spins a cocoon and overwinters inside, pupating there in the spring. The adult just has to cut through this window in order to escape. In contrast, most Phyllocolpa, Pontania and Eupontania species leave their galls as larvae to overwinter and pupate in the soil.

The galls of both E. lasiolepis and E. amerinae vary considerably in size depending on the vigour of the host plant and, in E. amerinae, on the number of larvae in the gall. If E. lasiolepis is unusually common, its galls may run together to form irregular swellings. Large galls create a stronger drain on the plant’s resources than small galls, especially if they are numerous. The stem beyond the gall often dies, which encourages lateral shoots to develop in future years, and results in a bushier plant. This, in turn, enhances the supply of young shoots and so benefits future generations of the sawflies. Large numbers of galls reduce the number of leaves per shoot, too, and may reduce or prevent flowering and seed production. Large galls protect their sawfly larvae better than small ones, at least against attack by parasitoids–the ovipositors of many parasitic wasps are not long enough to reach larvae deep inside large galls (see Chapter 10 for more discussion of this topic).

NEMATODE GALLS

Few nematodes cause galls–malformation and rotting of the host plants are more common effects of these plant parasites. A brief background to the worms is provided in Box 8.1. Their biology is not well known, apart from those that are pests, and some species, including a few gall causers, are important crop pests. Most nematode galls are swellings in roots and stems; a few affect leaves and flowers (especially the florets of grasses). Their structure varies from simple growths that vary considerably in size, to more constant, complex swellings. The simplest have an irregular cavity that only develops in the mature gall; in the more complex, well-defined layers of tissues surround a distinct chamber. All nematode galls develop nutritive tissue, usually in or near the vascular tissue, as long as the worms are actively feeding–single enlarged cells or distinct patches or layers of cells forming a ‘syncytium’ (cells whose walls break down so that their contents coalesce). The cells and worms act as a nutrient sink, as in the galls of mites and insects. Nematode galls are included in this chapter because they usually develop cavities and have no opening to the outside. The worms are usually released into the soil only when the gall and its host plant rot. The Fergusonina–Fergusobia galls are an exception, see type (5), below. Inevitably there is some overlap with other chapters: some of the simplest swellings are more or less solid until the end of their life (and overlap with Chapter 3) and, in one case, in a relationship of the nematode and a bacterium that is not fully understood, a virescence or fasciation of the shoot of the host develops (relevant to Chapter 2).

There are four main types of nematode gall, plus a fifth in which nematodes are involved in a remarkable association with flies and Eucalyptus:

(1) Simple swellings on plant roots that vary from pinhead to grape in size and may occur in clusters. They may be solid but usually develop large air spaces that form irregular cavities, and these are the main cause of the swelling. The nutritive tissue takes the form of one to many giant multinucleate cells (up to 100 in potato cyst nematodes forming the syncytium) that develop near the heads of the feeding worms. Examples are the root-knot and false root-knot nematodes (Meloidogynidae and Pratylenchidae), which are common in warm temperate and tropical climates rather than in Britain, and many species are major crop pests (the potato cyst nematodes Globodera spp. are pests in Britain).

(2) Swellings mainly in stems caused by Ditylenchus dipsaci (Anguinidae), the commonest nematode gall that naturalists are likely to come across in Britain. It is very polyphagous, recorded from more than 450 hosts, although it does not cause distinct galls in all of these. At first the galls are solid but, as in type (1), large air spaces develop due to cells separating and collapsing. Typical nutritive cells (see Chapter 1), rather than giant cells, develop in patches where the nematodes are feeding.

(3) Distinct, well-structured galls with a cavity, in stems, leaves, florets and occasionally roots, usually on grasses (Poaceae) and daisies, thistles, knapweeds and their relatives (Asteraceae). Typical nutritive tissue occurs in a distinct layer around the cavity.

(4) ‘Cauliflower’ galls on strawberry Fragaria caused by Aphelenchoides ritzemabosi in the shoot tips and crowns of the plants. The typical fasciation develops only if the bacterium Rhodococcus fascians is present inside the cells of the plant. The nematodes live in a film of water between the stunted, crowded flower stalks and leaves. Little is known about the development of this gall; it could be caused mainly by the bacterium, which causes virescences on its own on a variety of plants (Chapter 2, Table 2.4 and Bacteria and phytoplasma virescences).

(5) Galls on Myrtaceae (Eucalyptus and its relatives) caused by a unique mutualism between gall flies of the genus Fergusonina (family Fergusoninidae) and nematodes in the genus Fergusobia (Neotylenchidae). The galls develop in meristematic tissue of shoot tips and stems, and also of flower buds. They contain one or more chambers lined with nutritive tissue, each with a fly larva and a number of nematodes. Their detailed structure is not well known. Both gall fly and nematodes are required for these galls to develop–a three-way association (with the plant) reminiscent of gall midge ambrosia galls (see below and Chapter 9).

BOX 8.1. Biology of nematodes. (Information from Southey in Redfern et al., 2002; Taylor et al., in Raman, et al., 2005.)

Nematoda is a large phylum of very small worms, variously known as roundworms, eelworms, threadworms and hookworms. They are very common but are rarely encountered by naturalists as most are hidden in the soil or are parasitic inside the bodies of plants and animals, and are mostly microscopic in size. They are unsegmented worms, transparent or pale-coloured, with a fairly uniform structure despite their variety of lifestyles. Most gall causers in Britain belong to the families Anguinidae, Meloidogynidae and Pratylenchidae (all in the order Tylenchida). In Australia, an unusual three-way association results in galls on eucalypts and other Myrtaceae: plant–Fergusonina gall flies–Fergusobia nematodes cooperate in a mutualistic association reminiscent of the plant–fungus–cecidomyiid midge relationship characteristic of ambrosia galls. Fergusobia belongs to the Neotylenchidae, also in the Tylenchida. The biology of gall-causing nematodes is not well known apart from pest species that damage crops; species that are parasites of humans or stock animals are better known too.

Nematodes feed by piercing and sucking, using their needle-like stylets and strong muscular pharynx. The plant parasitic species pierce plant cells, inject enzymes that dissolve the cell walls and suck out the juices. They feed from the surface of the plant or burrow inside, destroying more cells as they burrow.

Reproduction is usually sexual and the sexes are separate. The female produces a pheromone that attracts the male, who swims towards her. They mate and she lays eggs, and subsequent development is straightforward: the larva that hatches is a small immature version of the adult; it moults through three or four stages before becoming an adult male or female. In the potato cyst nematodes, the female dies after mating and her body hardens into a cyst to protect the eggs developing inside her.

In addition to these five groups, the dagger (Xiphinema) and needle (Longidorus) nematodes (both belonging to the Longidoridae) cause solid swellings of the root tips of a variety of plants (Fragaria, Rosa, Rubus, Daucus, Apium, etc.) but do not live inside the swellings. They are not true galls under the definition used here (see Chapter 1) but are clearly gall-like growths. These are important crop pests because they are often vectors of virus diseases.

Two nematode species are described that illustrate the extremes of complexity of nematode galls. Both belong to the family Anguinidae: Ditylenchus dipsaci causes a simple swelling and Subanguina picridis (not a British species) causes a more complex, well-structured growth. Both induce the plant to produce special nutritive cells on which the worms feed, making these galls rather more complex than the beetle and moth galls already described although, superficially, the galls of all three groups can look similar. In addition, the fascinating mutualism of the Fergusonina–Fergusobia galls on eucalypts is described in more detail.

Ditylenchus dipsaci is unusual in that it causes galls in a wide variety of host plants. Usually the galls are in the stems, e.g. on creeping thistle Cirsium arvense and germander speedwell Veronica chamaedrys, but they also occur in flower stalks and in leaf veins, e.g. on ribwort plantain Plantago lanceolata. The worms also distort and rot the bulbs, tubers and roots of many species without causing galls. Some host species are important crop plants in which the nematode causes considerable damage (e.g. ‘bloat’ disease of onion Allium cepa and ‘tulip root’ of oats Avena sativa). In all hosts, young plants are more susceptible to attack than older plants and galls formed on them tend to be larger. The species D. dipsaci contains many biological races and it is likely that each is more restricted in its range of hosts.

The following description applies particularly to the stem gall on creeping thistle. Larvae enter the young stem from the soil by piercing the epidermis and this causes the cortex cells to enlarge and separate and their walls to break down. At this stage, the swelling is due mainly to enlarged air spaces within the cortex of the stem. The worms feed on the cell contents and grow, and move upwards, congregating amongst the young cells beneath the apical meristem. These cells respond by dividing and forming patches of nutritive cells around the heads of the worms, and vascular strands grow to link with the main vascular bundles of the stem. Many of the nutritive cells are typical with dense cytoplasm and enlarged nuclei like the nutritive cells of insect galls (see Chapter 1, Types of Galls), while some contain many nuclei. The worms destroy the nutritive cells as they feed producing irregular cavities, and new cells develop to replace them. The worms tend to concentrate around the borders of the cavities especially within or near the vascular strands. As numbers of worms in the gall increase through the summer, the whole plant may become stunted with its terminal internodes crowded together, and the stem and leaves turn yellow or brown. A large population of worms can kill the plant. Two months after infestation, all stages from eggs to adults are present in the gall. The worms escape from the gall when the plant dies back and rots at the end of summer and they can survive in the soil for several years before infesting new stems in the spring.

Subanguina picridis is restricted to Russian knapweed Acroptilon repens and possibly a few closely related knapweeds, and causes more complex galls than D. dipsaci. Russian knapweed is native to southern Russia, Tadzhikistan and Armenia, and was accidentally introduced into Canada with Turkestan alfalfa in the early 1900s. It has become a noxious weed of grassland, reducing the amount of pasture for cattle. In the 1980s, the nematode and its gall were investigated for the biological control of this weed, and the gall was introduced into Canada (see Chapter 13). Neither the plant nor the nematode is found in Britain, but its galls (Fig. 120a) are similar to those of a related species Subanguina millefolii on yarrow (Achillea millefolium, Fig. 120b).

Galls develop in spring in stems, leaf stalks and veins, and the root collar of the plant. The worms invade the apical meristem of the rosette as it sits on the soil surface for a few weeks before extending upwards. The stems become distorted and the leaves develop rounded swellings up to 12 mm across. A section through a mature gall shows distinct layers of cells with nutritive cells lining a central cavity containing the nematodes. Some of the nutritive cells are sucked dry and collapse, while new cells develop in response to feeding by the worms. Outside the nutritive layer is a broad zone of tightly packed parenchyma cells. Vascular strands run through this zone and join the vascular bundles of the leaf, so supplying the gall from elsewhere in the plant. The epidermis of the gall has a thicker cuticle and is hairier than the normal leaf. At the end of summer, the plants die back and rot and the worms escape into the soil. The second-stage larvae become dormant and overwinter. Next spring, they become active and must live free in the soil for at least a month before they are capable of penetrating the terminal bud and inducing a gall. This is true also of the species on yarrow.

FIG 120. (a) Gall of Subanguina picridis at the base of a leaf of Russian knapweed Acroptilon repens (scale = 5 mm; modified after Watson, 1986). (b) Galls of Subanguina millefolii on yarrow Achillea millefolium (from Redfern et al., 2002, with permission).

Fergusoninid flies were described from Eucalyptus in Australia in the 1920s and their mutualistic association with nematodes first worked out a decade later. About 80 galls have been identified and there are probably many more to be discovered. It seems that all are species-specific; each Fergusonina fly species has its own Fergusobia nematode species and their galls are specific to particular eucalypt host species. They develop in stems, leaf petioles, leaf or flower buds, and most are small, 2–3 mm across, rounded and with one chamber, or twice as large and with several chambers.

The lifecycles of the fly and the nematode are complex (summarised in Fig. 121) and are synchronised with development of the host plant. The nematode is at first parasitic in the adult fly: a female worm lays eggs inside the female fly, they hatch and juvenile worms are deposited into the meristematic tissue of eucalypt shoots or buds as the fly lays its eggs. The gall grows before the eggs of the fly hatch; probably the feeding of the young nematodes stimulates cell proliferation and development of nutritive tissue. The fly larva and nematodes develop separately in a gall and both feed on the gall tissue. Four weeks after oviposition, the fly larva hatches and grows through three instars before pupating in the gall. Meanwhile, the nematode develops through two generations. The juveniles deposited with the fly’s egg grow into parthenogenetic females that produce more juveniles. These develop into males and females that mate in the gall, and a fertilised female worm enters the full-grown fly larva before it pupates. So, when the fly adult emerges from the gall, it contains a female nematode, which lays eggs that hatch into juveniles ready to start the next generation. The nematode causes the gall and feeds on plant tissue, but is also parasitic inside the body of the fly, while the fly larva is entirely phytophagous. It seems that both depend on the other for survival: Fergusonina depends on the nematode for the growth of the gall, its food and shelter, while Fergusobia relies on the fly to transport it to new host plants and to insert its offspring into tissues suitable for their development.

FIG 121. A summary of the lifecycles of Fergusonina flies (dashed arrows) and Fergusobia nematodes (solid arrows) in galls on Eucalyptus (modified after Taylor et al., in Raman et al., 2005).

GALL MIDGE STEM GALLS

Gall midges that cause stem galls belong to different genera from those that cause leaf rolls, pouches and rosettes (described in Chapters 4, 5 and 6). Several unrelated species attack a wide range of plants, trees and shrubs as well as herbaceous species, though each species is specific to only one or a few related host plants. Some genera have specialised on particular plant families, e.g. Mayetiola on grasses (Poaceae), Planetella on sedges (Cyperaceae) and Rabdophaga on willows and poplars (Salicaceae). Most of the stem galls are not well known and so their range of complexity is difficult to assess, but most include specialised nutritive and other tissue layers. Of special interest are the ambrosia galls–galls that contain a fungus, with both gall midge and fungus dependent on each other and the plant (mutualism). Ambrosia galls often have a relatively simple structure, without specialised layers of tissues, but the relationship between the plant, gall midge and fungus is anything but simple. Stem galls of several gall midges are described: two species of Lasioptera followed by the more complex galls of Giraudiella and Rabdophaga. Other genera (e.g. Asphondylia) gall buds, flowers and fruits (see Chapter 9).

Lasioptera arundinis causes ambrosia galls in side shoots of the common reed Phragmites australis. Side shoots develop when the terminal bud of the main stem has been damaged. This often happens and is due, for example, to grazing caterpillars or to attack by Lipara gall flies (see Chapter 6, Cigar Galls, caused by Lipara Species). When galled by L. arundinis, the shoot is slightly thicker than normal with shortened, crowded internodes. The leaf sheaths must be removed to reveal slits stuffed with a black fungus. The fungus obscures the entrance to tunnels, and each tunnel contains a pale orange gall midge larva. The fungus is an anamorphic hyphomycete, i.e. the asexual stage of a hyphomycete species, and has proved difficult to identify (Yukawa & Rohfritsch, in Raman et al., 2005, name it as Ramichloridium subulatum; but see also Chapter 9, Gall Midge Leaf, Bud and Fruit Galls). The development and lifecycle of the gall midge and the fungus are closely integrated and adapted to the plant. The gall midge is specific to Phragmites australis, although the fungus may also be able to survive in other ambrosia galls.

L. arundinis has one generation a year. Adults emerge from the old galls in late June or July, mate, and the females lay batches of eggs on or just above the nodes of the side shoots, under the bases of the leaf sheaths. The larvae hatch and crawl towards the tip of the shoot, in a line one behind the other. The hatchlings bear long bristles and spines that trap spores and fragments of hyphae of the ambrosia fungus and, as they move up the shoot, they deposit the spores on the surface. The female deposits the spores with the eggs–she picked up the spores earlier after she emerged from her old fungus-infested gall. During their migration, the larvae suck and wound individual cells of the shoot epidermis, and these stop growing so enabling hyphae germinating from the spores to penetrate between them. Each hypha grows between the cortex cells of the stem penetrating deep until it reaches a vascular bundle. It macerates cell walls and kills some cells along its route, creating a channel. The larva follows it into the stem and enlarges the channel by destroying cells too. It is clear that both the larva and the fungus produce enzymes that digest the middle lamella (the layer that glues the cells together). In the normal Phragmites stem there are two rings of vascular bundles, one inside the other. The fungus penetrates the parenchyma of the inner, larger, bundles and causes these cells to modify into typical nutritive cells (Chapter 1). Both the larva and the fungus feed on the nutritive cells and the larvae may also feed on fungal hyphae (although this is not known for sure), and they, the larvae, soon moult into the second instar. Damage to the outer vascular bundles and cortex is prevented by the plant, at least when the stem is young and actively growing; the cells of the outer ring react to the fungus by producing lignin, which prevents growth of the hyphae. So, damage is limited and the shoot can continue to grow–and so benefit the fungus and gall midge in the long run as well as the plant. The fungus continues to penetrate deep into the pith, followed by the larva. In late August or September, the larva moults into its final instar. The cells of the inner vascular bundles and the pith cannot produce lignin, interestingly, unless the same part of the stem is attacked by another gall midge Giraudiella inclusa (see below), which is not an ambrosia species. If this happens, lignified cells are produced and they prevent the fungus from contaminating the G. inclusa gall. By mid-or late September, the galls of L. arundinis are mature, the larvae full-grown, and the stem is becoming hard and dry. Each larva lines its chamber with silk and overwinters. In the following May, it uses its sternal spatula to cut an exit hole through the fungal mycelium that blocks the original entrance tunnel, leaving a thin window of plant epidermis across the exit. It retreats back down the burrow and pupates in June. In July, the pupa moves out through the tunnel, revolving like a screw and using the spiny horns on its head to break the window, and the adult emerges. After this, the fungus matures and produces spores near the top of the Phragmites shoot. The midges mate and females collect the spores into ‘mycangia’ (special pockets) at the tip of the ovipositor, before flying to new reed shoots to start the next generation. The pockets and sensory hairs of the spore-collecting apparatus ensure that spores of the right size (5–8 x 1–2 µm) are collected and therefore likely to include the correct species.

Survival of the fungus and the gall midge is bound together, each benefiting from the behaviour of the other and both dependent on the host plant. Initial damage by the hatchling enables the fungus to penetrate the stem, which then allows the larva to enter. The fungus modifies the vascular parenchyma into nutritive tissue and this creates a strong nutrient sink and feeds both the larva and the fungus. The fungus helps to provide an exit route for the adult–without this, the gall midge might have to wait until the plant rotted before it could escape. Another benefit to the larva is protection from parasitism. The fungus makes the galls difficult for parasitoids to detect and deters them from drilling into the stem. A final advantage to the fungus from the gall midge is dispersal of its spores. The female carries them to new reed shoots, a much less chancy process than dispersal by the wind, and the behaviour of the hatchling larvae ensures that the spores can penetrate precisely the correct spot on the stem.

FIG 122. (clockwise from above) Stem galls on bramble Rubus fruticosus of: (a) Lasioptera rubi (Michael Chinery) and (b) young and old galls of Diastrophus rubi. (Michael Chinery & Tom Higginbottom)

Lasioptera rubi causes ambrosia galls in the stems of brambles (Rubus fruticosus, Fig. 122a) and, less commonly, in other Rubus species (e.g. raspberry R. idaeus, dewberry R. caesius and stone bramble R. saxatilis). The females collect spores from the surface of their old galls; the fungus sporulates soon after the adults emerge. The L. rubi gall is quite common but its structure is less well known than the L. arundinis gall. The mature L. rubi gall is rounded and woody, up to 5 x 2 cm in size and with characteristic splits and cracks in its surface. These occur because growth of the bark and outer tissues of the stem does not keep up with the swelling within. Inside are several irregular cavities lined with fungus and containing larvae that are reddish orange when full-grown. Superficially, this gall is similar to that of Diastrophus rubi (Hymenoptera: Cynipidae, Fig. 122b), especially when both are young. Their structure is entirely different, though (see below for D. rubi).

Like L. arundinis, L. rubi has one generation a year. The females lay eggs in May, at the base of buds or on actively growing young bramble shoots. The larvae enter the stem, grow though the summer, overwinter when full-grown and pupate in the gall in spring. The females carry the spores of the ambrosia fungus and probably deposit them with the eggs, and the fungus and hatchlings probably help each other to enter the stem, perhaps in much the same way as described for L. arundinis. Inside the stem, the larvae in separate tunnels burrow across the cortex and vascular tissue to the pith in the centre, aided by the fungus (both producing digesting enzymes). The cambium of the vascular tissue reacts by producing extra cells; perhaps these modify into nutritive cells as in the L. arundinis gall or perhaps the larvae feed entirely on fungal hyphae. In galls with many larvae, the larval tunnels often coalesce as they enlarge, and the mature gall becomes hard and woody.

The galls of Giraudiella inclusa in stems of common reed and of two species of Rabdophaga in willow twigs are not ambrosia galls. These galls have a more complex structure, with several layers of specialised tissues encircling the larval chambers, their complexity reminiscent of the galls of gall wasps (Chapter 9).

The structure of the Giraudiella inclusa gall is quite different from the gall of Lasioptera arundinis, although both may occur in the same shoot of the reed. G. inclusa galls do not cause the stem to thicken and are not usually obvious from the outside. Peeling away the leaves and leaf sheaths reveals the slit-like opening to each gall. G. inclusa has two generations a year in the warmer areas of Europe but usually only one in the colder north and west. The galls of the two generations appear so different that they were originally described as two species, but development of each generation is similar. Eggs are laid on the surface of an internode, under its enclosing leaf. The larvae hatch and crawl downwards and settle in a row above the node to feed. Beneath each larva, the epidermis cells stop growing while, either side and around it, the cells multiply, causing the larva to sink into a pit. The pit deepens into a pouch and its lips grow up and over the larva. The first-generation larvae are almost completely enclosed inside except for a narrow slit opening. When full-grown their galls are 3–10 mm long, ovoid, and look like grains of rice attached to the inner surface of the stem. They occur low down in the main stem and if there are many, they may coalesce. The second-generation galls occur higher up in the main stem or in lateral shoots. They are about the same size as first-generation galls, but with wide openings that are closed by silk spun by the larva when it is full-grown. In both generations as the gall enlarges, it projects into the parenchyma of the stem and links up with nearby vascular bundles. These supply the nutritive tissue that develops around the larva. The galls develop in a similar way to the pit and pouch galls caused by gall midges on leaves (see Chapter 5). In reed stems, although the galls are not completely closed, they are tightly covered by the leaf sheath, which forms a protective covering. The full-grown gall (Fig. 123), of both generations, has a layer of nutritive tissue lining the chamber on which the larva feeds. The outer gall eventually becomes hard and woody, surrounded by the remains of the vascular tissue that supplied it, and projects into the central lumen of the stem, now hollow. If a gall of Lasioptera arundinis develops in the same part of a lateral shoot, the G. inclusa gall may not have space to sink into the centre of the stem. Instead, it projects above the stem surface, forming a woody bump. The G. inclusa gall resists being overrun by L. arundinis’s ambrosia fungus by developing between them a line of resistant lignin cells that the fungus cannot penetrate. So, the two galls can live together in the same stem without interfering with each other.

FIG 123. Diagrammatic section through mature first generation galls of Giraudiella inclusa enclosed in the stem of common reed Phragmites australis (the opening at the top is slit-like; simplified after Meyer & Maresquelle, 1983.) Larva (black) in its chamber; nutritive tissue (stippled); woody layer (hatched); vascular bundles (open circles).

Adults from overwintered G. inclusa galls emerge in May and lay eggs on the new reed shoots. In the south of its range, the galls and larvae develop rapidly and produce adults in July. These mate immediately and lay more eggs on plants that are now considerably taller, and the second generation develops. These larvae are full-grown in September and overwinter inside the dry and dead reed shoots. They pupate in April and early May, and the adults emerge later in May. Where there is just one generation, development is slower and larvae are fully fed in late summer. These individuals overwinter and emerge the following May at about the same time as the second generation of adults.

Rabdophaga is a large genus of gall midges that has specialised on willows and several species cause complex closed galls in the stems. R. salicis on sallows (Fig. 124) is common. Mature galls are woody and contain several larvae, each in a separate chamber enclosed in layers of specialised tissues. The lifecycle of R. salicis and development of it’s gall are not known in detail but a related species, R. degeeri on Salix purpurea, has been described (Meyer & Maresquelle, 1983; Meyer, 1987) and R. salicis may be similar. Both species also gall the petioles and midribs of leaves.

R. degeeri has one generation a year in Britain and western Europe. The female has a long, telescopic ovipositor that cannot pierce plant tissue; in May, she lays clutches of eggs on the young stems just above the nodes. The larvae hatch in a few days and burrow into the stem and deep into the pith, excavating individual galleries that cut across the vascular tissues. The stem starts to swell immediately due to increase in size and number of most of its cells. Strands from the vascular bundles grow towards each larval chamber and, ten days after the eggs hatched, a thick layer of nutritive tissue has developed around each larva. After 28 days, a woody shell develops outside the nutritive layer and, two weeks later, the gall is full-sized. Vascular strands pierce the woody shell and continue to supply the nutritive tissue until the larva is fully fed. The larvae are full-grown in September and overwinter in their chambers. In April, each larva cuts a tunnel to the surface probably using its strong sternal spatula, leaving a window of bark across the exit. It pupates back in the chamber and, in May when the adult is ready to emerge, the pupa wriggles to the exit, breaking the window with tough horns on its head (in a similar way to Lasioptera arundinis, above). After the adults have emerged, their pupal skins are left projecting from the exit holes. They live only for a day or two, during which time they must mate and the females lay their eggs to start the next generation.

FIG 124. Stem gall of Rabdophaga salicis on grey willow Salix cinerea. (Tom Higginbottom)

TEPHRITID STEM GALLS

In Britain, tephritid galls are characteristic of the flower heads of thistles and other Asteraceae (Chapter 7). Worldwide, though, stem-and root-galling species are more common and they also specialise in the Asteraceae. Two stem gallers have been studied intensively: Eurosta solidaginis on goldenrod Solidago in North America (not a British species; Fig. 125) and Urophora cardui on creeping thistle Cirsium arvense (Fig. 126). Both species cause complex, closed, woody galls that are highly organised. E. solidaginis’s gall contains one chamber with one larva, although several galls may run together if they are common, while U. cardui’s gall usually contains several larvae in individual chambers and is slightly more complex. Otherwise their galls are similar rounded or ovoid swellings in the stems.

Eurosta solidaginis has a wide distribution in North America from southern Canada to Texas and northern Florida, and from the east coast to the foothills of the Rockies. It galls several closely related goldenrods, with distinct subspecies or races specialising on different host species across this range. It has one generation a year. Adults emerge in late May or June from last year’s galls and live for about 10 days. The male sits on the tip of a shoot and, if a female approaches, he starts an excited courtship display (similar in all tephritids, see Chapter 7) and attempts to mount her from the rear. Often she reacts by walking or flying away but, if she is receptive, she will display too and allow the mating. She lays her eggs in the long thin buds of goldenrod and, before laying each egg, she checks out the bud. She climbs to its apex and rubs it rapidly with her forelegs, drawing the tip through her mouthparts. If it passes muster, she walks to the base of the bud and finds a suitable spot to insert her ovipositor, forcing it down between and into the young leaves. She normally pierces the leaves several times before depositing one egg on the surface of an inner leaf. If she is in a good patch of goldenrod shoots, she flies less than 1 m before laying another egg, but she is a strong flier and can fly several kilometres to find a new host patch if necessary. Each female contains over 200 eggs, though she is unlikely to have time to lay all of them, particularly if the weather is unfavourable during June.

FIG 125. Ball gall of Eurosta solidaginis on goldenrod Solidago altissima. (Robert Cameron)

FIG 126. Stem gall of Urophora cardui on creeping thistle Cirsium arvense. (Michael Chinery)

The egg hatches in about a week and the first instar larva crawls down to the meristem of the bud and burrows several millimetres into the stem beneath, feeding as it goes. The plant can fight back at this stage and may kill the young larva by enveloping it in wound cells that die and harden, effectively isolating the larva. If it survives, the larva mines into the centre of the stem, through the developing vascular tissues. Here, it burrows up and down, eating out a small chamber, and the cells around it multiply and differentiate into concentric zones of tissues, all derived from the pith of the normal stem. The swelling is apparent three weeks after the larva hatched. The gall grows rapidly and is full-sized, 2 or 3 cm across, three weeks later (the ungalled stem is only about 3 mm in diameter). At this stage, the central chamber is small and the larva, although now in its second instar, is growing only slowly. Lining the chamber are the nutritive cells, a 2 mm wide layer of small cells with dense cytoplasm on which the larva feeds. As it feeds it destroys these cells and new ones replace them from the next zone, a 2 mm wide layer of parenchyma cells, thin-walled with large vacuoles. Outside this layer and forming the bulk of the gall is the gall cortex, formed of large thick-walled parenchyma cells. Running through the gall cortex and parenchyma layers are numerous vascular strands that link with a ring of vascular bundles near the outside of the gall. These are continuous with the normal vascular bundles of the stem above and below the gall and are not greatly modified, and supply the nutritive tissue and the larva for as long as it is growing.

When the gall is full-sized, in early August, the larva starts growing rapidly. It moults into its third instar and is full-grown by mid-September. It prepares an escape tunnel to the edge of the gall, leaving a window of epidermis across the exit (Fig. 127). The gall soon dries out and becomes woody. The larva retreats back into the central chamber to overwinter. In spring, the puparium forms, the pupa develops inside and, in late May or June, the adult crawls out through the prepared tunnel, breaking the window with its ptilinum (the inflated balloon-like structure on the heads of emerging ‘higher’ flies, enabling them to break out of their puparia).

Although the galled goldenrod shoot grows to the normal height, it does so more slowly than its ungalled neighbours. The shoot will flower but produces fewer flower heads and fewer, smaller seeds (achenes) than if ungalled. Fewer resources pass to the rhizomes beneath the galled shoot too, which affects growth in future years. Advantages of the gall to the tephritid fly seem clear–as in all galls, the galler acquires food and a protected place in which to live. Protection against parasitoids and predators is two-edged, however. Although general predators cannot find the larva, many chalcid parasitoids and insectivorous birds (chickadees, i.e. tits, and woodpeckers) are attracted to the highly conspicuous galls and are adapted to oviposit or peck holes in them to reach the larvae inside (discussed further in Chapter 10). Protection against harsh weather is often assumed, too. The gall protects its larva from rapid changes of temperature, but it is not a good insulator. The temperature inside follows the ambient temperature outside although, if it is sunny, it can be warmer inside. The larvae can withstand being frozen (and temperatures on the prairies in winter can be severe). When overwintering, their bodies are rich in fat and an antifreeze prevents ice crystals from forming in the cells. The main physical protection that the gall provides the larva is against drought (it is always moist inside a plant) and direct contact with rain, ice, snow and sunlight.

FIG 127. Section across an overwintering goldenrod ball gall caused by Eurosta solidaginis on Solidago altissima (after Abrahamson & Weis, 1997).

Urophora cardui feeds only in creeping thistle (Cirsium arvense, known as Canada thistle in North America) and forms a distinct stem swelling (Fig. 126). Its lifecycle is similar to that of Eurosta solidaginis–adults fly and lay eggs in June or early July, larvae feed from July to September and the third instar overwinters in the gall. The larvae pupate inside puparia in late April and May, and adults emerge in early summer. Although creeping thistle is common throughout Britain, U. cardui is mainly found in the south, south of central Yorkshire and mid-Wales. It does seem to be slowly extending its range northwards, however, in mainland Europe as well as in Britain (it was found in S. Yorkshire for the first time in 2006).

FIG 128. Urophora cardui on creeping thistle Cirsium arvense. (a) A female laying eggs in a side shoot. (b–c) Vertical sections through the young shoot and mature gall: (b) eggs on the meristem and second instar larvae tunnelling into the stem; (c) the mature gall with full-grown larvae. (After Redfern & Askew, 1998, from Lalonde & Shorthouse, 1984.)

U. cardui males, like E. solidaginis, establish territories on thistles that include suitable egg-laying sites for the females, and their behaviour is similar to the goldenrod gall fly. The male probably marks suitable shoots with a pheromone, which attracts females and increases his chance of mating. Females lay small clutches of eggs (a maximum of about a dozen eggs in a clutch) between the immature leaves in an expanding terminal or axillary bud (Fig. 128a). The larvae hatch in seven to ten days. As in other Urophora species (see Chapter 7), it is the second instar that hatches, the first having moulted inside the eggshell. The larvae mine downwards in individual tunnels, skirting the meristem (Fig. 128b), until they reach the zone where the vascular tissue of the new shoot is differentiating. They move to the inside of the new vascular bundles and excavate individual chambers, and the gall tissues proliferate around them. Their tunnels fill with callus cells derived from cells wounded as they tunnel. The gall grows rapidly while the larvae, at first, grow slowly. The new cells of the gall are derived from the stem pith and the cambium of the vascular bundles. The pith cells multiply to form parenchyma that contributes most to the swelling, and new vascular strands appear in the gall parenchyma, linking with the normal vascular system of the stem and branching through the gall to form a network around each larval chamber. Nutritive cells, which develop from parenchyma cells next to the larvae, appear in irregular small patches on the chamber walls. These are similar to nutritive cells of other galls, with dense cytoplasm and enlarged nuclei (Chapter 1), and develop in response to feeding by the larva. More callus cells develop in the larva’s original tunnel, so that it enlarges as the gall grows. When the gall is full-sized, a second nutritive tissue appears in a patch at the bottom of the chamber and around the head of the larva. These nutritive cells are rich in lipids and proteins and their cytoplasm is dense but, unlike typical nutritive cells, they also have a large central vacuole. The larva now grows rapidly, demolishing any remaining primary nutritive cells, starting on the secondary cells and moulting into the third instar. The parenchyma cells start to lignify, at first either side of the callus-filled tunnel and then extending to the rest of the gall. By September, all of the nutritive tissue is eaten, the larva is full-grown and the gall is woody (Fig. 128c). Usually the gall falls to the ground, with the overwintering larva inside. Thistle stems tend to be knocked down in autumn and early winter by high winds and winter snows, and the galls become buried in litter on the ground. This is important if the larvae are to pupate successfully and the adults to emerge next June. In spring, rain and snowmelt cause the callus cells filling the larval tunnel to rot and loosen, while the woody tissue of the rest of the gall remains intact. Rotting allows air to enter the larval chamber and this stimulates pupation, and a few weeks later, the emerging adult is able to push the loose callus plug out of the tunnel with its ptilinum. If the galls remain on upright stems, perhaps because of a calm, mild and dry winter and spring, the callus plug may not soften sufficiently to allow pupation and adult emergence and the larvae will die. U. cardui galls are commonest on creeping thistles growing in damp valleys perhaps because of this requirement.

U. cardui galls vary considerably in size according to the number of larvae inside–this averages three or four but can be three times as many. The nutrient sink effect of the gall increases with number of larvae (up to five or six, then levels off in larger galls) and the weight of each larva increases too. This leads to larger, more successful adults and females with more eggs. Like other tephritids, the females have a limited time in which to lay their eggs and rarely lay all of them. They live 10–20 days and are also limited by the short time that suitable buds are available. Like Eurosta solidaginis, female U. cardui can judge the quality of the developing bud, preferring those with large meristems, and in these they lay clutches of about a dozen eggs. If the only available buds are smaller, they reduce the clutch size accordingly. Survival of the larvae is related to gall size too (discussed further in Chapter 10).

GALL WASP STEM GALLS

Most gall wasps gall buds and leaves and are described in Chapter 9; they are introduced there, and background information on their biology and classification is given in Box 9.1 (Chapter 9). Most species (in the tribes Cynipini and Diplolepidini of the family Cynipidae) gall oaks and roses and several of these cause galls in stems and roots, so should be included here, in this chapter. However, all cynipid galls are built on the same basic plan and, because some species gall buds or leaves as well as stems or roots, they are included with the rest in Chapter 9. Diastrophus rubi, which galls bramble stems Rubus fruticosus, is one exception that is included here. D. rubi is a member of the tribe Aylacini and, although its gall is similar in structure to other cynipid galls, it has a simpler lifecycle. Its gall is superficially similar to the gall of the gall midge Lasioptera rubi on the same host, described above, but its structure is quite different (both galls are shown in Fig. 122 a and b).

The gall of Diastrophus rubi is a multichambered elongated swelling in a bramble stem with bumpy surface, each bump indicating a larval chamber within. The galls are common but are often hard to find–they are more obvious when old and dead, with their many adult emergence holes, and seem to be most abundant on weak stems deep in a bramble patch.

In spring, adult D. rubi emerge from their old galls and mate. The females explore young elongating bramble shoots and insert eggs one at a time into the developing vascular tissue behind the shoot tip, at a time when the cells are capable of developing into a variety of tissues. The structure of the gall is similar to other cynipid galls (see Fig. 138, and detail in Chapter 9): an inner gall enclosing each larva and an outer gall enclosing this, each with several tissue layers. As soon as the larva hatches, nutritive tissue develops near its head and forms a single layer of cells around its chamber. Outside the nutritive cells, a thick layer of storage nutritive tissue develops, parenchyma cells packed with starch that are converted into nutritive cells as the larva feeds on and destroys the innermost cells. A woody shell consisting of about six layers of sclerenchyma cells surrounds these two layers and forms the outside of the inner gall. The outer gall is simple, consisting of a cortex of parenchyma cells and vascular tissues, and this merges with the cortex of neighbouring galls, resulting in a multichambered swelling that eventually becomes hard and woody.

The gall grows rapidly in early summer and is full-sized by July, though the larvae inside are still small. They grow rapidly through August and are full-grown in the autumn. They overwinter, pupate in the spring and the adults chew their way out as the new bramble shoots start to lengthen. D. rubi is bisexual with males making up about 30 per cent of adults (though sometimes considerably less than this). Reproduction is typical of the Hymenoptera, with fertilised eggs producing females and unfertilised eggs producing males (arrhenotokous parthenogenesis, see Box 9.1). D. rubi does not contain Wolbachia bacteria (that cause thelytokous parthenogenesis in Diplolepis species, see Box 9.2, Chapter 9) and its reproduction is less complex than in the rose and oak cynipids (described in Chapter 9).

CHAPTER 9

Galls in Buds, Leaves and Fruits

THE MAJORITY OF GALLS develop on leaves or in leaf buds, and the most familiar and well known are caused by gall wasps–gall wasp galls on roses and oak trees make up the bulk of this chapter. Gall wasps (family Cynipidae, order Hymenoptera) cause closed galls that are complex in structure and often bizarre in shape, and their relationship with their host plant is the most elaborate of any gall-causing group. Two other insect groups cause closed galls in leaves and buds. Sawflies (Tenthredinidae, Hymenoptera) cause simple swellings on willows–Pontania and Eupontania cause beans and balls on the leaves and Euura galls the buds (and also stems, see Chapter 8). The related genus Phyllocolpa induces simple leaf folds (see Chapter 4). Gall midges (Cecidomyiidae, Diptera) are included here too. This family causes the greatest variety of galls, having already appeared in Chapters 4, 5, 6 and 8. The gall midge galls on birch leaves and ambrosia galls in buds and fruits of broom and other plants provide examples of closed gall midge galls (ambrosia galls occur in stems, too; see Chapter 8). Although these sawfly and gall midge galls are simpler in structure than most gall wasp galls, the ambrosia galls are almost unique in involving a third organism–a fungus as well as the plant and insect interact to cause the gall (see also the Fergusonina–Fergusobia gall fly–nematode galls on eucalypts described in Chapter 8).

The galls in leaves, buds and fruits are described from the simpler to the more complex: sawfly, gall midge and gall wasp galls. Introductions to sawflies and gall midges are included in Chapter 4 and Boxes 4.7 and 4.8. Gall wasps are a new group, introduced here and in Box 9.1. Although most cause leaf and bud galls, a few species are included that gall fruits, specifically poppy seed capsules and acorns of oaks, and a stem gall is included in Chapter 8.

SAWFLY LEAF AND BUD GALLS

Leaf and bud galls are caused by Pontania, Eupontania and Euura sawflies on willows. Pontania and Eupontania species cause leaf galls–red bean galls in leaf blades and balls that hang down from the underside (Fig. 129a and b). Euura galls are swollen buds and stems, the same colour as normal buds, and are less obvious (Fig. 129c) than the beans and balls. Despite the considerable variations in shape, all closed sawfly galls are structurally similar. They are simple swellings caused by multiplication and enlargement of cells with no distinct tissue layers, and no nutritive tissue (see Chapter 1)–and often the original structure of the leaf, stem or bud can still be recognised in the gall.

In all sawfly galls, the egg-laying female inserts her ovipositor into the plant tissue and causes an initial swelling. In the more advanced Eupontania species, as in galls of other groups, the larva then must hatch and feed before the gall develops further. In Pontania proxima and most Euura species, the galls grow to almost full size before the egg hatches. P. proxima on crack or white willow (Salix fragilis or S. alba), Eupontania collactanea on creeping willow Salix repens and Euura mucronata on grey willow Salix cinerea are described in more detail, below.

Eupontania collactanea is specific to creeping willow, causing spherical galls that usually hang beneath the leaves (Fig. 129a). It is a northern species reflecting the distribution of its host plant, and is locally common in northern Britain and Ireland. The gall is 5–8 mm in diameter and is attached to the midrib or to a side vein with a scar on the upper side. It is green or yellowish in colour, often flushed with red, and with small warts scattered over its surface.

FIG 129. (clockwise from top left) Sawfly galls in leaves and buds of willows, Salix. (a) Eupontania collactanea on creeping willow S. repens (Tom Higginbottom); (b) Pontania proxima on white willow S. alba (Tom Higginbottom); (c) Euura mucronata on grey willow S. cinerea (Michael Chinery).

There is one generation a year. The adults emerge from cocoons in the soil in May and June when creeping willow is in full leaf, and live for about ten days. They feed on nectar and honeydew and mate, and the females start laying eggs almost immediately. The female examines the underside of leaves with her antennae, selects one and, if it passes muster, inserts her ovipositor obliquely from one side of the midrib, laying an egg into the spongy mesophyll of the other side. Two weeks later, the young larva can be seen through the eggshell and it hatches in another week. Each female initiates about 26 galls, though not all of these will grow to full size: she may fail to lay an egg or it may fail to hatch or the larva may die young. There are five larval instars that develop over the summer. The first two grow slowly while the gall matures, and then instars three to five grow more rapidly, reducing the thickness of the gall wall to little more than the epidermis. Like all sawflies, the larva is a caterpillar and it chews and destroys the cells of the gall, producing quantities of frass that accumulate in the cavity. When it is full-grown, the larva chews a circular hole, usually near the equator of the gall, and becomes a prepupa. At first it is active, like the larva. It drops out through the hole, burrows into the soil or surface litter, and spins a cocoon from silk produced by its salivary glands. The completed cocoon is a neat brownish cylinder with rounded ends, about 5.5 mm long. It has a waterproof wall with a smooth, polished inner surface and its colour darkens through the winter. Inside, the prepupa becomes short, stout and inactive and overwinters, requiring at least a month of cold temperatures (below 7°C) if it is to develop further. The pupal stage is short, lasting only a week in May, and the adult bites its way out of the cocoon later in May or in June. As in most sawflies, reproduction is sexual and haplodiploid (see Chapter 4, Box 4.7), and males are as common as females (in contrast to Pontania proxima).

Pontania proxima causes the familiar red bean galls common on leaves of long-leaved willows, particularly crack willow Salix fragilis and white willow S. alba (Fig. 129b). Bean galls grow to about 8 x 4 mm and develop in the thickness of the leaf, projecting from both surfaces. Galls are green, with the upper surface becoming red or purplish as they mature. Young galls are hard with a thick wall; this becomes softer and thinner as the larva inside feeds (Fig. 130).

P. proxima has two generations a year (Fig. 131) and is ‘thelytokous’, i.e. females produce more females (all diploid) parthenogenetically, eggs are unfertilised and males are unknown. In late May, females emerge from their cocoons in the soil and lay eggs one at a time into expanding leaf buds. The females live only a few days and do not feed, unlike Eupontania collactanea. The galls develop in the leaf blades, growing as the leaf grows, and are full-sized two weeks after oviposition, even though the egg has not yet hatched (Fig. 130b). Then the egg hatches and the larva grows rapidly. When it is half grown, in its third instar, it chews an exit hole in the gall wall and, four weeks after it hatched, it is full-grown and drops out of this hole. It burrows into the soil to spin a cocoon and pupate, and the adult female emerges in August or September. This second generation lays eggs in more young leaves and new galls develop. These grow quickly, each larva biting a hole in the gall and pushing out some of its frass, and the larvae reach their fifth instar in September or October. The larva is soon full-grown and becomes a prepupa. It drops to the soil, spins a cocoon and overwinters, its development now very like that of E. collactanea. Occasionally if the autumn is mild, some adults of this generation emerge prematurely, in December. Their adult lives are short and unsuccessful–there are no expanding buds and most leaves are old and have fallen, and the sawflies soon die. But most larvae overwinter successfully and pupate in the following May. After a week or so, the new females emerge and lay eggs as the willow leaf buds open and the new leaves expand. The size of the subsequent gall depends, partly at least, on the rate of growth and age of the leaf–eggs laid in younger, more vigorously growing leaves result in large galls whose larvae are more likely to develop successfully.

FIG 130. Sections through Pontania proxima galls (a) soon after oviposition (arrow indicates insertion of ovipositor); (b) full-sized, two weeks after oviposition; (c) with a full-grown larva, frass and exit hole, four weeks after oviposition (modified after Rohfritsch, in Shorthouse & Rohfritsch, 1992). Scale = 1 mm. Egg (open circle), oviposition fluid (dense stipples in [a]).

FIG 131. The lifecycle of Pontania proxima on crack willow Salix fragilis (modified after Leitch, in Williams, 1994).

The cause of the galls of Pontania proxima and of other sawflies has long been the subject of investigation. Beijerinck in 1888 was the first to suggest that the fluid injected by female sawflies with the egg might contain the ‘cecidogenic substance’. This idea was not new. In the 1680s, Malpighi in Italy had suggested that gall wasps caused their galls by injecting ‘ichor’, a ‘venomous liquor’, and this theory was later accepted by Linnaeus in the eighteenth and Darwin in the nineteenth centuries. Alternatively, or in addition, wounding by the ovipositor might stimulate cells to multiply into a mass of callus tissue, which then attracts nutrients from elsewhere in the plant. Because some sawfly galls also require a feeding larva in order to grow to their full size, it seems that more than one stimulus must be involved (see Chapter 1 for discussion on gall formation).

Euura mucronata galls the leaf buds of sallows causing them to become pear-shaped and twice their normal size (Fig. 129c). It is said to gall the three common sallows (S. cinerea, S. caprea and S. aurita), although recent work suggests that E. mucronata may be a complex of very similar species, each specialising on a particular host plant. Like the Pontania and Eupontania galls, its gall is structurally simple, a mass of parenchyma cells around a central cavity. The sawfly has one generation a year. The females lay eggs into young shoots, piercing through the base of a petiole and into an axillary bud, depositing one egg at a time, and the gall grows to its full size before the egg hatches. The lifecycle is similar to that of Eupontania collactanea and males are common. The larvae feed throughout the summer and drop out of the galls in late autumn to overwinter in the soil, they pupate in late spring and the adults emerge in June. Like the Pontania and Eupontania species, E. mucronata females are attracted to young, vigorously growing sallow bushes. The galls are commonest on young plants, their abundance declining to zero on plants over 18 years old–which is not even middle-aged in the life of the tree. Sallows in sites that are regularly disturbed and stimulated to produce abundant new growth year after year are galled more permanently (in a study of grey willow S. cinerea in Finland (Roininen et al., in Price et al., 1994), regular browsing by mountain hares was shown to promote the abundance of galls). Regular disturbance of the plants keeps them in a juvenile state, with a shrubby form and abundant vigorous lateral shoots, ideal for E. mucronata. If undamaged, the plants grow tree-sized and produce fewer shoots that grow more slowly, and are less favourable for galling.

GALL MIDGE LEAF, BUD AND FRUIT GALLS

Gall midges cause closed galls in leaves and buds as well as leaf rolls, pouches, rosettes and stem galls (described in previous chapters). Two groups are included here: birch Betula leaf gall midges and gall midge ambrosia galls. Ambrosia galls are almost unique amongst insect galls in that the larvae may not feed on directly on plant tissue. Some species probably feed on a fungus whose hyphae line the gall chamber, the plant providing nutrients for the growth of the fungus (for another three-way gall, see the Fergusonina–nematode–eucalypt example described in Chapter 8, Nematode Galls).

Birch leaf gall midges

Three gall midges gall birch leaves in Britain (Fig. 132). Massalongia betulifolia causes small, shallow blisters, about 3 mm across, the same colour as the leaf and raised more on the under than the upper side. It has two generations a year, the first causing blisters in the blade and the second forming more elongated blisters over the veins. New galls appear in late May; adults probably emerge from their overwintering sites earlier in May, and mate and lay eggs straightaway. The larvae are full-grown in June and leave their galls by cutting a slit in the lower epidermis of the leaf, and each larva burrows into the litter and spins a silken cocoon. In M. betulifolia, this is about 3 mm long and roughly cylindrical with a narrow neck, like a miniature bottle. The larva pupates inside and the adults emerge later in the summer to cause the second generation of galls. It is the pupa that breaks open the cocoon, by tearing it with a pair of tough sclerotised spurs, one at the base of each antenna. The pupa then wriggles up to the surface of the litter, enabling the delicate adult to emerge without damage. The second generation of larvae leaves the galls in September or October, spins cocoons as before and overwinters inside them. They pupate in the spring and produce adults in May, which start the lifecycle again.

FIG 132. Galls of birch leaf cecidomyiids (clockwise from top left): (a) Massalongia betulifolia on yellowing leaves in autumn (Keith Harris); (b) Massalongia ruber (Tom Higginbottom), both on silver birch Betula pendula; and (c) Anisostephus betulinus on downy birch B. pubescens (from Redfern et al., 2002, with permission).

The Massalongia ruber gall is a woody swelling in the midrib, often incorporating the bases of side veins or the petiole. The gall is green at first and becomes reddish or purple as it matures, later turning brown. It is more obvious on the leaf underside, about 10 mm long and broadens the midrib by about four times. Galls of both Massalongia species contain one larva, but occasionally two or more galls coalesce, which results in more than one larva in a common chamber. In autumn when the leaves are yellowing, ‘green islands’ form around the galls of both species, making them easy to spot and no doubt extending the nutritive value of the leaf. The third species is Anisostephus betulinus (Fig. 132c). This causes conspicuous circular blisters in the blade, 2.5 mm across. The blisters are raised more on the upper surface, and are slightly paler than the leaf, often with a purplish border. They turn brown after the larva has left and no green island forms on the yellowing leaf. M. ruber and A. betulinus apparently have one generation a year. Full-grown larvae cut holes in the underside of their galls and drop out in late summer and autumn, spin cocoons and overwinter. They pupate in the spring and adults emerge in May, as in M. betulifolia.

The larvae inside the galls are distinctive. M. betulifolia larvae are oval in outline, slightly flattened, white or pale orange and with no sternal spatula in the final (third) instar. M. ruber are more elongate and yellowish when young, becoming an intense orange-red when full-grown and with a small, indistinct sternal spatula. A. betulinus larvae are ovoid, bright yellow and with a strong sternal spatula.

Ambrosia galls

Ambrosia galls are caused by gall midges in three tribes, the Alycauliini (found in North and South America), the Lasiopterini and the Asphondyliini, which both contain British species (see Table 4.8 in Box 4.8, Chapter 4). These tribes are not closely related, indicating that a close relationship between gall midge, fungus and plant has arisen independently several times during the evolution of the subfamily Cecidomyiinae. The relationship between gall midge, fungus and plant is not always clear. Perhaps the larvae have retained the fungal-feeding habits of ancestral cecidomyiids (Box 4.8) or perhaps they have reverted to fungal-feeding from more recent plant-feeding ancestors, or perhaps the fungus is an inquiline, with both it and the gall midge larva feeding from the cells of the gall.

The structure of ambrosia bud galls is simple. Growth of the gall starts in the young meristematic tissue of buds (and see also Lasioptera species in stems, Chapter 8, Gall Midge Stem Galls). The newly hatched larva prevents normal development of the bud; the fungus does not seem to initiate bud galls although full development of the gall cannot occur without it. The bud enlarges due to an increase in cell number and size, and consists of a mass of unspecialised parenchyma tissue surrounding the central chamber. There is no differentiation of tissue layers and usually no development of nutritive tissue (see Chapter 1), although vascular strands increase around the chamber. Instead of nutritive tissue, the cavity of the gall is lined with fungal hyphae that grow between several layers of cells, absorbing nutrients via haustoria (suckers) inserted into the cells. The larvae may feed on fungal hyphae but this is not proven; they may feed on plant cells lining the gall chamber.

Identification of the fungus in ambrosia bud galls has proved to be a problem. The fungus is difficult to culture in the laboratory and does not produce sexual spores in the gall (these are necessary for certain identification). It seems that the larva is able to regulate the growth of the fungus so that it lines the chamber but does not grow to fill it or to overrun the larva. Inside the gall, the fungus is kept in a juvenile state and prevented from maturing and sporulating as long as the larva is alive. After the gall midge emerges, the fungus may produce asexual spores on the surface of the gall. The fungi are probably the asexual stages of the ascomycete Botryosphaeria, in the form-genera Macrophoma and Diothiorella. These are widely distributed fungi that grow on a variety of host plants, both living and dead. It seems likely that a few closely related species of fungi grow in the galls of several species of gall midge, i.e. the fungal diversity is less than the gall midge diversity. In some plants, the ambrosia fungus may be endophytic, living permanently in the host plant and appearing in the gall if one happens to appear, and so would not be limited to the galls alone. As well as the problem of the identity of the ambrosia fungus, the relationship between gall midge and fungus is often uncertain. The gall midge may feed on the fungus or on plant cells or maybe on both. Perhaps different ambrosia gall midge species have different behaviours. An association between fungus, insect and plant has developed in the ambrosia beetles that feed under the bark of trees, too, though these are not gall inducers.

The relationship between the gall midge and its fungus is highly evolved and mutualistic. Both species are likely to benefit. The gall midge larva probably gets nutritious, easily digested food and/or protection from enemies and the fungus gets distributed to a habitat in which it can thrive–some species of fungus may be incapable of infecting living plant tissue without the help of the gall midge. Spores and perhaps pieces of hyphae are carried by the adult female gall midge in special pockets called ‘mycangia’, which have evolved independently in the different gall midge genera (mycangia occur in ambrosia beetles too). In Asteromyia (a North American genus), the spores are carried in elongate pockets, one on each side of the ovipositor. In Lasioptera, the mycangia are patches of long, strong setae (tough hairs) on either side of segment 8 of the abdomen. In some Lasioptera species, each patch folds up to form a pocket enclosing the spores when the ovipositor is retracted. Asphondylia and Schizomyia females have a long needle-like ovipositor that is retracted within the abdomen until she lays her eggs. The mycangium is a membranous sac attached to segment 7 of the abdomen, a scoop-shaped segment able to shovel up spores into the sac, and spores are laid with the eggs as they are inserted into the plant. Sometimes the fungus sporulates on the surface of the old gall after the adult has left it and she may collect the spores from this source. Asphondylia pruniperda, which causes ambrosia galls on Prunus species, does this. Some species probably search for spores on leaf litter; sometimes miscellaneous debris is found in the mycangia together with the spores of the correct species. It seems likely that the fungi can survive free-living in the environment as well as living in the highly specialised habitat of the ambrosia galls.

Asphondylia sarothamni is a common species in Britain that causes ambrosia galls in the pods and buds of broom (Cytisus scoparius; Fig. 133). It has two generations a year. Adults emerge in mid-May, they mate and the females insert eggs singly into the base of young pods in flowers that are just opening, piercing the plant tissue with their long, pointed ovipositors. The egg is white and smooth, tapering slightly towards the anterior end, and fungal spores are inserted with it into the pod. The larva hatches and at first feeds in the meristematic tissues of the young pod causing both valves to swell to form the gall. At the same time, the fungal spores germinate and white cottony hyphae spread to line the larval chamber, and they probably supplement the food supply of the larva. The pod gall is 5 mm across when full-sized in early June (Fig. 133a). The young larva is cream-coloured, changing to bright orange in its third (final) instar, with a strong, heavily sclerotised sternal spatula. If the pod is very young when attacked it will be completely stunted, with no viable seed. If attacked after it has started to elongate, the pod may produce some normal seeds in its distal half. The larva is full-grown in mid-June, pupates in the gall a week later, and adults emerge from late June to mid-July. It is the pupa that breaks out of the gall. It wriggles violently causing two strong spines on its head to cut a hole through the side of the gall, then bores its way out until its head and thorax protrude, and the adult emerges leaving the pupal skin wedged in the hole. In July, after mating, these adults lay eggs in the tiny dormant buds in the axils of leaves, buds that would produce next year’s leafy shoots if unattacked. The eggs, one per bud, remain in the buds until next February or March when the buds start to grow; the eggs then hatch, and the buds swell into ovoid galls, 5 mm across (Fig. 133b). The larvae grow through March and April feeding on plant cells or on the fungal hyphae lining the chamber; they pupate in the galls in late April, and adults emerge in the first half of May from pupae that force their way out as described for the pod generation. These adults lay eggs into the young pods to complete the lifecycle.

FIG 133. Galls of Asphondylia sarothamni on broom Cytisus scoparius (left to right): (a) in pods and (b) in buds. (Michael Chinery)

GALL WASP LEAF, BUD AND FRUIT GALLS

The gall wasp family, the Cynipidae, is relatively small compared to the gall midges (Cecidomyiidae). But gall wasp galls are many and varied and, in Britain at least, are more familiar than those of any other group–many people are familiar with oak apples and robin’s pincushions. All cynipid species are associated with galls, either causing them or living as inquilines inside those of other species. The family has an unusual biology, particularly in its various methods of reproduction. Background information on its biology and classification is given in Box 9.1.

Most cynipid galls are found on trees and shrubs, particularly oaks (Fagaceae) and roses (Rosaceae; Table 9.1, Box 9.1). The exceptions belong to the tribe Aylacini, which makes up less than 10 per cent of the family, and most of these gall herbaceous plants in the families Asteraceae, Rosaceae, Lamiaceae, Papaveraceae and others. Galls are found on all parts of the plant: roots and stems, flowers and fruits, as well as buds and leaves, the commonest sites. Most gall wasps specialise on a particular plant organ on a particular species of host; a few are more catholic, causing galls in more than one plant part and on more than one species, though the host plants are always related in the same genus or family.

BOX 9.1. Biology of gall wasps (Information from Askew, in Ananthakrishnan, 1984; Roskam, in Shorthouse & Roskam, 1992; Shorthouse, 1993; Redfern & Askew, 1998; Ronquist, 1999; Stone et al., 2002; Csóka et al., in Raman et al., 2005; Abe et al., in Raman & Gupta, 2007)

Gall wasps are plant-feeding Hymenoptera belonging to the superfamily Cynipoidea, family Cynipidae, and all of them live in galls. Their closest relatives are parasitoids of other insects and they clearly evolved from parasitic ancestors. The gall wasps are divided into several tribes in three groups (Table 9.1), which at present are all included in the same subfamily, the Cynipinae. The woody plant gallers make up a monophyletic group, i.e. its species are closely related having evolved from common ancestors (but recent DNA evidence suggests that the woody plant gallers are not monophyletic; see Chapter 11 for discussion of this problem). Inquilines feed in the galls of other species; they can modify the shape and size of galls but cannot cause galls on their own.

TABLE 9.1. Hosts plants and numbers of species of the tribes of gall wasps (Cynipidae: Cynipinae) worldwide, with examples of British genera (after Ronquist, 1999; Stone et al., 2002; Csóka et al., in Raman et al., 2005; Abe et al., in Raman & Gupta, 2007)

They make up the tribe Synergini, which may be monophyletic or consist of several independent lineages. The ‘Aylacini’ is almost certainly not a natural, monophyletic group, i.e. its members have arisen from several ancestral types. This group may be basal to the other Cynipinae, thought to have evolved from aylacine-like ancestors. But recent DNA evidence suggests that the Diplolepidini–Pediaspidini–Eschatocerini group is monophyletic and basal to the rest (see Chapter 11).

In evolutionary terms, the Cynipidae probably arose in the Cretaceous, with the earliest known fossils of adult cynipoids about 85 million years old. The main radiation of flowering plants occurred in the early Cretaceous (135 million years ago, mya) and, by 85 mya, all of the angiosperm families had appeared. So, why are most gall wasps, and the Cynipini in particular, mainly associated with oaks (Quercus) in a relatively primitive family of plants, the Fagaceae? By the Miocene (25 mya) when the Cynipini were continuing to evolve, oaks were exceedingly abundant, dominating the deciduous forests of the northern hemisphere. The Cynipini evolved in temperate climates and, like many herbivorous insects, they colonised and adapted to the most common and largest, most apparent, plants available. The Fagaceae is not a tropical family, which may explain why the tribe has not spread into the tropics. Quercus is divided into two subgenera, Cyclobalanopsis, which occurs only in Asia, and Quercus, which is widespread. The subgenus Quercus is subdivided into four sections: Quercus sensu stricto (white oaks), Lobatae (red oaks), Protobalanus (golden cup oaks) and Cerris (black oaks). Most oak gall wasps are restricted to just one of these sections, apart from species that alternate between species from two sections (see below). The Eastern Palaearctic (eastern and southeast Asia, including China) may have the greatest variety of oak species, but they and their gall wasps are less well studied than elsewhere. In the western hemisphere, North America (the Nearctic), and especially Mexico, has the greatest variety of native oaks with representatives from the first three sections of Quercus, while northern Europe is much poorer, dominated by just two species, Q. robur and Q. petraea, both white oaks. Cerris is a small section endemic to Italy, the Balkans and Asia Minor. Q. cerris (Turkey oak) has been introduced by humans into western Europe including Britain. The species richness of oak gall wasps, Cynipini, follows the host plant richness; it is greatest in the Nearctic (as far as is known at present) and the oak gall wasps may have originated there. Of the 1,000 or so oak gall wasps known worldwide, about 700 species occur in N. America especially in Mexico, perhaps 200 species in Europe and western Asia, and only 50 species are known in central and eastern Asia. However, these figures are only approximate. Numbers will increase as more species are discovered, particularly in the potentially rich but not well-studied areas of central America, China and southeast Asia. Conversely, species numbers will fall as more becomes known about the lifecycles of particular species–many species have alternating sexual and asexual generations that cause different galls (see below) and are currently, mistakenly, classified as two species. The second largest tribe of woody plant gallers, the Diplolepidini (= Rhoditini), has also diversified on a single plant genus, Rosa, which is also a temperate genus. Diplolepidini and their host plants are also more diverse in the Nearctic than in the Western Palaearctic and may also have evolved there. But interpretations may change when more is known of Eastern Palaearctic roses and their gall wasps.

Gall wasp larvae are short, stout grubs tapering towards the posterior end and curved into a C-shape at rest. They are inactive, without legs, wriggling feebly when disturbed. They are white in colour with a smooth, non-hairy skin. Their main feature is a pair of strong, toothed mandibles. All cynipids are herbivores and feed on specialised nutritive cells that develop in the gall. The inquilines also are herbivores, though they may cause the death of the gall inducer by starving it or crowding it out. Cynipid larvae feed by tearing open the plant cells and sucking up the mainly liquid contents, destroying the cells in the process. While the larva is feeding, its gut is in two separate parts, with no connection between the mid-gut and the hind-gut. Just before pupation the two parts join and the larva can defaecate–so, the larval chamber is not contaminated with faeces while the larva is feeding. The larvae pupate in the gall, without a cocoon, and the adult escapes by chewing out with its strong mandibles, through the gall wall which is often thick and woody. The adults may feed on nectar, honeydew and other sugary liquids but, probably, most of them do not feed.

Reproduction and lifecycles of gall wasps are complex, with sexual reproduction and several types of parthenogenesis occurring. The standard method of reproduction in the Hymenoptera is ‘haplodiploidy’, with males developing from unfertilised haploid eggs and females developing from fertilised diploid eggs. Thus, males are produced by the type of parthenogenesis known as ‘arrhenotoky’.

The Aylacini are the least specialised Cynipinae and include two types of lifecycle:

(1) Bisexual species, e.g. Isocolus rogenhoferi. This species may be a synonym of I. scabiosa (Nieves-Aldrey, 2001), not yet confirmed and unlikely because the galls of the two species are different. It galls greater knapweed (Centaurea scabiosa; Fig. 134a). Parthenogenesis in I. rogenhoferi is arrhenotokous as is usual in Hymenoptera. Both sexes are common, although there are usually more females (e.g. 52–70 per cent of the population). Males and females mate, fertilised eggs develop into females and unfertilised eggs into males. Males are haploid (n); in development of their sperm, the first meiotic division aborts, so that the sperm have the same chromosome number (n) as the parent male. Meiosis is normal in egg development so that the diploid (2 n) mother produces haploid (n) eggs. Diastrophus rubi, which causes galls on bramble stems Rubus fruticosus, belongs here although, because males are scarce in some populations, it may be moving towards ‘thelytoky’ (lifecycle type 2), in which unfertilised females give rise only to more females and males are redundant.

(2) Unisexual species with males rare or absent, e.g. species of Phanacis, Aulacidea, Liposthenes. Diploid (2n) females produce more diploid females by thelytokous parthenogenesis. During development of the eggs, normal meiosis occurs and results in haploid nuclei; these fail to separate and fuse together in pairs, so that the resulting eggs have two sets of chromosomes and are all female and diploid like their mother. This process allows some genetic variability so that daughters are not exact clones of their mother.

FIG 134. Examples of two types of lifecycle in cynipid wasps: (left) a bisexual species Isocolus rogenhoferi (Aylacini) and (right) Diplolepis rosae (Dipolepidini) in which males are rare (modified after Redfern & Askew, 1998).

Lifecycles of the woody plant gallers are modified from these basic types, forming a progressively more complex series through the tribes Diplolepidini to Cynipini (Table 9.1). Arrhenotokous and thelytokous parthenogenesis occur, ‘heterogony’ (sometimes known as ‘heterogyny’) is common (i.e. a sexual generation alternates with an asexual generation each year, also known as ‘cyclical parthenogenesis’) and some species are also ‘heteroecious’ (i.e. the sexual and asexual generation galls develop on two oak species in different sections of the genus). The following series (a–c) describes this increasing complexity:

(a) Diplolepidini:

Some Diplolepis species are bisexual and arrhenotokous ([1] above, e.g. the rose pea galls) but others reproduce by thelytokous parthenogenesis as in (2) and males are rare or are unknown (e.g. the multichambered rose galls; Fig. 134, right). In some species, the proportion of males is higher in more northerly populations–this occurs in Britain, Europe and in North America. So, the method of reproduction in the Diplolepidini varies by species and geographically, with some sexual reproduction occurring in more northerly, or perhaps more isolated, populations of otherwise thelytokous species.

(b) Pediaspidini:

This tribe includes Pediaspis aceris, which galls sycamore Acer pseudoplatanus in mainland Europe and is not found in Britain. It is heterogonous, causing sexual and asexual galls that are different in appearance and position, both on sycamore and a few other species of Acer. Both types of gall have a single chamber, the sexual galls on leaves producing either a female or a male, and the asexual galls on roots producing females only (of two different types). Fig. 135 summarises the lifecycle. The asexual galls produce two types of asexual females that produce the sexual generation without mating: females that lay haploid male eggs only (called ‘androphores’, reproducing by arrhenotokous parthenogenesis) and females that lay only diploid female eggs (‘gynephores’, and thelytokous parthenogenesis). The sexual galls produce males or females that mate (nearly always) to produce the next asexual generation, and all of their offspring are diploid and female. A complication in P. aceris is that, occasionally, a sexual generation female produces offspring without mating–these are diploid asexual females (as in Diplolepis, (a) above) and not haploid males.

(c) Cynipini:

This tribe includes species that show increasingly complex lifecycles. Most species are heterogonous, producing alternating sexual and asexual generations that cause different galls, like P. aceris (b). In some species, galls of one or other generation have not yet been found, or have been identified as caused by different species because their lifecycles have not yet been linked. Some species may have lost the sexual generation and become entirely parthenogenetic and a few may have only a sexual generation (though none in Britain, it seems). But both of these aberrations are rare. Biorhiza pallida causes the familiar oak apple (the sexual generation gall) that alternates with a root gall (the asexual generation). Most B. pallida populations behave like P. aceris (Fig. 135)–the asexual females emerging from root galls are either androphores or gynephores. Oak apples may produce only females or only males or a mixture of sexes, mostly one sex but with 2–6 per cent individuals of the other sex. So, not all asexual females of B. pallida are strict androphores or gynephores; some are ‘gynandrophores’, able to produce both males and females (Fig. 136). It is probable, too, that the individuals emerging from this type of oak apple are the offspring of several mothers, including androphores, gynephores and, perhaps, gynandrophores.

FIG 135. Lifecycle of the heterogonous gall wasp Pediaspis aceris (modified after Folliot, 1964, quoted in Askew, in Ananthakrishnan, 1984; Stone et al., 2002).

The lifecycle of several European species of Neuroterus, Andricus and Cynips (Fig. 137) is the most highly derived. As in Pediaspis and Biorhiza, there are two types of asexual females, androphores and gynephores (Fig. 135). As before, these produce haploid and diploid eggs respectively, and androphores produce only sons and gynephores only daughters in the following sexual generation. The sexual generation differs from Fig. 135 in that there are also two types of sexual female, which after mating can produce only androphores or only gynephores in the next asexual generation (Fig. 137). How are two types of sexual females produced? Two suggestions have been made:

FIG 136. Lifecycle of a minority of populations of Biorhiza pallida (modified after Folliot, 1964, quoted in Askew, in Ananthakrishnan, 1984). The commoner lifecycle is similar to Pediaspis aceris, Fig. 135.

FIG 137. Lifecycle of Neuroterus quercusbaccarum, which produces two types of both asexual and sexual females (a similar pattern occurs for species of Andricus and Cynips; modified after Folliot, 1964, quoted in Askew, in Ananthakrishnan, 1984; Stone et al., 2002).

The asexual gynephore females produce two types of sexual females.

The asexual androphore females produce two types of males.

Recent research suggests that the second suggestion is the more likely. Whether a sexual female produces androphores or gynephores in the next generation seems to depend on which type of male she mates with. But the sexual arrangements in this group of cynipids are not yet fully understood.

A group of closely related species of Andricus and of Callirhytis alternate between two species of oaks (i.e. they are heteroecious), both using host trees in the sections Quercus and Cerris. In the Andricus species, the sexual galls develop on Cerris and the asexual galls on Quercus, while in Callirhytis the reverse usually occurs. These species are restricted to areas where both host oaks occur together. Heteroecy seems to have evolved only in the Western Palaearctic. No examples are known from the Eastern Palaearctic or Nearctic nor from Quercus hosts in other sections.

Gall wasp adults are all rather similar and many are difficult to tell apart. Their galls are more variable and are usually characteristic of a species, and have often been used to identify the insect (a practice that can lead to misidentifications, though less often in cynipids than in other groups). The adults are small insects, with globular abdomens and the ‘wasp waist’ characteristic of the Hymenoptera, and are dull in colour–they are brown or black, sometimes yellowish or reddish brown, without the contrasting bright white, red or yellow of familiar bees and wasps and without the attractive metallic colours of many parasitic wasps. They have strong biting jaws, needed to chew their way out of their galls, which are often thick and woody. The adults probably do not feed. Feeding is the job of the larva inside the gall. The larval mandibles are designed to tear open cells so that their contents can be sucked up, first liquefied by enzymes produced by the larva. The female’s large abdomen is packed with eggs (over 1,000 in large females), with a long ovipositor coiled inside when not in use. Cynipid eggs are distinctive, with a long tail perhaps seven times as long as the rest of the egg. The egg is squeezed down an exceedingly fine ovipositor and is partially displaced into the tail. After it is laid it regains its characteristic shape. All cynipid galls start in young meristematic tissue or in the cambium of stems and roots. In many species, the eggs are laid into young leaves still inside their buds, the ovipositor inserting the eggs precisely into single cells in the position where the galls will develop. In some species, the egg is attached to the surface of a leaf and not inserted into a cell. Beneath the egg, the cells break down to cause a small hollow; the hatching larva crawls into this, and the cells around the hollow multiply to form a pad. This initial response of the plant is stimulated by the egg or by the ovipositing female. The gall will not develop further, however, unless the larva survives and feeds. It is probably the saliva of the young larva that stimulates growth of the gall. While the gall grows, it acts as a strong nutrient sink for the plant’s resources, and the larva remains small until the gall is fully developed.

The structure of all cynipid galls is similar; they are all built to the same basic plan (Fig. 138), despite their large variation in size and shape and ornamentation when mature. Each larva lives in a chamber surrounded by zones of distinct tissues (listed from the inside outwards):

A nutritive zone, with large cells rich in nutrients.

Nutritive storage cells, parenchyma cells containing starch and vacuoles; these cells become nutritive and continually replace the inner cells as they are eaten.

FIG 138. Diagrammatic section through the young gall of Liposthenes glechomae on ground ivy Glechoma hederacea showing the concentric zones of tissues typical of cynipid galls (modified after Meyer & Maresquelle, 1983). The nutritive tissue around the larval chamber is divided into an inner layer (finely stippled) and an outer storage layer (coarsely stippled); woody layer (cross-hatched), cortex (unshaded), with strands of vascular tissues (dashed lines), and the outer epidermis is hairy.

A woody shell with cell walls strengthened with sclerenchyma (this zone is missing in some galls). These three layers and the larval chamber make up the inner gall.

An outer cortex of parenchyma cells with vascular bundles encircling the inner gall and connecting to strands that extend inwards across the woody layer to the nutritive tissue, and outwards to link with the vascular system of the plant.

An epidermis, which is hairy in the L. glechomae gall and otherwise elaborated in many oak and rose galls.

The size of the inner gall is remarkably constant (diameter of about 2.6 mm when fully developed) in most cynipid galls, whether there is one chamber or many. If there is more than one chamber, an inner gall develops separately around each larva although the outer gall of each merges with the others. It is the outer gall that varies–in thickness of the cortex, whether air spaces are present, and in ornamentation of the surface. In the tiny sexual galls of Andricus kollari in buds of Turkey oak Quercus cerris, the outer gall is absent and the woody shell forms the outside of the gall. At the other extreme, for example in the oak apple of Biorhiza pallida and the hedgehog gall of Andricus lucidus on Q. robur, or in the robin’s pincushion of Diplolepis rosae on Rosa canina, the cortex is thick, enclosing several inner galls with their larvae, and the surface may be adorned with spiny processes or elaborate branched hairs. The oak apple is spongy with many small air spaces in its cortex and in the mature knopper gall (of Andricus quercuscalicis) there is a single large air space between the inner and outer galls. When the gall is full-grown and all of the layers are developed, the inner gall ceases to be a nutrient sink. The larva then starts to grow rapidly and soon devours all the cells of the nutritive and storage layers, and then lies in a large chamber enclosed by the woody shell.

Inquilinous species (cynipids of the tribe Synergini, see Box 9.1) cannot cause galls on their own but often modify the galls of their hosts, enlarging and distorting them. They live in chambers surrounded by concentric zones of tissues very like the inner gall of the gall causers. Synergines may all be related, evolved from a single ancestral type thought to have had the ability to cause a gall, an ability that was later lost. But recent DNA evidence suggests that they may have evolved from several aylacine groups (discussed further in Chapter 11). Inquilines are described further in the next chapter (10).

Development of cynipids in the gall is straightforward. There are five larval instars, a short pupal stage and an adult, which emerges inside the gall chamber before chewing its way out. Their reproductive arrangements are far from simple, however, and involve several types of parthenogenesis (Box 9.1). The less specialised species on herbaceous plants (many Aylacini, e.g. Aylax, Diastrophus, Aulacidea, Xestophanes, Phanacis centaureae) is bisexual with males making up a considerable proportion of adults. These reproduce like other Hymenoptera; females develop from fertilized eggs and are diploid and males are haploid, developing from unfertilised eggs (arrhenotokous parthenogenesis). Most bisexual cynipids have one generation a year. Adults emerge in late spring or early summer from the previous year’s galls. They mate and lay eggs in young plant tissues and the galls develop rapidly. The larvae are full-grown by autumn and overwinter in the gall, pupate in spring and the adults emerge about two weeks later to start the next generation. In a few bisexual species (e.g. Xestophanes potentillae), some populations in the southern parts of their geographical range have two generations a year. In other aylacines (e.g. Liposthenes, some species of Phanacis), males are unknown and diploid females produce more diploid females without mating (‘thelytokous’ parthenogenesis). Most of these also have one generation a year. Many Diplolepis species (tribe Diplolepidini), too, are all female and reproduce by thelytoky, and have one generation a year. Males are not unknown, however. In some European and Canadian Diplolepis species, males make up nearly 50 per cent of the adult population and these, presumably, reproduce like the bisexual Aylacini. Examples in Britain are the rose pea galls, D. eglanteriae and D. nervosa. Even in the thelytokous species, D. spinosissimae, D. rosae and D. mayri, males appear occasionally. They may have a role in some populations or be redundant, a relic of a sexual past.

Thelytokous parthenogenesis has advantages. Reproduction can be rapid when resources are plentiful and conditions benign because all individuals lay eggs so that many offspring are produced in a short time. The main disadvantage is that genetic variability in the species is reduced, which means that the species may not be able to survive if environmental conditions were to change. ‘Heterogony’, common in the oak and sycamore gall wasps (below), overcomes this problem.

Oak cynipids (the Cynipini) and sycamore cynipids (the Pediaspidini) may be related tribes. They have complex lifecycles and methods of reproduction, which involve cyclical parthenogenesis (heterogony), i.e. regular alternation between a sexual and an asexual (thelytokous) generation. Sometimes these generations alternate between different host trees, too (‘heteroecy’ see Box 9.1). Most species have two generations a year, one sexual and the other asexual, and their galls look very different. The gall containing the sexual generation develops rapidly in the spring or early summer and the asexual gall matures more slowly in late summer or autumn. Either the asexual females emerge from the galls in autumn and lay eggs that remain dormant until the following spring, or the full-grown larvae overwinter in the gall and produce females next spring. In some species, the asexual generation requires a further year to develop (e.g. in Biorhiza pallida). In other species, most or all individuals in populations in the north of their range require two years to complete the asexual generation, while annual lifecycles are commoner in southern populations (e.g. Andricus kollari). In a third variation, the sexual and asexual generations each take a whole year (e.g. Andricus paradoxus; A. albopunctatus is a synonym).

Heterogony has several advantages, in addition to maintaining the variability caused by sexual reproduction. The first generation gall is produced quickly, early in the year when new leaves are abundant and are growing rapidly. This gall is initiated by an asexual female that can lay her eggs as soon as conditions are suitable, without having to spend time finding a male and mating. The larvae and pupae in these spring galls tend to be less heavily parasitised than in later-developing galls; most adult parasitoids are active later and attack the second generation heavily. The second generation, produced after the sexual adults have emerged from the spring galls and have mated and laid eggs, develops more slowly in tough, often woody, galls able to protect the asexual females through the following winter.

Until the late nineteenth century, no one realised that heterogony occurred in oak cynipids; it was discovered in 1873 by Bassett in the USA and, independently, by Adler in 1881 in Germany. The sexual and asexual females and their galls are so different that at first they were identified as different species, sometimes even in different genera. This happens still–in several species, only the sexual or the asexual generation gall and adult are known, the pairs not yet linked. In a few species, the alternate generation is genuinely lost, but this is rare. Even in the well-known fauna of Britain, the two galls of a few species have been recognised only recently (e.g. Andricus grossulariae/A. mayri and A. lucidus/A. aestivalis were both linked in 2000). All plants, herbs as well as trees, have two main phases of growth that provide young, meristematic tissue suitable for gall formation. The first occurs in spring or early summer when the plant germinates or produces its new leaves, and the second happens later in the year when the seeds and fruits develop. Many galling species exploit one or other growth phase (e.g. the flower head tephritids, see Chapter 7), but it is unusual for one species to take advantage of both. The oak and sycamore gall wasps are unique in that they do this and also in that they cause two types of gall that involve sexual reproduction as well as both arrhenotoky and thelytoky.

How has thelytoky and loss of sex in gall wasps arisen? And how has heterogony, involving both sexual and asexual reproduction, evolved? The answer to the first question seems to involve a bacterium called Wolbachia, first discovered in 1990 in the eggs of a tiny parasitic wasp, Trichogramma (a chalcid, not a cynipid), whose larvae feed inside the eggs of other insects. Populations of Trichogramma infected with Wolbachia were all female; females produced more females by thelytokous parthenogenesis and were unable to produce males. In other populations of the same species, males were common and reproduction was normal (involving arrhenotoky as is usual in Hymenoptera). When asexual females with Wolbachia were fed antibiotics, the bacteria were killed and the females then mated with males and produced healthy sons as well as daughters. The thelytokous females had reverted to arrhenotoky. Some aylacine and some Diplolepis gall wasps are affected by Wolbachia in a similar way–see Box 9.2 for more detail. The origin of heterogony is a more complex problem and is not yet understood, but does not involve Wolbachia. So, it seems likely that Wolbachia is responsible for thelytoky in aylacines and in Diplolepis but not in the sycamore or oak gall wasps.

BOX 9.2. Wolbachia and loss of sex in gall wasps (Information from Plantard & Solignac, in Csóka et al., 1998; Plantard et al., in Csóka et al., 1998; Rokas, 2000; Stone et al., 2002)

In the 1990s, Wolbachia bacteria were discovered in several groups of insects and in some woodlice and, in all, were found to affect sex and reproduction. The bacterium was first found in the eggs of some populations of Trichogramma (a chalcid wasp), which parasitises the eggs of other insects. Infected Trichogramma were all female and produced more females by thelytokous parthenogenesis; they were unable to mate or to produce males. Uninfected individuals of the same Trichogramma species produced males as well as females, by arrhenotokous parthenogenesis as is normal in Hymenoptera. In experiments in which asexual females were fed antibiotics, the Wolbachia bacteria were killed, and the females could mate and produce offspring of both sexes. In Trichogramma, Wolbachia interferes with meiosis, the process that produces normal haploid gametes, and infected mothers were unable to produce haploid eggs. Thus, they produced diploid eggs and daughters but no sons. In the gnat Culex pipiens and in fruit flies Drosophila (both Diptera), and sporadically in other insects belonging to a variety of insect orders, Wolbachia causes sexual incompatibility–infected females mating with males carrying the same strain of Wolbachia cannot produce viable offspring. In woodlice, the bacterium feminises males even though they have normal male chromosomes.

Since 1990, Wolbachia has been found in many cynipids. In the Aylacini, it is present in about half of the species that have been tested: Liposthenes glechomae and three Phanacis species contain Wolbachia, while Aulacidea hieracii, Aylax papaveris, Xestophanes brevitarsis and Diastrophus rubi do not. All of the infected species are thelytokous, producing females (males are not unknown but are rare), while the others are bisexual, with males contributing up to 50 per cent of adults. In Diplolepis (Diplolepidini) in North America as well as in Europe, about half of those tested are infected with Wolbachia and, again, it is these species that are thelytokous, with males usually very rare (D. rosae and D. mayri in Europe). D. spinosissimae is intermediate; some populations are thelytokous with few males while in others, males are common, Wolbachia is absent from the females and sexual reproduction occurs. The rose pea gall females (D. eglanteriae and D. nervosa in Europe and four out of six species that have been tested in Canada) do not contain Wolbachia and are bisexual, with considerable numbers of males, and they reproduce only by arrhenotoky. The heterogonous gall wasps, the Cynipini and Pediaspidini, are more complex because they have alternating asexual and sexual generations. Wolbachia has been found in only 6 out of 42 of the oak gall wasp species tested. In the oak apple gall wasp Biorhiza pallida, for example, females of some populations contain Wolbachia while others do not, as in D. spinosissimae. And species of Andricus that are entirely asexual (and in those in which a sexual generation is not known) are all without Wolbachia.

It seems that Wolbachia is responsible for thelytoky in the aylacines and Diplolepis but not in the heterogenous species–in these, thelytoky alternates with arrhenotoky, and so the mechanism involved is clearly different (and is not yet understood). In the rose gall wasps, the single bisexual generation of some species becomes asexual in others due to bacterial interference with development. In the few oak gall wasps with only asexual generations, the sexual generation has been lost for a reason that does not involve Wolbachia, and may have an ecological explanation. All of the purely asexual species known are closely related, in a group that also contains sexual species and, in all of these, the two generations occur on different species of oak (e.g. in Andricus quercuscalicis: the asexual knopper gall occurs on Quercus robur or Q. petraea, and the sexual gall on Q. cerris). Perhaps loss of sex in some of the group allowed them to survive in areas where the alternate oak species was absent.

How has Wolbachia come to be associated with cynipids? Several strains of the bacterium are associated with the single genus Diplolepis and it is clear that different gall wasp species have been infected independently during the course of evolution. Transmission can occur vertically from mother to daughter in the cytoplasm of the egg; this is straightforward and, once a female is infected, all of her daughters will be also. Horizontal transmission, from species to species, is more difficult to visualise. It may occur via parasitoids that attack several cynipid host species. Insects that cause similar galls in similar positions on a plant often have parasitoids in common. This occurs in thistle head tephritid flies (see Chapter 10) as well as in the gall wasps on roses and on oaks. A parasitoid female often feeds on a host before she lays an egg, and she may transmit Wolbachia as she feeds first on one species and then on another inhabiting a similar gall. The host would have to survive in order to pass the bacterium on to its offspring. This does occasionally occur–the parasitoid may not kill the host if she was disturbed while feeding or failed to lay an egg.

Most cynipids gall leaves and buds. Examples of species from each tribe (except the Eschatocerini, Box 9.1) are described here and illustrate a series of increasing complexity. The poppy Papaver gall wasps, Aylax minor and A. papaveris, gall poppy seed capsules, the fruits. They provide examples of the simplest cynipids, the bisexual aylacines that reproduce sexually (most of these species in Britain cause galls in stems, e.g. Diastrophus rubi, see Chapter 8, Gall Midge Stem Galls, Gall Wasp Stem Galls). Thelytokous aylacines are represented by Liposthenes glechomae, which galls ground ivy Glechoma hederacea, and is a female-only species. Diplolepis species are described next, both bisexual and all-female species, and all of them gall roses. The host-alternating gall wasps are represented by the sycamore gall wasp Pediaspis aceris, included as an example of its tribe (the Pediaspidini) although it does not occur in Britain, and by several oak gall wasps (the tribe Cynipini), and these include the most complex and elaborate of all galls.

The poppy seed capsule gall wasps

Aylax minor and A. papaveris gall the seed capsules of the field poppy (Papaver rhoeas) and the long-headed poppy (P. dubium). The A. minor galls may not distort the capsules and are often not apparent from the outside. They are separate, small, rounded galls, 2–3 mm across, attached to the septa that divide the inside of the capsule into segments. A. papaveris usually enlarges and distorts the capsule, due to the swollen septa inside (Fig. 139). The individual galls develop in the septa, which swell and coalesce around each larva. Males and females emerge from their old galls in early summer, in roughly equal numbers. They mate and, in A. papaveris, the females lay eggs into the developing seeds (ovules) attached to the septa, the ovipositor piercing deep into the young capsule to its centre. The cells into which the eggs are inserted die, and the surrounding ovule and adjacent septa tissues enlarge to enclose the newly hatched larvae. The concentric zones of tissues that make up the inner gall, typical of all cynipid galls (Fig. 138, Chapter 9), develop rapidly around each larva: large nutritive cells line the chamber, storage nutritive tissue lies outside this, enclosed by a woody zone half a dozen cells thick. The outer gall is represented by the swollen septa, which enclose each inner gall and coalesce to produce a mass that is spongy in June and July, becoming hard and dry in late summer when the larvae are full-grown.

FIG 139. Gall of Aylax papaveris in the seed capsule of the field poppy Papaver rhoeas. An ungalled capsule appears in the background. (Tom Higginbottom)

Lifecycles of the two species are similar, typical of bisexual Aylacini with one generation a year. Galls are full-sized in midsummer and the larvae fully fed by early autumn. They overwinter in their galls, pupate in the following spring and the adults emerge in early summer.

The ground ivy gall wasp, Liposthenes glechomae

Liposthenes glechomae is a more specialised aylacine gall wasp, without males. It causes hairy ball-shaped galls in the leaves (galls called ‘cat’s bollocks’ in Hungarian folklore!) and occasionally in the stem of ground ivy (Glechoma hederacea; Fig. 140). The gall is green, often flushed with red when in a sunny spot, with one chamber inside (sometimes two or three chambers if galls coalesce). Each chamber contains a larva surrounded by the concentric zones of tissues typical of the inner galls of cynipids (Fig. 138, Chapter 9). When mature, the outer gall is a thick mass of parenchyma derived from the palisade mesophyll of the leaf and traversed by vascular strands that extend inwards to the nutritive zone. The strands link with the veins of the leaf and a hairy epidermis encloses the whole.

FIG 140. Galls of Liposthenes glechomae on ground ivy Glechoma hederacea. (Michael Chinery)

L. glechomae is thelytokous and infected with Wolbachia bacteria (see Box 9.2). The females emerge in May from galls of the previous year and lay eggs next to veins on the underside of tiny new leaves, just bursting from the bud. As is usual in cynipid galls, the hatching larva crawls into the hollow caused by the breakdown of cells beneath its egg, and starts to feed. At first, nutritive cells develop in patches near the larva’s head; later they form a continuous layer one cell thick lining the chamber. Vascular strands differentiate early, before the cells of the woody layer are lignified. The woody cells remain alive and store starch and, like the rest of the gall, they only develop if the larva is alive and actively feeding. The gall matures in midsummer when the larva is still small. Then the larva feeds rapidly and is full-grown in late summer. It overwinters in the gall and pupates in the spring, with females emerging in May to start the next generation.

THE ROSE GALL WASPS

The rose cynipids belong to the genus Diplolepis, the largest genus in the tribe Diplolepidini (previously known as Rhoditini), and all species are associated with roses Rosa. There is one other genus (Liebelia) in the tribe, which is found mainly in the Eastern Palaearctic and is also associated with roses. There are five Diplolepis rose gall wasps in Britain, seven Diplolepis plus one Liebelia in mainland Europe, 15–20 species, mostly Liebelia, in central and eastern Asia, and 31 native species, all Diplolepis, in North America (and three species have been introduced from Europe). Parts of central America and southeast Asia are not well studied, though, and no doubt there are more species to be discovered.

The basic structure of all Diplolepis galls follows the usual cynipid pattern, with nutritive, storage nutritive and woody tissue zones around a larval chamber. The outer gall consists of parenchyma cells and vascular tissues enclosed in an epidermis as in all cynipid galls, but varies in complexity and in the number of inner galls it contains. The rose pea galls (D. eglanteriae and D. nervosa in Britain) are very similar and each contains one chamber. The other Diplolepis galls are multichambered, with the outer layers of each individual gall coalescing, and with each chamber separate and containing one larva (the three British species are the bedeguar gall or robin’s pincushion Diplolepis rosae, D. mayri and D. spinosissimae). All species in Britain gall leaves and buds. In North America, although leaf and buds galls are common, several species gall the stems, causing single-or multichambered galls also. British species are shown in Fig. 141.

FIG 141. Galls of British Diplolepis on leaves and buds of roses Rosa spp.: (a) smooth pea galls of D. nervosa; (b) spiked pea gall of D. nervosa; (c) D. mayri; (d) D. spinosissimae; (e) bedeguar galls of D. rosae. ((a, b & e) Michael Chinery, (c & d) Gyuri Csóka)

Reproduction in Diplolepis varies; some species are bisexual, others are female-only and, in some, populations are either all female or with both males and females. The pea galls on leaflets, in Canada as well as in Britain, are bisexual with males making up 35–50 per cent of adults. Most of these species reproduce by arrhenotokous parthenogenesis and their females are without Wolbachia (see Boxes 9.1 and 9.2). Some Canadian pea gall species are thelytokous in at least some of their populations; in D. bicolor, for example, females usually contain Wolbachia and males are uncommon (e.g. 14 per cent of adults). In species that cause the more complex rose galls, in Canada as well as in Britain and Europe, males are absent or rare (usually!), most females contain Wolbachia, and thelytoky is the norm.

Despite these reproductive complexities, the lifecycles of Diplolepis species are straightforward and similar to the aylacine species: adults emerge in late spring or early summer, lay eggs into buds or young leaves, the larvae are full-grown by late summer or autumn and overwinter, they pupate in the spring, and the adults chew their way out of the gall a week or so later. All species have one generation a year, both the bisexual and all-female species.

The bedeguar or robin’s pincushion caused by Diplolepis rosae

The bedeguar is the best known of Diplolepis galls (Fig. 141e). It is very distinctive, large, rounded and covered with a tangled mass of branched wiry hairs that can be bright red on galls in sunny spots. It has been recognised since antiquity–by Pliny in the first century AD and probably earlier by Aristotle and his student Theophrastus (371–286 BC). ‘Bedeguar’ is a Persian word that refers to a kind of thistle; perhaps the branched hairs of the gall suggest spinyness or are reminiscent of thistledown. The word also means ‘wind-brought’, perhaps referring to thistledown blowing in the wind.

The bedeguar gall is widespread and common in Britain, mainland Europe and western Asia, and occurs from sea level to 1,500 m (5,000 ft). In Britain, it is found from southern England to Sutherland in northern Scotland, and in Wales and Ireland, although its distribution is patchy. It prefers warm, dry summers (with an average temperature of 18ºC or more and less than 760 mm rainfall per year) and seems to select weak and spindly host plants on light, well-drained soils that may be short of water. Perhaps vigorous plants can ‘fight back’ and prevent a gall from developing–it is rarely found on cultivated roses in gardens. The commonest host in Britain is the dog rose Rosa canina, though sweet-briar R. rubiginosa and field rose R. arvensis are often galled too. D. rosae was accidentally introduced into North America in 1868 and is now widespread from southern Canada to Virginia and west to the Rockies. There, it occurs on European wild roses, particularly dog rose and sweet-briar that were already established, introduced by early settlers.

Nearly all D. rosae individuals are females (reproduction is by thelytokous parthenogenesis, see Boxes 9.1 and 9.2). Males are always rare but are not unknown–their numbers increase in more northerly populations, though the maximum known is only 4 per cent (in northeast England). Probably they are redundant and in the process of disappearing. The females emerge over a week or two in May and June from galls of the previous year. In warm, sunny weather they are active and live for only a few days. If June is cold and wet, they live longer but are sluggish and lay fewer eggs. They lay eggs into leaf buds that are beginning to swell (Fig. 142 shows a D. rosae female ovipositing), and each female investigates a bud thoroughly, perhaps for an hour, before deciding it is suitable. She inserts her ovipositor under the edge of a bud scale and between the tiny leaflets inside without piercing any tissues, and lays 30 or more eggs. Each egg is inserted into an individual cell, usually in the main vein at the base of a leaflet. Oviposition takes an hour or two per bud, and all the time she sits motionless or spirals slowly round the bud, withdrawing and inserting her ovipositor from time to time. The eggs are typical of cynipids, sausage-shaped with a short projection at one end and a long tail at the other–in D. rosae, the total length is 2.4 mm, of which 2.0 mm is the tail. The short projection is inserted into the cell with a drop of fluid that glues it in place, and this cell dies. The egg has an immediate effect that is typical of cynipid galls. The leaflet cells adjacent to and below it become highly active, producing RNA, proteins and other cell substances and the cells enlarge, and then multiply to form a small pad (Fig. 143a and b), apparent within two days of the egg being laid. A cavity develops in the pad due to breakdown of cell walls and, a week after oviposition, the egg hatches and the larva crawls into the cavity (Fig. 143c), leaving its eggshell behind. As soon as it starts to feed, nutritive cells develop in patches near its head, and soon it becomes enclosed in a chamber that is lined with large nutritive cells, forming a layer three cells thick (Fig. 143d). Outside the nutritive tissue a cambial zone develops. The cells in this zone divide continuously, producing cells to the inside that become the storage nutritive cells, and cells to the outside that differentiate into vascular strands, woody cells and the parenchyma of the cortex. The epidermis, originally a single layer of cells, multiplies to form bulges (Fig. 143d) that become the complex hairs that envelop the gall, hairs that are multicellular. The new vascular strands grow inwards before the woody layer of the inner gall is complete and supply the nutritive tissue, and outwards into the multicellular hairs, and also to link with the veins of the leaf. This ensures that a continuous supply of water and nutrients reaches the developing tissues and the feeding larva. In the mature gall, as in all cynipid galls, the larva destroys inner nutritive cells as it feeds and cells from the nutritive storage layer replace them. The outer gall merges into a single mass enclosing the many inner galls, and all is enveloped in a mass of entangled green and reddish hairs that is the robin’s pincushion. There are several types of hairs, all multicellular and sticky when young. Some are long and branched, others shorter with a red glandular knob at the tip, and others are sharp spines. Red knobs are found on the ungalled plant too, on the edge of the normal leaflets when young, and so are not unique to the gall.

FIG 142. Diplolepis rosae female laying eggs into a leaf bud of wild rose Rosa sp. (Michael Chinery)

FIG 143. Development of the young bedeguar gall of Diplolepis rosae on dog rose Rosa canina, shown diagrammatically. (a) An egg glued into a cell. (b) Three days after oviposition, a pad is developing beneath the egg with a cavity caused by breakdown of cell walls. (c) A week after oviposition, the larva hatches and enters the cavity. (d) 12 days after oviposition, the larval chamber lined with nutritive tissue (stippled) is formed and the branched hairs typical of this gall are developing from bulges on the outer surface (modified after Rohfritsch, in Shorthouse & Rohfritsch, 1992).

The size of the bedeguar gall depends on the number of inner galls inside, which depends on the number of eggs laid and can vary between one and about 60, and the largest galls may be the product of more than one female. Galls are enlarged by inquilines, too (see Chapter 10). The choice of bud also affects size–a bud that is terminal on a shoot, or near to its tip, gets the lion’s share of the plant’s resources and grows vigorously, able to support many larvae, while further back down the stem, resources are less abundant. The smallest galls are those in unusual places, on the blade of leaflets or on petioles, stems or hips. Galls in odd places probably are more common if there is a shortage of the preferred buds with a large number of females competing for them. Gall size also depends on survival of the larvae inside. If they die very early, before or just after hatching, the pad that has started to develop is soon obliterated by normal growth of the plant. If death occurs a bit later, the inner galls may stop developing but the hairs on the outside continue to form, resulting in almost solid galls that are often bright red long after the hairs of healthy galls have turned brown. The effect of the gall on the plant is not usually serious. The plant loses some nutrients and the affected bud is killed, but damage is minor compared to other problems the bush has to face (like shortage of water or nutrients, competition from taller vegetation or browsing by larger herbivores). But a small sapling, especially if it is already struggling to survive in thin dry soils, can be knocked back or even killed if most of its terminal buds are galled.

By July or early August, four to eight weeks after eggs were laid, the bedeguar gall is full-sized. The larvae are still small; they grow slowly while the gall develops and in August are only in the second or third instar (out of five). When the gall is mature, they start feeding rapidly and put on most weight in the final instar, and are fully fed by October. By then, they have eaten all of the nutritive and storage nutritive tissues, and so the chambers are large enough for the fully grown larvae and each is surrounded by the woody shell of the inner gall. The larvae overwinter in the gall, which often does not fall off the plant, and pupate there in late spring. The adults emerge inside their chambers a week or two later, and tunnel out of the gall using their strong jaws.

Reproduction in Diplolepis rosae always involves thelytokous parthenogenesis despite the rare male that appears from time to time, and the females contain the bacterium Wolbachia (see Box 9.2). A related species is D. spinosissimae, which causes galls in the petiole or main vein of leaflets of burnet rose Rosa pimpinellifolia (Fig. 141d). This plant has a patchy distribution, growing in light, well-drained, chalky and sandy soils, and is common on coastal sand dunes and in limestone grassland. The gall populations are patchy too and are often isolated, separated from neighbouring populations by long distances. Like D. rosae, D. spinosissimae males tend to be rare (e.g. 1.5 per cent of adults), females contain Wolbachia and reproduce by thelytokous parthenogenesis. But not always. Occasionally, males make up more than 25 per cent of the adult population and the females are without Wolbachia. They mate with the males and reproduce by arrhenotokous parthenogenesis as in the bisexual Diplolepis species. Reproduction in D. spinosissimae varies geographically and the reason seems to be due to isolation. If Wolbachia were lost from an isolated population, perhaps due to antibiotics occurring naturally in the environment, reinfection would be difficult because no infected females are present nearby. Without Wolbachia, reproduction reverts to arrhenotoky, males are produced and the population becomes bisexual. In D. rosae, this does not happen probably because its host plants are much more common and widespread than the burnet rose.

A final example of a rose gall wasp involves a species that causes a stem gall. The gall of Diplolepis nodulosa is built on the same plan as other Diplolepis galls and so is included here rather than in Chapter 8 with the other stem galls. There are no Diplolepis stem galls in Britain. D. nodulosa is a North American species, common in Ontario on a native rose, Rosa blanda. It causes an inconspicuous spindle-shaped swelling in the internode of a stem–unless it is attacked by a Periclistus inquiline, when the diameter of the stem is quadrupled making the gall much more conspicuous (see Chapter 10 for the effect of inquilines on galls). The female lays a single egg into the procambium beneath the apical meristem of a bud, i.e. into the cells that produce the vascular bundles of the stem. Most Diplolepis species do not pierce the bud and leaflet tissues when they lay their eggs but, in D. nodulosa, the ovipositor cuts a channel through the tissues below the bud to deposit the egg. A cavity forms by breakdown of cell walls, as in D. rosae, the larva crawls into this and the usual zones of tissues develop. The destruction of cells by the ovipositor is reminiscent of oak cynipids (e.g. Biorhiza pallida) and is unusual in Diplolepis.

The sycamore gall wasps

The sycamore gall wasps belong to the genus Pediaspis, in the tribe Pediaspidini (see Box 9.1). Pediaspis aceris is the only well-known species, causing galls on Acer in mainland Europe and probably extending eastwards into Asia; it is not found in Britain. P. aceris causes two types of gall on sycamore Acer pseudoplatanus (Fig. 144) and on two other Acer species in mainland Europe. It is heterogonous (see above and Box 9.1) with sexual and asexual generations that alternate to produce two generations a year.

The sexual gall is a smooth ball 5–6 mm across that develops in spring on the underside of new leaves (Fig. 144a) or on petioles. The structure of the gall is not complex. Inside there is one large chamber, surrounded by concentric layers of tissues; these differ from the usual cynipid pattern (see Fig. 138) in that the nutritive tissue occurs in patches around the chamber and there is no woody layer (a structure similar to the currant gall of Neuroterus quercusbaccarum, see below). The asexual gall develops in autumn on thin rootlets, 1 mm in diameter or less. Each gall is rounded and can occur separately but, because their eggs are often laid in a row on the same rootlet, they tend to coalesce into an irregular mass (Fig. 144b). Asexual females emerge from the root galls in March and lay eggs into expanding leaf buds. The spring gall develops rapidly and is full-grown a month or so later. Males and females emerge from these galls in June and mate, and the females lay their eggs into the young rootlets. Most of these eggs hatch in the summer, producing the root galls in autumn of the same year. Fully fed larvae overwinter in the galls, pupate there in early spring, and the asexual females gnaw their way out in March to start the next sexual generation. Some eggs do not hatch the previous summer but remain dormant in the rootlets, perhaps for two years, and the length of the larval stage may be extended too. Their adults emerge in March one or two years later than individuals from the same batch of eggs, and so mix with asexual females belonging to later generations.

FIG 144. (left to right) The galls of the sycamore gall wasp Pediaspis aceris, (a) sexual galls on the underside of leaves (Michael Chinery) and (b) a coalesced mass of asexual root galls. (Gyuri Csóka)

The sycamore wasp tribe, the Pediaspidini, is possibly the sister group of the better-known oak gall wasp tribe, the Cynipini, and both tribes are heterogonous (see discussion of cynipid evolution in Chapter 11). P. aceris is simpler than most oak gall wasps in that a sexual female that emerged from a leaf gall and has mated produces both types of asexual female (i.e. androphores that produce males and gynephores that produce females, Fig. 135 in Box 9.1). The sexual females of most oak gall wasps can produce either androphores or gynephores, but not both (Fig. 137 in Box 9.1), which means that the sex of the offspring is determined earlier in the oak than in the sycamore gall wasps. But some individuals of the oak apple gall wasp Biorhiza pallida behave like P. aceris and produce both types of asexual female.

Pediaspis may be the only genus in the Pediaspidini. A second genus, Paraulax, may, however, be part of the same tribe. Paraulax includes several species that gall the southern beech Nothofagus in Chile and probably elsewhere in the southern hemisphere. More work needs to be done on this group before its relationship to other cynipids is known.

THE OAK GALL WASPS

Like other cynipids, most oak gall wasps can be identified from their galls–the outer galls vary in size, shape and structure even though the inner galls follow the standard cynipid pattern (Fig. 138, Chapter 9). The galls and lifecycles of several species are described, with examples from four genera: Biorhiza pallida, Cynips divisa, four Neuroterus species, and five Andricus species, including three heteroecious (host-alternating) species, the most specialised.

The oak apple gall wasp Biorhiza pallida

Oak apples are amongst the most familiar and best known of oak galls (Fig. 145a). The oak apple is the sexual gall and develops from a leaf bud in spring. The asexual gall (Fig. 145b) occurs on shallow roots and develops in summer and autumn. Both galls occur only on white oaks (in the section Quercus, see Box 9.1), which include pedunculate and sessile oak (Q. robur and Q. petraea) in Britain and several more species in mainland Europe. B. pallida is common and widespread wherever these oaks grow in Europe and is found also in North Africa bordering the Mediterranean and eastwards into Turkey and Iran. It occurs throughout Britain, as far north as Sutherland and Ross-shire.

The structure of the galls is shown in Fig. 146. The oak apple is multichambered, with several inner galls enclosed in a cortex of parenchyma cells and air spaces. The cortex is spongy in the young gall and later hardens as its cells become impregnated with tannins. The epidermis around the young gall has no cuticle or stomata and is soft like the skin of a mushroom; later it is parchment-like. The root gall is simpler, sometimes with one chamber but often with several when neighbouring galls coalesce. When mature, the outer gall is hard and rich in tannins, which deter attack by predators and pathogenic fungi during its long life underground (see below).

FIG 145. Younger and older (a) oak apples (left, top and above) and (b) root galls (right, top and above) of Biorhiza pallida on pedunculate oak Quercus robur. ((a) Robert Cameron, (b) Michael Chinery)

FIG 146. Sections through mature galls of (a) the oak apple and (b) the root galls of Biorhiza pallida (after Redfern & Askew, 1998). The scale line indicates 5 mm.

The oak apple develops after an asexual female has laid a batch of eggs into a dormant leaf bud. She is wingless and emerges from a root gall in December or January; she climbs the trunk of an oak tree, probably the one her root gall developed on. She searches for a suitable bud and investigates it thoroughly before accepting it or moving on to check another bud. Then, positioning herself head downwards (Fig. 147), she drills repeatedly into the bud with her ovipositor, first preparing many channels and then laying many eggs (100 or more is not unusual), each into a channel with its tail extending up it. Her behaviour during oviposition may vary a bit; some observations suggest that the ovipositor cuts a horizontal slit across the bud almost decapitating it, and that the eggs are laid in a cluster on the cut surface of the lower half. The process of checking the bud and laying the eggs can take three hours, and all the while the female is exposed to high winds, cold and rain in the leafless canopy in the middle of winter. The eggs remain dormant until early spring, when they hatch and start the chain of events that results in the oak apple. As in other cynipid galls, a swelling develops under each egg as soon as it is laid. When the larvae hatch and burrow into these swellings, nutritive tissue develops around each separately, the inner gall (Fig. 138) develops, and vascular strands grow to link the nutritive layer with the vascular tissues of the bud (so establishing a nutrient sink that will ensure the larva grows to maturity). The gall is fully developed in May or early June. At this stage the larvae are small, surrounded by nutritive tissue about three cells thick and by storage nutritive cells rich in starch. The larvae now feed actively and grow rapidly, destroying the inner cells, which are successively replaced by the cells of the storage layer. In June, the larvae are full-grown and the nutritive layers eaten, and the larva fills its chamber, now lined by the woody shell. Most oak apples drop to the ground in June, the larvae pupate and adult males and females chew their way out in late June or July through individual tunnels, leaving the gall peppered with holes. A few galls remain on the tree and their dried-up remains may survive there through the next winter. Males usually emerge a day or two before the females and mate with them as soon as they appear. The sexual female is smaller than her mother and, like the males, is fully winged. Wingedness varies geographically, though–in Germany, sexual females may have full-sized wings, reduced wings or no wings.

FIG 147. Asexual female of Biorhiza pallida laying eggs into a leaf bud of oak (after Redfern & Askew, 1998). The scale line indicates 2 mm.

In July or August, the mated female flies down to the ground and finds a thin rootlet exposed on the surface or buried a few centimetres. She lays an egg deep into it, positioned in the cambium layer between the phloem and the woody xylem and, as in the oak apple, this causes a pad of cells to develop and a cavity. In September, the new gall breaks through the outer layers of the root and appears on the surface as a soft, pale yellow or pinkish cushion, 2 mm across. Growth slows down in the winter and stops in harsh weather; it resumes the following spring and the gall is full-grown in June. A single root gall when mature is 10–15 mm across, spherical and light brown in colour. If coalesced with others, which happens when eggs are laid close together in the rootlet, the shape of the galls may be distorted (Fig. 145b). After the gall has become full-sized, the larva grows rapidly, eats all of the nutritive layers and is full-grown by the autumn. Some larvae pupate and their adults emerge in December; others sit in their galls for a further year or two, emerging in December along with individuals one or two years younger. These are all asexual, wingless females, and they climb the tree trunks to lay eggs in dormant buds and to start the cycle again. So, the complete lifecycle takes a minimum of two years, and sometimes four.

The sexual arrangements of B. pallida are complicated, with different females producing different types of offspring (see also Box 9.1). All individuals emerging from one oak apple are usually all males or all females and until recently were thought to be siblings, all the offspring of one asexual mother (either an androphore, producing sons only, or a gynephore producing daughters). But some oak apples (perhaps 25 per cent) produce offspring of both sexes, though often biased to one or the other. So, there could be a third type of asexual mother, capable of producing sons as well as daughters (a ‘gynandrophore’). Alternatively, perhaps an oak apple is produced by more than one mother–and this multiple founding could occur in oak apples whether they give rise to offspring of one or of both sexes. Detailed studies tracing the genetic fingerprints of mothers and their offspring have shown that both of these ideas occur: there are three types of asexual female and oak apples are often the product of more than one female. Is multiple founding the chance outcome of asexual females laying into the same buds, perhaps due to a shortage of suitable buds? Or do females purposely lay their eggs into the same buds because this confers some sort of advantage to their offspring? The answer to the second question is ‘yes’, and the first scenario may also occur in some places and in some years. A bud containing many eggs develops into a larger gall and this gives the larvae advantages: (i) Those in chambers deep in a large gall are more likely to escape parasitism–the ovipositors of many parasitoids are not long enough to reach them. (ii) Females emerging from galls containing both sexes will find males waiting for them, so they do not have to go through the chancy process of searching for a mate. A large gall would also be produced if one female laid all of her eggs into one bud. This would be highly risky, though, as a chance event could destroy the bud and all of her offspring with it. Laying eggs in small batches into buds already containing eggs spreads the risk and provides the advantages of a large gall to the larvae. The tip of the ovipositor probably can detect the presence of eggs, at least of the same species. Sometimes chimeras occur, i.e. galls of two species form in the same bud and fuse together. This is not advantageous to either species and usually no offspring survive–it arises because females probably cannot detect eggs of other species. A final complication in B. pallida is that some sexual females produce offspring without mating (like some individuals of the sycamore gall wasp Pediaspis aceris). All of these are diploid asexual daughters and not haploid males (thelytokous parthenogenesis as in Diplolepis rosae rather than the arrhenotoky of other Hymenoptera).

The oak pea gall wasp Cynips divisa

Cynips divisa causes pea galls on the underside of oak leaves in summer (the asexual generation) and red wart galls in leaf buds in spring (the sexual generation, which are not always red; Fig. 148b). It attacks pedunculate and sessile oaks (Quercus robur and Q. petraea) and other white oaks (in the section Quercus, see Box 9.1) in mainland Europe. The galls are common and widespread over most of Europe and North Africa and, in Britain, they occur from the Isle of Wight to northern Scotland. Abundance is patchy, however, and it is clear that some host trees are much more susceptible to galling than others. Like many oak galls, pea galls are more abundant on young shrub-sized trees than large mature trees. The Cynips species in Britain (there are five: C. divisa, C. agama, C. disticha, C. quercusfolii, C. longiventris) are often attracted to the same trees, too (and this is probably also true for the oak spangle galls, see below). Abundance of C. divisa also fluctuates dramatically over time–in some years, each galled leaf might carry 25 or more pea galls, while in other years, it is difficult to find a single gall on the same tree.

The asexual pea galls are more familiar and are better known than the red wart galls. Their structure is relatively simple. The inner gall is standard with a woody shell enclosing the nutritive layers, and the outer gall is hard and woody, forming a thick resistant wall around the larva. In late August or September, the asexual female emerges from her pupal skin and sits inside the gall until her eggs mature (on average, she has 78 eggs). She chews her way out between November and January from a gall that has fallen to the ground still attached to its leaf, and lays eggs into dormant buds. She is fully winged, unlike the asexual female of B. pallida, which is active at the same time. In the spring, the buds develop rapidly into the red wart galls. These galls are slender cylinders with rounded ends, 6 mm long, in a bud or at the end of a vein on the edge of a young leaf. Like many small bud galls on oak, the gall structure is not much more than a single inner gall. The adult, either a sexual female or a male, emerges in May. After mating, the female lays eggs into the veins of expanding leaves, and the pea galls appear in mid-June, when most leaves are fully expanded. Pea galls occur on the main side veins of the leaf and on the tip of the midrib, where it is similar to the side veins. The galls tend to cluster in the middle of the leaf and on the proximal half of the veins (i.e. near the midrib)–the female carefully chooses the larger leaves and the spot where her offspring are most likely to survive. When numbers are high, three or four pea galls may occur on a single vein, and those near the midrib may be twice as large as the rest. Larvae in large galls are more likely to survive than in small galls, because attack by inquilines, parasitoids and fungi is more common in small galls (see Chapter 10).

FIG 148. (a) Pea galls (left) and (b) red wart gall (right) of Cynips divisa on pedunculate oak Quercus robur. (Michael Chinery, Gyuri Csóka)

The oak spangle galls, Neuroterus species

There are four kinds of spangle galls in Britain, caused by Neuroterus quercusbaccarum, N. numismalis, N. albipes and N. tricolor. The first two, the common and silk button spangles, are usually very common, while the smooth spangle N. albipes is less so and the cupped spangle N. tricolor is often rare. The common names refer to the galls of the asexual generation, found on the underside of oak leaves. Their usual hosts in Britain are pedunculate and sessile oak (Quercus robur and Q. petraea) and other white oaks (section Quercus; Box 9.1) in Europe, but they have been found on red oaks (section Lobatae) imported from North America and grown in parks and gardens. They are distributed throughout the Western Palaearctic, from Scandinavia south to North Africa and from Ireland eastwards to Iran. In Britain, the three commoner species occur wherever the host trees grow, in Scotland as far north as Sutherland, Ross-shire and the Outer Hebrides. N. tricolor may be less widespread. The spangle galls are shown in Fig. 149 and the sexual galls in Fig. 150. The descriptions that follow refer mainly to the common and silk button species, which are better known than the other two.

FIG 149. Asexual spangle galls of (a) Neuroterus quercusbaccarum; (b) N. numismalis; (c) N. albipes; and (d) N. tricolor. (Michael Chinery)

FIG 150. Sexual galls of Neuroterus: (a) N. quercusbaccarum (also occurs on leaves), (b) N. numismalis, (c) N. albipes and (d) N. tricolor. (Michael Chinery [a, b & d], Gyuri Csóka [c])

The structure of the sexual galls when mature is simpler than the normal cynipid pattern. The currant gall of N. quercusbaccarum and the blister gall of N. numismalis lack a woody layer between the nutritive tissues of the inner gall and the cortex of the outer gall; the inner gall does not have its usual woody shell. The bulk of the currant gall is made up of a thick succulent cortex, which consists of large parenchyma cells filled with water. Both of these galls develop on the young leaves as soon as they appear in spring and grow rapidly. The currant gall occurs on the catkins, too, and several on one catkin look remarkably like a string of redcurrants. The lifecycles of N. quercusbaccarum and N. numismalis are similar. Most asexual females emerge from their spangle galls in February and March, fly up into the canopy of an oak tree and lay eggs on to the surface of leaves or catkins inside buds that are beginning to swell. As in other cynipids, a cavity forms in a swelling pad and the larva crawls into it. Nutritive cells form wherever the larva feeds–all around the chamber in the currant gall so that a spherical layer of nutritive tissue develops, and in a horizontal plane in N. numismalis, fitting inside its blister. Vascular strands develop in the cortex to supply the nutritive cells and link with the veins of the leaf or catkin. The galls are full-grown in April and the larvae then grow rapidly. They pupate in the gall in May and males and females emerge in late May and June. They mate and the females lay eggs into the blade on the underside of a leaf, each one in a small cavity next to a vein, where the spangle galls develop.

FIG 151. Development of the spangle gall of Neuroterus quercusbaccarum: at (a) 12 days, (b) 3 weeks, (c) 4 weeks, (d) 5 weeks, (e) 7 weeks, and (f) in October before it falls off the leaf; half of the gall is shown in (f) (modified after Rohfritsch, in Shorthouse & Rohfritsch, 1992). Larva (black); nutritive tissue (stippled); woody tissue (hatched); vascular tissue (open circles and dashed lines). (See text for further explanation.)

Fig. 151 summarises the development of the common spangle gall of N. quercusbaccarum. This takes more than four months, in contrast to the rapid development of the currant gall. Early development follows the usual cynipid pattern. Soon after the egg is laid, the pad and cavity develop (a), and the larva, which hatches after three weeks, crawls into this (b). After another week (c), nutritive cells have developed near the head of the larva, supplied by a new vascular strand linked to a nearby vein, and the pad has broken through the lower epidermis of the leaf and is developing the clusters of red hairs that are characteristic of this gall. Five weeks after oviposition (d), a layer of woody cells has started to form beneath the nutritive tissue and the young spangle is extending laterally, due to a new meristem around its margin. The disc shape that is characteristic of all spangle galls results from the feeding pattern of the larva; it circles in its chamber but only in a horizontal plane. After two further weeks (e), nutritive tissue encircles the chamber, and a sclerenchyma plate has developed above it as well as below and, in contrast to most cynipid galls, these plates remain distinct and do not join into a continuous woody shell. In October, just before it falls off its leaf (f), the spangle is a flattened disc. Unlike most galls, the spangles continue to develop and the larvae to grow after they have fallen to the ground; if they remain attached to the leaf on the tree, the larvae will not survive. The spangles fall before most of the oak leaves and are protected from drying out and from frosts by a blanket of leaves. In the gall, the remaining nutritive cells and the parenchyma cells outside them extend in height, changing the shape of the spangle–it deepens and becomes fat. This growth requires nutrients, which come from starch stored in the cortex. The larva finishes off the enlarged nutritive cells and then overwinters in the gall, protected by tannins in the gall cortex. The tufts of hairs and thick cuticle on the outside of the spangle, as well as the blanket of leaves, all help to prevent the gall from drying out. Without the unique double disc of woody cells that allows the expansion of the spangles on the ground, the larvae would not survive.

The timing of development varies slightly between the four spangle species. The asexual females of N. albipes, and most individuals of N. quercusbaccarum, emerge in late winter and lay eggs in February and March. N. numismalis oviposits in March and some N. quercusbaccarum in April. N. tricolor females are not active until April and May. The sexual females oviposit in the same order too: N. albipes in May, slightly earlier than N. numismalis and N. quercusbaccarum, and N. tricolor in June and July, the only species late enough to cause spangles on the second flush of leaves, the lammas leaves.

The distribution of the galls is notoriously patchy. Some trees are heavily galled by both generations year after year, while others nearby are avoided. This may be due to the timing of bud burst, which varies considerably in different trees. If bud burst regularly coincides with emergence of the asexual adults in spring, the tree is likely to be heavily galled. Some trees may be genetically resistant too. Sometimes the spangle galls are so common that they cover the entire underside of leaves and seem to be growing on top of each other. Do they compete for space and nutrients? Possible competition is reduced because each spangle gall species tends to specialise in a particular area of both the tree and the leaf. N. numismalis is commonest at the tops of trees and near the tip of leaves, N. albipes prefers leaves on branches near the ground and the basal third of the leaf, and N. quercusbaccarum tends to be intermediate both on the tree and on the leaves. This pattern may vary if one species is much more common than another, and age of the trees and their geographical distribution may affect the pattern too.

The lifecycle and sexual arrangements of N. quercusbaccarum are summarised in Fig. 137 (in Box 9.1) and are typical of the other spangle gall wasps and of other advanced cynipids, i.e. Cynips, Neuroterus and most Andricus species. The sexual females as well as the asexual females are of two types. The asexual offspring of one sexual mother that emerged from a currant gall are either androphores (producing sons) or gynephores (producing daughters). She cannot produce both and which type she produces probably depends on the type of male she mated with. The males can mate with many females but the females mate only once. In these more specialised gall wasps males are essential and make up at least 40 per cent of the sexual generation. There are no gynandrophores as in Biorhiza pallida or Pediaspis aceris.

A selection of Andricus galls

Andricus is the largest genus of oak gall wasps with 28 species recorded in Britain (and more in mainland Europe). Some species are thought to have evolved earlier than others: A. curvator and A. foecundatrix (previously A. fecundator) are examples of basal species, while the heteroecious species (those alternating between two oak species), e.g. A. kollari and A. quercuscalicis, are more specialised.

The sexual gall of A. curvator (Fig. 152a), which distorts the leaves of pedunculate and sessile oak (Quercus robur and Q. petraea) in spring, is widespread and common. When full-grown in May and June, it is thin-walled with a large cavity, and the larva lives inside a small spherical inner gall (Fig. 153a). The gall when young consists of the usual concentric zones of tissues (see Fig. 138, Chapter 9). The sclerenchyma cells of the woody layer are alive while the gall is growing, so that there is no barrier to the transport of nutrients to the nutritive cells and the larva within. The outer gall at first consists of a thick cortex of large parenchyma cells with vascular strands that deliver nutrients to the nutritive cells. The gall grows rapidly, and the inner gall soon reaches its full size (Fig. 153b [i]). The outer gall continues to grow, forcing the cortex cells next to the woody shell to pull away and split apart from it so that an air space appears around the inner gall and increases in size (Fig. 153b [ii and iii]). The larva remains small while the zones of tissues develop and then grows rapidly as the air space expands (Fig. 153b [iv]). Males and females emerge from these galls in late May and June and mate, and the females lay eggs into dormant buds. The asexual gall develops slowly and is full-grown in autumn (September–November). This gall is inconspicuous, brown and woody, and only 3 mm long. At first it is enclosed in the brown scales of the bud; when full-grown, it pushes out of the scales and can be recognised from other small bud galls by its white collar (Fig. 154). Inside, the larval chamber takes up most of the space. The larva overwinters, pupates, and the asexual female emerges in March or April, ready to lay eggs into leaf buds as they are beginning to swell.

FIG 152. (left to right) Basal Andricus species: (a) A. curvator, two sexual galls at the base of the right-hand leaf; (b) A. foecundatrix, the asexual artichoke gall (Michael Chinery).

FIG 153. Sections through the sexual gall of Andricus curvator. (a) The mature gall and a section (from Redfern et al., 2002, with permission); (b) stages (i–iv) in development of the inner gall and air space (modified after Meyer & Maresquelle, 1983)–the gall is developing while the larva (not shown) remains small; in (ii), the air space is starting to develop; (iv) the nutritive cells have been eaten by the larva and most of the outer gall is a large air space. Nutritive tissue (finely stippled); storage nutritive tissue (densely stippled); woody shell (black); midrib vein (open circle).

FIG 154. Asexual gall of Andricus curvator in an oak bud (Quercus robur or Q. petraea), one specimen with bud scales partly removed and the other with scales only around the base; in both, the white collar is clear (from Redfern et al., 2002, with permission).

In Andricus foecundatrix, the oak artichoke gall (the asexual gall, Fig. 152b) is more familiar than the sexual gall. The sexual gall is small and hairy (Fig. 155); it grows rapidly on a catkin in spring and when full-grown is only 2 mm long. Galled catkins persist longer on the tree than normal catkins, ensuring that the gall is supplied with nutrients for long enough for the larva to develop. The larva grows quickly and pupates in May and June, and the adult male or female emerges later in June. The artichoke gall develops in a bud during the summer. When full-grown the gall is large, up to 30 x 20 mm, a cluster of enlarged bud scales. These enclose an ovoid inner gall, which replaces the meristem of the bud. The inner gall is 6–9 mm long, shiny brown in colour, and contains the single larva. The larva is full-grown in August. Then, the scales dry out and shrink, and push out the inner gall, which falls to the ground. It remains buried in leaf litter over one or two winters, the larva inside pupates in the spring, and the asexual female emerges in March or April of the first or second year following the shedding of the inner gall. The old outer artichoke galls can persist on the tree for years.

FIG 155. Sexual gall of Andricus foecundatrix on an oak catkin (Quercus robur or Q. petraea) (from Redfern et al., 2002, with permission).

The most specialised gall wasps are the heteroecious species. None of these is native in Britain. All require an oak in the section Cerris (see Box 9.1) for one of their generations and they spread to Britain only after the Turkey oak Quercus cerris arrived–this species was introduced to parks and gardens as an ornamental tree in 1735. Only two cynipid genera, Andricus and Callirhytis, contain heteroecious species and it is likely that heteroecy evolved only once (but see Chapter 11 for further discussion). In Britain, we have eight species, all Andricus, whose asexual galls occur on pedunculate and sessile oaks and their sexual galls on Turkey oak. There are four additional species known only from their asexual galls. These may have lost the sexual stage of the lifecycle, or their sexual galls may await discovery. Andricus kollari, the marble gall, was the first heteroecious species to arrive in Britain, introduced intentionally from the eastern Mediterranean in the 1830s; its high tannin content made it valuable for tanning leather, dyeing cloth and ink making (see Chapter 13). The other species are all accidental introductions to Britain: the knopper gall Andricus quercuscalicis arrived in the late 1950s, the cola nut A. lignicolus and A. corruptrix were first certainly recorded about 1970, and the hedgehog gall of A. lucidus and A. grossulariae have appeared more recently. The latest arrivals are the ram’s horn gall A. aries in 1997 and A. gemmeus in 2008, and there may be more in the future. A. kollari and A. quercuscalicis are the best known of these species and their galls and lifecycles are described next.

FIG 156. Asexual marble galls of Andricus kollari on pedunculate oak Quercus robur, which often grow in clusters. (Michael Chinery)

FIG 157. Section through a mature marble gall of Andricus kollari (from Redfern et al., 2002, with permission).

The oak marble gall, the asexual gall of Andricus kollari (Fig. 156), is probably the most familiar of all galls on any plant. A typical gall is spherical and smooth, and often has small bumps on its surface (caused by a coelomycete fungus). Marbles of odd shapes are common, stunted or fused together, or distorted and enlarged, due perhaps to the early death of the larva, or to aberrations during gall development, or to the presence of inquilines (see Chapter 10). Fig. 157 shows a section through a typical gall. It is bright green when young and later brown, hard and woody, and old galls may persist on the tree for years. Its inner gall is similar to that of other cynipid galls, with nutritive tissues enclosed by a woody shell. The cortex of the outer gall includes a fine network of cells with small air spaces. When young, this is slightly spongy; when older, it dries out and becomes friable, and fuses with the woody shell of the inner gall. Beneath the epidermis the cortex cells are more closely packed and are rich in tannins. Tannins protect the insect inside against harsh weather and from the invasion of moulds, but are probably not very effective against attack by parasitoids and inquilines. The sexual galls on Turkey oak are tiny bud galls, inconspicuous, yellowish or light brown in colour, and they often occur in clusters of two or more (Fig. 158). Like other small bud galls, they consist of the inner gall only with a thin woody shell, and the larva takes up most of the space inside.

FIG 158. Sexual galls of Andricus kollari on Turkey oak Quercus cerris (from Redfern et al., 2002, with permission).

Andricus kollari is unusual in that its asexual gall develops more rapidly than its sexual gall. Males and females emerge in May and June from the bud galls on Turkey oak, they mate and the females fly to pedunculate or sessile oaks. They lay eggs into leaf buds, which develop into the marble galls. The females prefer swelling buds that have not yet burst into leaf, and marble galls are common on young and coppiced trees that are often late to flush their leaves. The larvae are fully fed in August when they pupate, and the asexual adult females chew their way out with most emerging in September. These fly to a Turkey oak, which may be some distance away, and deposit their eggs between the tiny immature leaves inside dormant buds (Fig. 159). The sexual galls develop slowly during the winter and become apparent when they are full-sized in March or April. The larvae now grow rapidly as temperatures increase in the spring, they pupate, and males and females emerge in May and June ready to start the next asexual generation. The lifecycle of A. kollari usually takes a year to complete, at least in mainland Europe. In Britain, a proportion of most populations takes two years to develop–the females remain inside their marble galls over the winter, emerge in the following May or June, and lay eggs into dormant buds of Turkey oak, buds that would produce the following year’s leaves. It is now too late for the sexual galls to develop and so the eggs and the buds remain dormant until the autumn. Then they develop slowly during the winter along with the next cohort of sexual galls, and produce males and females in May and June of the following year. The proportion of individuals with a two-year lifecycle increases from south to north in Britain, with most or all in northern Scotland (Sutherland and Ross-shire) developing over two years. If none develop in one year, the populations of alternate years would be genetically distinct, unless one-year migrants fly in from further south. In the far north, numbers often fluctuate violently from year to year, and a two-year lifecycle may be partly responsible (discussed further in Chapter 12).

FIG 159. Andricus kollari asexual female ovipositing into a dormant bud of Quercus cerris. (Michael Chinery)

Knopper galls, the asexual galls of Andricus quercuscalicis, are well known and can be as common as marble galls even though the species only arrived in Britain in the 1950s. It spread rapidly and now is found throughout England and Wales wherever pedunculate oak Quercus robur occurs (it is very rare on sessile oak Q. petraea, although it occurs on the hybrid between them). Knopper galls are found on acorns (Fig. 160) in summer and autumn. They are green and sticky when young with a thick wall that becomes brown and woody later; inside is a large air space and an ovoid inner gall (Fig. 161). The sexual galls (Fig. 162) occur on the male catkins of Turkey oak Q. cerris in spring, and are tiny, simple, thin-walled galls only 1–2.5 mm long. Each gall replaces two to four anthers in the catkin, and one catkin may carry up to 20 galls. The distribution of these galls varies greatly–the catkins on one tree may be smothered in galls year after year while its neighbour is never galled. If a catkin is galled, its stalk is thickened and its life span prolonged, although beyond the gall the catkin withers. Normal catkins of Turkey oak survive for about 12 days. The tiny galls of A. quercuscalicis require about two weeks to develop, prolonging catkin life only slightly, but other Andricus species with larger galls may extend the life of a catkin by two or three months. The prolonged life ensures that nutrients from the tree supply the larva until it is fully fed. In May, the sexual adults emerge from these tiny galls. The males emerge first and sit on galls containing females, waiting for them to bite their way out. They battle with other males in order to secure the best position to mate with the females. The mated females fly to pedunculate oak and, in late May, lay eggs into newly pollinated female flowers, placing each egg precisely between the developing acorn and its cup. Early-developing trees whose leaves flush early have the most knopper galls. In these trees, most female flowers have opened and been pollinated and are in the right state for galling when the sexual females arrive. The buds of late-burst trees are still closed and are ignored by the ovipositing females. The knopper gall develops over the summer. In September, the galls with their larvae fall to the ground before the ungalled acorns and overwinter in the litter. Most asexual females emerge the following February and March, fly to a Turkey oak and lay eggs into male flower buds to start the cycle again. A proportion of females remains in their knoppers in the leaf litter for up to three years after they fell in September, i.e. they join later cohorts and emerge with them in February or March. In Britain, most (70–80 per cent) emerge in their first year; this proportion decreases eastwards across Europe, with less than 20 per cent emerging in their first year in eastern Austria and Slovenia.

FIG 160. Knopper galls of Andricus quercuscalicis on pedunculate oak Quercus robur. (Michael Chinery)

FIG 161. A mature knopper gall of Andricus quercuscalicis in section, showing the large internal cavity around the ovoid inner gall (after Schönrogge et al., in Williams, 1994).

FIG 162. Sexual galls of Andricus quercuscalicis on a catkin of Turkey oak Quercus cerris (from Redfern et al., 2002, with permission).

A second invading heteroecious species, Andricus lucidus, forms hedgehog galls on the edge of an acorn cup (Fig. 163a). Before knopper galls arrived, this spot had not been exploited by any Andricus species in Britain and provided a resource that the invaders could exploit. The hedgehog gall, the asexual gall, was first recorded in Britain early in the twentieth century but, later, this was considered to be an error. Then, in the early 1990s, it was found in Regent’s Park in London–probably a new, accidental, introduction. It is spreading slowly but, after nearly 20-years, it is still confined to southeast and central southern England. Hedgehog galls also occur in terminal buds and are found on both pedunculate and sessile oaks and other white oaks (Box 9.1). When mature, the gall is spherical, 5–10 mm in diameter, and covered with thick spines, each ending in a knob. It is pale green, sometimes flushed with red, and is sticky. Old galls, which can remain on the tree for years, become dark brown or black and lose their spines. The young gall appears in mid-July as a small, sticky, reddish bulge. As it grows, the surface becomes fissured and the spines develop. Inside, the cortex of the gall is pale yellow and soft at first, with several chambers each containing a larva. When mature, it is dry and woody and loses its red colour and stickiness. On acorns, the galls with their fully fed larvae fall to the ground in autumn; galls on buds remain on the tree. The asexual females emerge in the spring, fly to a Turkey oak and lay eggs into buds containing catkins. The sexual gall (Fig. 163b) is a cluster of small galls with many folds and indentations, greenish yellow in colour and soft when young, becoming bright red and woody as it matures. It causes the catkin stalk to thicken and prolongs its life by several months–galled catkins remain on the tree until July or August.

FIG 163. (a) The asexual hedgehog gall of Andricus lucidus on an acorn of oak Quercus sp. (Michael Chinery); (b) the sexual gall on Turkey oak Quercus cerris. (Gyuri Csóka)

PART FOUR

Galls in Context

CHAPTER 10

Interactions and Adaptations

GALLS ARE DISCRETE LITTLE UNITS, rather distinct from the rest of the plant they are found on. Their occupants have a more passive and enclosed way of life than their free-living relatives. Hence, while galls and their causers are part of a wider ecological community, they can be the centre of a subcommunity of specialists that depends on them. The gall itself, made up of nutritious and often succulent plant tissue, attracts herbivores. The gall causer has enemies that attack it, and the whole complex attracts detritus feeders that live off debris and decaying plant tissue. Collection and dissection of galls, and the rearing and identification of their occupants, can reveal almost self-contained microcosms of ecological communities.

Some of the systems have been studied in detail, exploring the interactions involved and attempting to answer some fundamental ecological and evolutionary questions. What determines the distribution and abundance of gall-causing animals? Is it the availability or quality of the host plant (‘bottom-up’ effects) or the attacks by predators and parasitoids (‘top-down’)?* What, if any, are the advantages of living in a gall rather than in the open? How stable and balanced are these miniature communities?

These interactions have been explored in detail for only a few kinds of gall, all of which are caused by mites or insects. We know very little about communities associated with the galls of bacteria, fungi and nematodes. The patterns revealed vary from very simple to some very complex ones involving many species. While some of the species concerned are specialists, completely dependent on the gall or its occupants, there are others for which galls and their contents meet only a small part of their requirements for food or shelter.

BOX 10.1. Definitions and explanations of feeding categories and the food web arrow symbols used in this chapter

In the food web diagrams, the gall causer is enclosed in a box with a double border.

Carnivore: A general term for animals that feed on other living animals. It can be subdivided into the several more or less distinct categories below.

Parasitoid: An insect parasite whose larva feeds on a single host individual, usually another insect, eventually killing it (a true parasite does not kill its host); in galls, all parasitoids belong to the order Hymenoptera (ants, bees and wasps).

Endoparasitoid (): A parasitoid that lays its eggs inside the host and whose larva feeds within it.

Ectoparasitoid (): A parasitoid that lays its eggs on the outside of the host or nearby in the gall chamber; the larva feeds on the host from the outside.

Hyperparasitoid: A parasitoid of a parasitoid; sometimes it parasitises other individuals of its own species, i.e. it can be a cannibal. (In galls, ‘hyperparasitoids’ are actually

pseudohyperparasitoids because the primary host is killed before the hyperparasitoid attacks the parasitoid.)

Pathogen (): A bacterium or fungus that parasitises an animal or plant.

Predator (): A carnivore that kills its prey and feeds on several individuals before it is fully fed.

Herbivore (): A general term for an animal that feeds on living plant tissue; many specialise on a particular plant part (e.g. roots, stems or leaves) and some, particularly moth caterpillars, also feed on galls, burrowing in from the outside. In the context of galls, there are two specialised categories of herbivore in addition to the gall causer itself.

Inquiline (): A specialised herbivore that feeds on the ‘home’ of the gall causer; most inquilines develop inside the gall, feeding from within, and leave as fully fed larvae or as adults. Many species are lethal inquilines, i.e. they kill the gall causer either directly or indirectly.

Cleptoparasite (or kleptoparasite): An inquiline that usurps the gall causer, driving it out of its gall and often killing it, and taking over the gall; it is a term applied particularly to thrips galls, containing many individuals.

Detritivore: An animal that feeds on dead and decaying animals and plants, or on the fungi and bacteria that grow on the decaying remains.

None of these feeding categories is totally exclusive–they can merge one into another so that the boundaries between them become blurred. For example, an inquiline feeding on gall tissue may break into a chamber and eat the larva within, so acting as an ectoparasitoid; or it may open several chambers in a multichambered gall and eat the occupants one after another, and so behave more as a predator. The role of an enemy can change over time too: it may start as an ectoparasitoid when young, becoming inquilinous as it matures (or vice versa).

In the food webs, the main role of each enemy is shown; its role is elaborated and modified in the accompanying text.

Two further terms are applied to parasitoids:

Koinobiont: A parasitoid that does not kill its host at oviposition; the host larva continues to grow and usually develops until it is fully fed, when the parasitoid kills and feeds on it. Because the host larva (the gall causer) continues to feed, the gall usually grows to its full size.

Idiobiont: A parasitoid that kills or paralyses its host when ovipositing so that it does not develop further. If the gall is not full-grown when attacked, it will remain undersized.

The results of some of these studies are displayed here as food webs, centred on the species causing the gall. The food web diagrams use standard terms and symbols (see Box 10.1). They concentrate on the major and specific interactions and cannot show all the known links and species. Many galls and their occupants are attacked by birds and mammals, either herbivores that accidentally kill the occupants or insectivores destroying the gall to extract occupants. None of these birds and mammals are dependent on galls, and they are not shown in the webs.

Some of the species shown have multiple effects. Some parasitoids are polyphagous, and parasitise other parasitoids (i.e. they are ‘hyperparasitoids’) as well as the gall causer or inquilines. Some parasitoids also consume gall tissue (and so are also inquilines). The webs depict just one role for each species, and other roles are described in the text. Herbivorous moth caterpillars, which attack the gall from the outside, have their own parasitoids; these are not shown.

After the gall causer and any inquilines and parasitoids have left the gall, ‘successori’ or squatters often invade it and live inside the old gall. They may just shelter there, attracted by the small dark space, or they may feed on detritus that accumulates or on fungi and bacteria that colonise the old gall. Some species are carnivores and feed on other squatters, and any of them may have their own suites of parasitoids. Some old galls are used as nest sites by solitary bees and wasps; they pack their brood cells inside, one above the other or side-by-side to fill the available space. A wide range of animals may be successori: psylloids, scales, aphids, thrips, springtails, fly maggots, beetles, ants, spiders and mites and, if the gall becomes buried in litter and soil, woodlice, millipedes, snails, slugs and worms. None of these are included in the food webs.

The various subcommunities that have been studied are grouped by the kind of gall-causing animal at their centre. Galls caused by mites and by some aphids, adelgids, gall midges, tephritids and sawflies often support rather simple subcommunities. Others, especially those of cynipids, can become very complex; in many of these, more connections and interactions remain to be discovered. The end of the chapter uses information from some of the case studies given to look for some general rules. Later chapters explore some questions, such as adaptation, in more detail.

ERIOPHYOID MITE GALLS

Most mite gall communities are simple, whether in erinea, pustules, pouches or big buds. Because of the small size of the mites, they are not parasitised–a parasitic wasp cannot survive on just one individual eriophyoid mite. The Eulophidae is a wasp family that is often parasitic in insect galls, but in mite big buds they are predators, feeding on several or many individual mites in their lifetime. The predator larvae crawl about as they feed and they pupate inside the gall, surrounded by the skins of mites they have sucked dry. A single larva of Aprostocetus eriophyes is found in big buds on yew Taxus baccata (Fig. 164) and blackcurrant Ribes nigrum, the galls caused by Cecidophyopsis psilaspis and C. ribis respectively, while the larger big bud of Phytoptus avellanae on hazel Corylus avellana can support two (these galls are described in Chapter 6, Big Buds). A. eriophyes can survive on several species of gall mite even though they occur on unrelated plants; the female clearly recognises and is attracted by the shape and structure of the big bud itself. A related eulophid, Quadrastichus sajoi, feeds on mites of Acalitus phloeocoptes, which causes hard pustules in the bark of plum and blackthorn trees Prunus spp. It invades the galls in their second season when they are larger and contain more mites, and by the time a larva is full-grown there will be few mites left alive in the gall.

FIG 164. Aprostocetus eriophyes in the yew big bud of the mite Cecidophyopsis psilaspis. (a) The gall cut open to reveal the full-grown larva; (b) the pupa (scales = 1 mm) (after Redfern & Askew, 1998).

No true inquilines, feeding on the gall tissue and leaving the gall causer unharmed, are known in eriophyoid galls. But ‘usurping’ species have recently been reported (Jim Amrine, pers. com.). Aceria ulmi is a North American mite that lives on leaves of native elms in the eastern USA. The mite cannot cause its own galls but usurps the small pouch galls of Aceria hindali (a new species, not yet formally described). A. ulmi behaves as a ‘cleptoparasite’ (see below, under thrips galls), overwhelming and killing the gall causer, or driving it out of the gall (which amounts to the same thing). The cleptoparasite is sometimes so common that nearly every gall on the leaf contains it rather than the host mite.

Eriophyoids in big buds, and in erinea and pouches too, are attacked by a variety of less specific predators, active species that roam over the foliage of trees and bushes, entering any gall they encounter–though they have to be able to push past the hairs guarding the mouths of the pouches. Predatory mites of the families Phytoseiidae, Tydeidae and Stigmaeidae are common in the galls of Colomerus vitis on grapevine Vitis vinifera (described in Chapter 4) more or less anywhere vines are grown, and may be effective in keeping the erineum and leaf curl strains of this pest at low levels. They do not have much effect on the bud strain, though. General insect predators include anthocorid bugs and larvae of lacewings and ladybirds, and the predatory larva of the cecidomyiid Lestodiplosis has occasionally been found in big buds on yew (Fig. 165). The simple food web of the yew big bud is shown in Fig. 166, which is probably typical of communities in eriophyoid galls. Parasitic fungi (pathogens) sometimes attack the gall mites too, although how specific they are is unknown. Plasmopara viticola infects Colomerus vitis in erinea, and four species (Verticillium lecanii, Macrosiphoniella sanborni, Hirsutella thompsoni and Metarhizium anisopliae) are pathogens of Cecidophyopsis ribis.

FIG 165. Full-grown larva of the predatory gall midge Lestodiplosis sp. (after Redfern & Askew, 1998). (The head is to the right; scale = 1 mm.)

FIG 166. The food web in big buds of Cecidophyopsis psilaspis on yew Taxus baccata (from Redfern & Askew, 1998, slightly modified). Arrows and symbols are defined in Box 10.1.

Mortality of the eriophyoid mites when inside their galls is probably not severe, in erinea, rolls and folds as well as inside the big buds. This changes in the spring; then the mites disperse from their old galls to new leaves and buds, and are exposed on the surface of leaves and shoots, prey to any passing predator and to adverse weather–they may get blown about and dislodged by wind and rain and large numbers will fail to reach new buds and leaves at a stage when they are suitable for galling. Control of pest species by spraying with pesticides has most effect during dispersal.

APHID AND ADELGID GALLS

The most elaborate of these galls are the closed pouches caused by poplar and pistacia aphids (e.g. Pemphigus species on Populus in Britain and species belonging to the subfamily Fordinae on Pistacia in Israel; see Chapter 5), and the pineapple galls of Adelgidae on spruce Picea (see Chapter 6), and many of these contain large numbers of aphids. But despite the large potential food supply, the communities inside these galls are not complex. They are particularly simple in Adelges galls–there are no known parasitoids and predation is negligible, on the fundatrices as well as on the gallicolae (adelgid lifecycles are described in Chapter 6, Pineapples). The main causes of mortality are of the first instar gallicolae if they have to sit for some time on the gall surface waiting for the aperture of chambers to open, or of the winged stages during dispersal. The communities in poplar and pistacia aphid galls (summarised in Figs 167 and 168) are larger and are remarkably similar, despite occurring far apart geographically. They include parasitoids, inquilines and predators, though relatively few species of each compared to the galls of some other insects. Free-living aphids are much more heavily attacked too, with hundreds of species of parasitoids recorded, far more than in the galls.

FIG 167. Food web for Pemphigus gall aphids in Britain (information from Dunn, 1960; Jones, 2002). Arrows and symbols are defined in Box 10.1.

FIG 168. Food web for pistacia gall aphids in Israel (information from Wool & Manheim, 1986; Wool & Burstein, 1991; Wool & Bar-El, 1995; Wool & Ben-Zvi, 1998; Wool, in Raman et al., 2005).

Parasitoids only attack the aphid fundatrices, transforming them into ‘mummies’: hard, dry, brown aphid bodies that have been consumed from within by the parasite larva, the adult biting its way out through a circular hole. These parasitoids belong to the family Aphidiidae (a family related to the Braconidae, Hymenoptera: Ichneumonoidea) and few species have been recorded. Monoctonia pistaciaecola is widespread across southern Europe to Iraq, and attacks the fundatrix of several pistacia species (e.g. Smynthurodes betae, Aploneura lentisci, Forda marginata), killing perhaps 10–20 per cent. It also attacks Pemphigus fundatrices on poplars in the same area. In Britain, although mummified fundatrices of Pemphigus with emergence holes have been found, the parasitoid species has not been determined. Of course, if the fundatrix dies, no gall develops. In both poplar and pistacia galls, a hyperparasitoid of the fundatrix is found (i.e. a parasitoid of the parasitoid) and belongs to the same chalcid genus Pachyneuron (family Pteromalidae) whether in Britain or Israel.

Predators are more common. Insect predators can do a lot of damage even though most can enter only when the galls are open, i.e. early in the season when the gall is forming or a few months later when the aperture opens to release the winged aphids. Ladybirds attack the newly hatched fundatrix of Pemphigus bursarius as it is attempting to establish itself. Once the gall has closed, the fundatrix is safer and can produce offspring without interruption–unless moth caterpillars attack the gall. Caterpillars are primarily herbivores and chew through the gall wall attracted by the succulent gall tissue, but they often also eat the aphids and may destroy the whole colony. Geometrid, pyralid and gelechiid larvae have been recorded (Figs 167 and 168) and are often parasitised themselves. The caterpillars are not specific to these particular galls; they are free-living, feeding on leaves of poplars, pistacias and other trees, and they enter the galls only when they happen to encounter them. On pistacia, the caterpillars often pupate in the gall, first cutting a hole through the wall and sealing it with a sheet of silk–the pupa later breaks through this, allowing the adult moth to emerge. A large gall, of Geoica wertheimae for example, may contain several pupae, with each adult emerging through a different hole. Although caterpillars of Palumbrina guerrini (Fig. 168) do not eat the aphids, they feed on the inner tissues of the gall so that the aphids starve. Caterpillars often attack the galls before they, the galls, are fully grown; even if some aphids survive these galls are smaller than usual, because there are fewer individuals to stimulate gall growth. Caterpillars of some species actively avoid the galls while feeding on the rest of the leaf–the galls are left hanging by threads, suspended by the veins of skeletonised leaflets.

Caterpillar holes allow the predators to enter the galls, although most do not depend on these holes. In Britain, hoverfly larvae belonging to two species of Heringia (Diptera, family Syrphidae; Fig. 167) specialise on the waxy aphids of Pemphigus and Thecabius, becoming camouflaged by their wax. The larvae are small and brown, 5 mm long when full-grown, and feed on the aphids in August and September before and after the gall apertures have opened. When fully grown, they crawl out of the galls to overwinter in a crevice on the tree; they pupate in April and the adults emerge in May. H. heringi, and less often Pipiza festiva (Fig. 168), also occur in galls of Pemphigus and Baizongia pistaciae and other Fordini in southern Spain and in Israel. There, they apparently pupate inside the galls. In southern Europe, Leucopsis larvae (Diptera, family Chamaemyiidae; Fig. 168) also specialise on aphids in both pistacia and poplar galls. The eggs of L. steinbergi are brown and sausage-shaped, and in May or June are laid near the hairs of gall apertures that have not yet closed. The hatchlings crawl inside, feed on the aphids and, when fully fed, crawl out of the gall when it opens in late summer. They pupate on the ground or drop with the gall when leaves are shed in the autumn. The larvae rarely destroy all of the aphids in a gall; sometimes only a few (e.g. 2–6 per cent of Aploneura lentisci), sometimes up to 45 per cent (e.g. of Forda formicaria and Paracletus cimiciformis). Leucopsis palumbi and L. grunini have more complex lifecycles in southern Europe. Early in the season, the larvae feed on Tetraneura ulmi galls on elms (Ulmus; see Chapter 5, Plant bug pouches), entering through the narrow opening on the underside of the leaf before it closes. Later, the adults emerge when the gall splits and fly to pistacia leaves to lay eggs near Fordini galls. They require galls on the margins of the leaflets and with long slit-like openings through which the maggots can enter, e.g. the galls of A. lentisci, P. cimiciformis and Forda species.

Anthocorid bugs (Hemiptera, family Anthocoridae) are common in poplar and pistacia galls and can inflict major damage, sometimes destroying the whole colony. Anthocoris minki specialises in Pemphigus galls on poplar, but polyphagous species are found too (Fig. 167; e.g. A. nemorum, A. nemoralis, Orius minutus). One or two A. minki bugs enter soon after the gall opens, and can kill the entire contents of the gall before starting to feed. The dead aphids are in a relaxed state but without feeding scars or other injury, and have been killed rapidly and highly efficiently. How are they killed? Perhaps a secretion from the bugs’ stink glands ‘gasses’ them, all at once, the secretion particularly effective in the confined space of the gall. Anthocorid eggs are inserted into plant tissue, probably outside the gall. The first instar crawls inside, kills the aphids and completes its development in one gall. The less specialised species may enter the gall as older nymphs or adults.

Interactions of ants with free-living aphids are well known; they are often mutualistic, with the ants milking the aphids for their honeydew and the aphids getting protection from their enemies in return. This may occur in gall aphids too, with the ants defending the galls from predators. The root-feeding generations of pistacia aphids, which infest grasses but do not cause galls, are sometimes closely associated with the ants. The host grasses grow in or near to ants’ nests and the ants tend the aphids carefully, carrying them deep within the nest in hot dry weather and back to the surface when the rains start. But ant relationships are not always so beneficial–sometimes the ants become predators of the aphids.

Some gall aphids have ‘soldiers’ that drive enemies away, behaviour that also occurs in social wasps, bees, ants and in termites. In Pemphigus spyrothecae (and also P. dorocola in Japan), hoverfly larvae and caterpillars are attacked as they attempt to enter the gall, the soldiers emerging from the gall and stinging them with their stylets and piercing them with their armed hind legs and tarsal claws. Sometimes an intruder is quickly covered with soldiers and all of them drop to the ground. This behaviour is also effective against vertebrate predators. Ceratoglyphina bambusae is a tiny aphid in Taiwan, which forms large galls 9 cm in diameter on Styrax, each gall containing perhaps 100,000 aphids. Its second instars are aggressive soldiers that patrol the surface of the gall, attacking any invertebrate that comes near. If the galls are disturbed from below, by grazing mammals or humans brushing against the branches, large numbers of soldiers drop on to the intruder and inflict painful stings that intensely irritate the skin. Vertebrates are predators too. Tits and chickadees, squirrels and chipmunks readily slice open poplar galls and eat all of their aphids. Bird predation is uncommon on pistacia galls. One observation (Burstein & Wool, 1992) on a single Pistacia palaestina tree in Israel records a pair of great tits Parus major systematically attacking the galls of Paracletus cimiciformis and Forda formicaria. A bird removed a galled leaflet, hopped with it to a larger branch, and held it down with one foot while tearing the gall open and methodically removing every aphid. The tit dropped the spent leaflet before moving on to another, and a litter of empty galls with characteristic jagged tears piled up beneath the tree. But Israeli tits do not often learn this behaviour, it seems.

A final enemy recorded on pistacia gall aphids in Israel is a pathogen, the rust fungus Pileolaria terebinthi (Pileolariaceae: Uredinales; see food web Fig. 168). It can destroy whole galls of Forda formicaria and all of their aphids. Galls already damaged by caterpillars of Alophia are more likely to be attacked–perhaps the caterpillars spread the fungal spores or the fungus may gain access to the gall via caterpillar wounds.

Apart from the stages of the lifecycle that live in the gall, most poplar and pistacia aphids have non-galling stages that live on the roots of herbaceous plants, and these are attacked by a host of other enemies. Pemphigus bursarius migrates to lettuce and other Asteraceae in July, and feeds in dense colonies during the summer. They are attacked by a variety of non-specific predators: ladybirds, hoverfly larvae, anthocorid bugs and pathogenic fungi while they are migrating to and from their summer hosts, and chloropid fly larvae and carabid and staphylinid beetles when on the roots. Mortality can be high, much higher than it is in the galls. If temperature and humidity are high, fungi can destroy whole colonies.

THRIPS GALLS

Most thrips galls are leaf folds and pouches (see Chapter 4, Gall Thrips (Thysanoptera)) and, like aphid galls, they may contain thousands of individuals. No parasitoids have been noted in thrips gall communities, but they may occur. Otherwise, the community is similar to that in aphid galls with predators, inquilines and herbivorous caterpillars, and is more complex than in adelgid galls. Cleptoparasites are common; these are thrips of other species that attack the gall causers, killing them or driving them out, and taking over the gall (an extreme form of inquilinism, see Box 10.1). The gall is a valuable resource, so competition for it can be fierce. If the cleptoparasites are successful, the galls are often enlarged or distorted, as occurs in other galls attacked by inquilines. Unlike the aphids, the gall-causing thrips have more enemies than their free-living relatives.

Fig. 169 shows a generalised food web that applies to many thrips galls. Most galls are permanently open so that insects and mites can come and go, although soldiers belonging to the resident species often attack them (see Box 4.3, Chapter 4). Some predators are specialised, e.g. eulophid wasps, while many are more general, attracted to prey in any small space. These feed on other species in the gall as well as the gall causer. Spiders are opportunists and feed on anything they can catch. They sit on the outside of the gall near its aperture, waiting for thrips and other insects to leave or enter. Moth caterpillars may be predatory although, as in aphid galls, they are mainly attracted by the succulent tissue of the gall. The inquiline thrips mainly feed on the gall tissue too, but are also cleptoparasites. They fight the rightful owners of the gall and often succeed in taking it over–they tend to be larger than the gall-causing species and have soldiers with large stylets and well-armed forelegs. Other insects and mites are ordinary inquilines, feeding on the gall tissues and doing no harm to the founding thrips, and some may specialise in the galls of particular species. Five species of scale insects are found in the leaf rolls of Gynaikothrips ficorum; the scales produce honeydew that attracts ants and these often drive away other insects and mites so benefiting the host and the inquilines.

FIG 169. A generalised food web for thrips galls (details for specific galls are not well known). (Compiled from Mound, in Williams, 1994; Mound, 2005; Crespi et al., 1997.) Arrows and symbols are defined in Box 10.1.

An unusual, woody, thrips gall grows on Casuarina trees (family Casuarinaceae) in dry inland areas of New South Wales and southern Queensland in Australia. The gall is a growth up to 15 cm long on the branches, and increases in size over several years. It is initiated by the fundatrix, a single-winged Iotatubothrips crozieri female, and she produces young that add to the size of the gall as they feed. The gall surface is covered in tiny cracks through which two cleptoparasitic species gain entrance, Thaumatothrips frogatti and Phallothrips houstoni. The usurping thrips are larger than I. crozieri and both produce wingless soldiers that kill the owners with their large toothed forelegs. It takes some time to take over the gall. An invading P. houstoni female crawls to the end of a gall lobe, separates herself by producing a thin cellophane-like partition, and lays five to ten eggs in the enclosure. The eggs hatch into the soldiers and these break through the partition and set about destroying the host thrips. The female continues to lay eggs, producing normal offspring that mature and also reproduce, and the usurpers eventually kill all of the original inhabitants.

TEPHRITID AND CHLOROPID GALLS

In this section, the gall communities of tephritid flies (Diptera: Tephritidae) in stems and flower heads of thistles, knapweeds and other Asteraceae, and of Lipara flies (Diptera: Chloropidae) in the shoot tip of the common reed Phragmites, are described. The adults of these two families are robust flies, quite different from the delicate gall midges (Cecidomyiidae, also Diptera). Regular inhabitants are specialist endoparasitoids, several less specialised ectoparasitoids that sometimes become inquilinous, other inquilines, herbivorous caterpillars and predatory fly maggots. In addition, birds often peck open the galls and extract the larvae, and small mammals feast on them if they fall to the ground in winter. The tephritid stem gall communities are simpler than those in flower heads and both are simpler than the Lipara community in reed shoot tips (Figs 170–4). In Lipara galls, the variety of ichneumonid parasitoids and of inquilines is striking–and two of these inquilines are also gall causers in their own right. The galls of all of these flies are large and probably easy for enemies to recognise; both tephritid and chloropid larvae inside galls are more likely to be parasitised than their free-living relatives (in contrast to galling and free-living mites, aphids and sawflies). Two tephritid stem galls, tephritids in burdock, thistle and knapweed flower heads, and the cigar gall of Lipara lucens are described in more detail.

FIG 170. The food web of Eurosta solidaginis in stems of tall goldenrod Solidago altissima (information from Abrahamson & Weis, 1997).

FIG 171. The food web of Urophora cardui in stems of creeping thistle Cirsium arvense (modified after Redfern & Askew, 1998).

FIG 172. The food web of Tephritis bardanae and Terellia tussilaginis in flower heads of lesser burdock Arctium minus (from Straw, described in Redfern & Askew, 1998, slightly modified).

FIG 173. The food web of Urophora stylata in flower heads of spear thistle Cirsium vulgare; U. jaceana on black knapweed Centaurea nigra is very similar (modified after Redfern & Askew, 1998).

FIG 174. The food web of Lipara lucens in the shoot tip of common reed Phragmites australis (information from De Bruyn, in Williams, 1994; De Bruyn, in Raman et al., 2005). Arrows and symbols are defined in Box 10.1.

The tephritid stem galls are rounded or spindle-shaped woody swellings in the upper part of the stem: the goldenrod ball gall of Eurosta solidaginis, a North American species that galls several closely related species of Solidago, and the gall of Urophora cardui on creeping thistle Cirsium arvense, which occurs in England from Yorkshire southwards and in Wales. The galls of both species are described in Chapter 8 and their food webs are shown in Figs 170 and 171. Tephritid flower head galls are described in Chapter 7. They vary in structure from the simple accumulations of callus tissue of Tephritis bardanae and the swollen achenes of Terellia tussilaginis in burdocks (food web, Fig. 172) to the complex woody swellings of Urophora stylata and U. jaceana in thistles and knapweeds (food web, Fig. 173). The cigar gall of Lipara lucens in the shoot tip of the common reed is also woody when mature, and is enwrapped in overlapping leaves rather than hidden in flower heads. Its gall is described in Chapter 6, and its food web is shown in Fig. 174.

The tephritid galls contain specialist chalcid endoparasitoids found almost nowhere else, plus occasional eulophid endoparasitoids that are less specific. The Lipara gall contains five endoparasitoids, chalcids and braconids that specialise on Lipara species. Apart from the eulophids, all of these in tephritid as well as Lipara galls attack young host larvae, soon after or even before they have hatched from the egg, and before there is any discernible swelling of the gall (they are ‘koinobionts’, see Box 10.1). In the goldenrod gall (Fig. 170), the female Eurytoma obtusiventris lays an egg inside the egg or the newly hatched host larva in June. In the thistle galls (Figs 171 and 173), E. serratulae and E. tibialis insert their eggs into the second instar hatchling in July (Urophora larvae hatch as second instars having moulted once inside the eggshell). Pteromalus albipennis on the burdock tephritids (Fig. 172) lays its egg into early third instars in late July or August. In the Lipara gall (Fig. 174), Polemochartus liparae and P. melas lay single eggs into the host’s eggs glued to the underside of a reed leaf. These braconids do not have to pierce the gall in order to oviposit; the Lipara larva carries the parasitoid into the shoot when it burrows inside. The chalcids on the tephritid hosts use their ovipositors to drill through the gall wall while standing on the outside and, because the gall is still small and soft, they can reach larvae in all parts of the gall. Stenomalina liparae on Lipara attacks the young larva when it is still feeding above the meristem, when it is easier to reach. After a few days, all of these koinobiont parasitoids hatch inside their host larvae but grow very little until the host larva is fully fed in late summer or autumn, by which time it is large enough to provide the parasitoid with sufficient food. Then the parasitoids grow rapidly, inducing the host larva to form its puparium eight months or so early, before killing it and consuming it completely from within. The puparium is the hardened brown skin of the full-grown host larva and normally develops in the spring just before it pupates, and protects the pupa inside. The parasitoid overwinters in the gall inside the puparium. It pupates in the spring and the adult emerges by biting a circular hole in the puparium and then chewing its way out of the gall. An adult male and female of E. tibialis are shown in Fig. 175 and the juvenile stages in Fig. 176; all Eurytoma species are similar in appearance. On Terellia tussilaginis, P. albipennis emerges head downwards, making use of the hole in the base of the achene cut earlier by the host larva (see Fig. 109, Chapter 7). In the Urophora galls, E. serratulae and E. tibialis push and chew their way out of the debris that blocks the entrance tunnel to the gall chamber (Fig. 128c, Chapter 8; Fig. 114, Chapter 7), while E. obtusiventris must chew through the solid wood of the ball gall to escape (Fig. 127, Chapter 8). In the Lipara gall, the two Polemochartus species burrow out through the enwrapped leaves at the top of the gall making use of the channel cut earlier by the young Lipara larva, and the Stenomalina liparae adult chews a small round hole through these layers of leaves.

FIG 175. Eurytoma tibialis adults, a male (lower left) and a female (lower right); Torymus chloromerus adult female (top; note the long ovipositor typical of this genus). (From Redfern, 1995, with permission).

All of these endoparasitoids have one generation a year, closely synchronised with the single generation of their host. Eurytoma obtusiventris and E. serratulae are monophagous, each attacking a single host only. The flower head and shoot tip species are less restricted, specialising in the type of gall and in its position rather than in a particular host species. Thus, E. tibialis attacks Urophora hosts in woody galls in thistle and knapweed flower heads, and Pteromalus albipennis is the equivalent on Tephritis bardanae and Terellia tussilaginis in burdock heads and also attacks non-gall-causing species of these genera in thistle and knapweed flower heads. Polemochartus liparae, P. melas and Stenomalina liparae attack Lipara rufitarsis, L. pullitarsis and the non-galling stem-borer L. similis as well as L. lucens, but tend to specialise on one particular host: S. liparae and P. liparae mainly parasitise L. lucens while P. melas prefers L. rufitarsis.

FIG 176. Egg (top left), first instar larva (top right; scale = 0.2 mm), full-grown larva (bottom left) and female pupa (bottom right; scale = 1 mm) of Eurytoma tibialis (after Redfern & Askew, 1998).

The endoparasitoids described above are all solitary parasitoids, the host capable of supporting only one parasitic larva. The eulophids Baryscapus daira, Aprostocetus serratulae and A. legionarius (Figs 173 and 174) are gregarious, with one host larva containing several or many tiny parasitic larvae. The female lays eggs into a full-grown larva and kills it (they are idiobionts, see Box 10.1). The parasitoid larvae are full grown by winter and pupate the following spring, and the adults break out of the host’s skin, by now dry and papery. They do not allow the host to form a typical puparium as the solitary endoparasitoids do. The L. lucens larva may contain 40 A. legionarius larvae, its skin being stretched to twice its normal size to accommodate them. This species is primarily parasitic on L. lucens, but occasionally it is reared from L. rufitarsis and L. similis.

Besides endoparasitoids, all of these gall fly communities contain ectoparasitoids, often several species. Eurytoma robusta on Urophora and E. gigantea on Eurosta solidaginis (Figs 171, 173 and 170) are ectoparasitic at first and later become inquilinous. They attack in the summer when the galls are full-sized and the host larvae nearly full-grown. The females drill through the gall and deposit an egg on the larva or in its chamber. E. gigantea is restricted to galls that are not too large or too hard; its ovipositor cannot reach the central chamber of galls more than 12 or 13 mm in diameter and, by the end of July, the galls become too woody for the ovipositor to penetrate. In large galls of U. cardui and U. stylata, E. robusta is restricted to the peripheral larvae, with those in the centre out of reach of the ovipositor. In contrast, the thistle endoparasitoids occur in any size of gall and in any chamber because they attack well before the gall is full sized. The eggs of Eurytoma are distinctive (Fig. 176 and Fig. 207, below) and E. robusta’s egg is dark brown (Fig. 177). The Eurytoma larva hatches, quickly devours the host larva, and then feeds on gall tissue, selecting the more nutritious nutritive cells that remain lining the gall chamber. If the host contains an endoparasitoid, it will also be eaten, as long as the hard puparium has not yet formed. E. robusta may break into adjacent chambers if they are crowded close together and will eat their larvae too–so, it can behave like a predator and even grow larger than its host Urophora. The fully fed larvae overwinter in the host’s chamber and pupate in early summer, and the adults emerge when the new galls are well grown. Occasionally E. gigantea adults emerge in the previous autumn; these cannot find hosts until the following year because the galls are too hard for oviposition, and they probably fail to survive the winter.

FIG 177. The dark brown egg and newly hatched larva of Eurytoma robusta (scale = 0.2 mm) (after Redfern & Askew, 1998).

The Eurosta gall community contains a beetle inquiline (Fig. 170). The larva of Mordellistena convicta (previously known as M. unicolor) feeds primarily on the gall tissue, but usually also kills the E. solidaginis larva–occasionally adult gall fly and beetle emerge from the same gall. The adult beetles live for several months feeding on pollen from a variety of flowers and they lay eggs into older as well as younger galls, undeterred by the gall’s woodiness. Using her jaws, the female excavates a shallow pit in the surface of the gall and lays an egg into it. The hatchling burrows into the gall, feeding on the cortex and cutting a tunnel that fills with fine light-coloured frass, and its entry hole seals over. In late autumn, the larva reaches the central chamber, encounters the host larva and usually eats it. The beetle larva then overwinters in the chamber. It pupates in the spring, and the adult chews its way out at the same time as the new galls appear. M. convicta may attack galls already containing Eurytoma obtusiventris in its host’s puparium or E. gigantea, and the beetle feeds on these with no problem. It can also survive as a stem borer in ungalled goldenrod stems, but the resulting adults are smaller than those from galls–and are probably less fecund too (i.e. the females produce fewer eggs). The beetle larvae have their own parasitoids, two chalcids and a braconid.

The goldenrod gall community is simpler than that in the thistle galls, with only one ectoparasitoid, Eurytoma gigantea. In the thistles and burdocks, there are two or three. Each plant has a pteromalid, Pteromalus elevatus in the thistles and P. albipennis in burdocks, and Torymus chloromerus can attack both groups (Figs 171–3). These chalcids have similar lifecycles. The female stings a full-grown host larva, killing it before laying an egg on it or loose in the chamber. A tephritid larva already containing an endoparasitoid may also be a host as long as the puparium has not yet hardened. The larva hatches and grows rapidly, is full-grown by late September or October, and overwinters in the gall chamber alongside the host’s shrivelled skin. Pupation occurs in April or early May and the adult emerges two or three weeks later. P. elevatus has one or two further quick generations in late May and June on host larvae before the puparium hardens. The egg, first-and final-instar larvae and female pupa of both ectoparasitoids are shown in Figs 178 and 179, and adult females in Figs 180 and 175. An infrequent ectoparasitoid on Tephritis bardanae and Terellia tussilaginis in burdock heads is Bracon minutator (Fig. 172). It also parasitises non-gall-causing tephritid larvae in thistle and knapweed flower heads and can attack caterpillars that tunnel through the flower heads too. The adults emerge in July; they mate, and females lay eggs later in the summer on or near to host larvae that are full-grown. The parasitoid larva hatches and kills its host straightaway. It feeds rapidly and in September or October when full-grown, it spins a tough, silvery-grey papery cocoon of silk, often with the skin of the host attached (Fig. 181 shows the cocoon on T. tussilaginis). The Bracon larva overwinters in its cocoon and pupates the following June, and the adult bites its way out in July. B. minutator will kill host larvae already containing Pteromalus albipennis as long as the host’s puparium has not formed and, turning the tables, P. albipennis sometimes parasitises B. minutator.

FIG 178. Pteromalus elevatus, clockwise from top left, egg, first instar larva (scale = 0.2 mm), full-grown larva and pupa (scale = 1 mm) (after Redfern & Askew, 1998).

FIG 179. Egg (top left), first instar larva (top right; scale 0.2 mm), full-grown larva (bottom left) and female pupa (bottom right; scale = 1 mm) of Torymus chloromerus (after Redfern & Askew, 1998).

The Lipara lucens gall community is without chalcid ectoparasitoids, contrasting with the thistles and burdocks. Instead, in Europe, a rich variety of ichneumons and braconids has been recorded, some of them not common, and many of them not restricted to Lipara galls; Fig. 174 illustrates the commoner species. Scambus phragmitidis is often the commonest ectoparasitoid and specialises on Lipara, though not just on L. lucens. The female has a long ovipositor and lays an egg into the gall chamber, drilling through from the outside of the gall. The larvae are fully fed in autumn and overwinter in the gall. They pupate in spring and, in early summer, the adult chews its way out of the top of the gall, through the layers of enwrapped leaves. Exeristes arundinis and two Gelis species are also restricted to Lipara, though not much is known about their lifecycles. The other ectoparasitoids, including five further species of Scambus, are polyphagous, attacking other hosts as well as Lipara.

FIG 180. Pteromalus elevatus adult female (from Redfern & Askew, 1998, with permission).

FIG 181. Bracon minutator cocoon with skin of its host attached (scale = 1 mm) (after Redfern & Askew, 1998).

Many inquilines attack the cigar gall and most feed in the gall wall or in the enwrapped leaves above the chamber. Steneotarsonemus phragmitidis is a tiny mite that can kill the host if many are present in the gall. The mites distort the leaf blades and sheaths above the chamber and cause them to decay and fuse together into a hard mass, which blocks the exit to the gall chamber so that the adult fly cannot push its way out. The mite also causes a simple gall in its own right involving the youngest leaves at the growing point of the reed. Giraudiella inclusa (Cecidomyiidae) also causes its own galls in Phragmites, in the stems (see Chapter 8), and can construct them in the wall of the L. lucens’ gall. It does not seem to affect the growth of L. lucens, though. The other inquilines (Fig. 174) may be common, but do little damage to the host larva.

Moth caterpillars are regular herbivores in the tephritid flower head galls (Figs 172 and 173) although not in the stem or cigar galls. They are attracted by the plentiful food supply in the flower heads and by the succulent tissue of young, growing galls. As they tunnel through the heads, they break into the achenes and gall chambers, and readily eat any host and parasitoid larvae they encounter. Aethes rubigana and Metzneria lappa are specialists in burdock heads, while Eucosma cana is common in thistles and knapweeds, and all three species feed in ungalled as well as in galled heads. In summer, they lay eggs on the outside of the heads, attached to the bases of the bracts. A. rubigana eggs are pink at first, becoming brick-red, and are laid singly or in small groups of two or three. The hatchling chews out of its eggshell and immediately burrows into the head, through several layers of bracts and up through the receptacle. The young caterpillars mine round the edge of the receptacle leaving tunnels filled with black frass, and then turn inwards towards the young achenes, eating them and the larvae of T. bardanae and T. tussilaginis if present. Older caterpillars eat through one achene to the next and produce a lot of frass and debris held loosely together with silk. Large larvae may burrow into a neighbouring head if it is touching. A. rubigana is fully fed in September and drops out of the head, spins a loose cocoon in litter on the ground and overwinters inside. It pupates the following June and emerges in July. Eucosma cana in thistle head galls has a similar lifecycle and can cause considerable mortality of Urophora stylata; it breaks into gall chambers, leaving them packed with frass and partly destroyed. In burdocks, the caterpillar of Metzneria lappa is distinctive like others in its genus, fat in the middle and tapering to either end, and has no prolegs (Fig. 182). It feeds more neatly than A. rubigana, eating through the width of one achene to the next, leaving frass tightly packed into the empty seed coats, and lining its tunnel with silk. In September it is full-grown; it reinforces the silk tunnel and firmly cements together 9–12 achenes to form a hard cocoon. It overwinters inside and pupates the following June, and the adult emerges in July or August. Occasionally a caterpillar remains in its cocoon over a second winter, in a head by now fallen to the ground. All of these caterpillars are attacked by a number of parasitoids, most of them polyphagous braconids and ichneumons that attack several hosts in flower heads, or in stems and rolled leaves.

FIG 182. Larva of Metzneria species (scale = 1 mm) (after Redfern & Askew, 1998).

FIG 183. Larva of Palloptera species (scale = 1 mm) (after Redfern & Askew, 1998).

Fly maggots of the family Pallopteridae (Fig. 183) and Lestodiplosis (Cecidomyiidae) are occasional predators of tephritid larvae in burdock, thistle and knapweed heads (Figs 172 and 173), and will also attack caterpillars and any other larvae they encounter–but have not been recorded in Lipara galls. Sometimes they are found curled up inside the prey’s empty skin. When fully fed in autumn, Palloptera larvae drop out of the head and overwinter in the soil, and the adults emerge the following May. Lestodiplosis larvae spin cocoons, in the head or in the ground.

The final group of enemies of the tephritids and Lipara are birds and small mammals (not shown in the food webs), feeding on the larvae particularly in winter. In the USA, downy woodpeckers Picoides pubescens specialise on goldenrod ball galls. They peck neat conical holes, usually accurately locating and enlarging the tunnel excavated by the Eurosta larva ready for its adult’s emergence. The woodpeckers tap the surface of the gall to locate this weak spot and finding it informs the bird that there is a juicy grub inside. A gall without an emergence tunnel indicates that the larva died young or has been parasitised, and the woodpecker may not persevere: opening the gall will take more effort and any parasitoid larva inside will be a less good meal. Fully fed larvae of the parasitoid Eurytoma obtusiventris produce a resin that the woodpeckers dislike–they often reject these galls when partly opened. Tits and chickadees (both belong to the Paridae) attack both Eurosta and the tephritid galls. They are less tidy feeders than the woodpeckers. Black-capped chickadees Parus atricapillus make large irregular holes in the goldenrod galls by chiselling into the gall, grabbing loosened bits and tugging them free. Goldfinches and greenfinches (Fringillidae) pull thistle and burdock heads apart and extract achenes and larvae. Mice and voles attack galls that have fallen to the ground, gnawing into the chambers and extracting the larvae one by one. They often carry the galls away to store in their runs over the winter.

Although mortality of these gall flies is probably high before the gall starts to grow, when the hatchlings are free-living, it also can be high in the gall. In the Lipara species, mortality is usually higher in the more complex galls: only about 10 per cent die in the simple L. pullitarsis galls, mostly from bird predation, while parasitoids can kill 30 per cent in the larger, more complex galls of L. rufitarsis and L. lucens. L. pullitarsis often avoids much parasitism because, by late spring when the parasitoid adults are active and are laying eggs, most L. pullitarsis adults have already emerged. The parasitoids are synchronised better with the full-grown larvae of L. rufitarsis and L. lucens. Within each Lipara species, mortality is usually more severe in small galls than in large ones. Stenomalina liparae is often the commonest parasitoid of L. lucens (Fig. 174) but, because the female has a short ovipositor, it can only lay eggs successfully in host larvae in thin Phragmites shoots and before the galls are full-sized. Hosts in galls on thicker shoots are relatively safe from Aprostocetus legionarius as well as from S. liparae, but not from Polemochartes liparae, which attacks hosts in thicker shoots also.

Gall size affects mortality in tephritid galls too. Eurytoma robusta, the ectoparasitoid that becomes inquilinous and attacks several Urophora species (Figs 171 and 173), is the main enemy of U. cardui and can cause a small colony of galls to die out. In contrast, the specific parasitoid E. serratulae never causes more than about 7 per cent mortality. E. robusta females lay eggs into young galls, not yet full-sized nor hard and lignified. If host larvae in several chambers are killed, the gall stops growing; E. robusta is commoner in undersized galls. So, survival of U. cardui, and of U. stylata too, increases with gall size and with gall toughness. The ectoparasitoids are less successful in large galls when the host larvae are in chambers with thick, woody walls. These galls take longer to drill into, and larvae deep in the centre of a large gall are beyond the reach of the ovipositor. But deaths caused by caterpillars can partially restore the balance between large and small galls. Caterpillars (mainly Eucosma cana, Fig. 173) are attracted to large flower heads of spear thistle especially if they contain large galls packed with lots of chambers–the caterpillars plough through the thistle head, feeding on receptacle, florets, achenes, gall tissue and U. stylata larvae or its parasitoids. E. cana caterpillars are attracted to the largest flower heads, which are the most likely to contain galls. And a gall with many chambers increases the size of the head because of the nutrient sink effect caused by the larvae, so large-galled flower heads are a higher-quality food resource for caterpillars than small ungalled heads.

GALL MIDGE GALLS

Communities in cecidomyiid galls vary in complexity and, as in other galls, consist of parasitoids, predators, inquilines and caterpillar herbivores, with roles that often overlap and merge. They may start as true parasitoids of the gall causer, then attack other inhabitants of the gall so behaving more like predators, before becoming inquilinous, feeding on the gall tissue. A few parasitoids are endoparasitic and highly specific to a particular gall midge species, but most are less specific and are ectoparasitoids. The fungi of ambrosia galls provide another potential food source for their inhabitants. Figs 184, 190, 191 and 195 show the food webs of a range of gall midges. The community in Taxomyia taxi galls is particularly simple compared to that of the birch gall midges Massalongia and, particularly, Semudobia. Asphondylia gall communities are less complex too despite, or perhaps because of, the presence of the ambrosia fungus.

Yew gall midge galls

Taxomyia taxi causes leafy artichoke galls and small swollen buds on yew Taxus baccata (its lifecycle is described in Chapter 6, Rosettes and Artichokes). Its community is simple; the two-year galls contain two ectoparasitic chalcids, two herbivorous caterpillars that are often also predators, a predatory gall midge larva, and a gall mite that is a benign inquiline (Fig. 184a). The small one-year galls are missing one parasitoid and the caterpillars (Fig. 184b).

Torymus nigritarsus (Torymidae) is probably specific to T. taxi and attacks host larvae only in the large artichoke galls. It has one generation a year and, apart from the month-long adult stage, spends all of its life in the gall. Adults emerge in late March or April from galls nearly two years old and mate; both males and females are common. In May, the female lays eggs on to host pupae or larvae that are about to pupate, drilling with the long ovipositor from the surface of the gall and piercing the host to paralyse it, and leaving a characteristic small round scar. Like many chalcids, the female often feeds on host larvae before she is ready to lay eggs. She stings the host and remains motionless for a short time while a liquid produced from the ovipositor hardens into a tube around it. She then withdraws the ovipositor and drinks the juices of the larva that ooze out of the tube, drawn up it by capillarity. The larva’s body shrivels to not much more than an empty skin and the female moves on to other hosts on which to lay her eggs. The chalcid is, therefore, predatory as well as parasitic as an adult female. Its lifecycle is straightforward. The first instar (Fig. 185) is active, with a relatively large head and mandibles, and crawls rapidly over the host larva or pupa, killing any other eggs or young larvae of its own species (Fig. 186) or of Mesopolobus diffinis (see below) laid at more or less the same time–the host can support only one parasitoid to maturity. T. nigritarsus can also be a hyperparasitoid of T. taxi, parasitising larvae of M. diffinis (Fig. 187). The larvae of T. nigritarsus are fully fed in July and pupate in August, and the black pupae (containing fully developed adults) overwinter in the galls, adults emerging next spring. The numbers of T. nigritarsus vary dramatically; in some years it kills 90 per cent of host T. taxi, while in other years it is very rare. It does not attack hosts in the one-year galls probably because the female’s ovipositor is too long to be manipulated into these small galls.

FIG 184. Food webs in the galls of Taxomyia taxi on yew Taxus baccata (a) in the 2-year artichoke galls and (b) in the small 1-year bud galls (after Redfern & Askew, 1998).

FIG 185. Egg (top left), first instar (top right; scale = 0.25 mm) and full-grown larvae (bottom left) and female pupa (bottom right; scale = 1 mm) of Torymus nigritarsus (after Redfern & Askew, 1998). Note the smooth eggshell, hairy larvae and long ovipositor sheath of the pupa that distinguish this species from Mesopolobus diffinis (Fig. 188).

FIG 186. Taxomyia taxi pupa (stippled) inside its gall with eggs and first instar larvae of Torymus nigritarsus, all dead and shrivelled except one (scale = 1 mm) (after Redfern & Askew, 1998).

FIG. 187. Egg of Torymus nigritarsus on a full-grown larva of Mesopolobus diffinis that parasitised Taxomyia taxi; the shrivelled skin of the gall midge larva is shown at the top of the gall chamber (scale = 1 mm) (after Redfern & Askew, 1998).

Mesopolobus diffinis (Pteromalidae) is also ectoparasitic on Taxomyia taxi (Fig. 184) and attacks hosts in both one-year and two-year galls. It is polyphagous, also attacking other gall midge hosts, and has a complex lifecycle. It has three generations a year on T. taxi during winter and spring, and further generations on other hosts during the summer. Adult females arrive on the yew galls in October and November and lay eggs on to third instar larvae that are not yet full-grown. M. diffinis larvae grow rapidly, overwinter in the galls as full-grown larvae and pupate in early spring, and adults emerge in March. These produce a second quick generation in March and April, and eggs of the third generation are laid in April, on host larvae that are now fully fed or have pupated. The parasitoids pupate in May and the adults emerge in June. These adults fly off to gall midge hosts in herbaceous plants nearby where they have one or more further generations, returning to the next generation of T. taxi in the autumn. On T. taxi, most M. diffinis individuals are females (< 14 per cent are males) and the winter and spring generations are parthenogenetic, females producing more females without mating (thelytoky); the males at this time are probably redundant. The summer generations, though, are sexual, with roughly equal numbers of males and females. The juvenile stages are shown in Fig. 188. For a long time it was thought that two species of Mesopolobus attacked T. taxi because the adults varied so much in size. Only when the lifecycle was worked out in detail was it clear that small mothers arising from the overwintering generation gave rise to large daughters in the spring, their contrasting size due to the different sizes of their larval hosts. M. diffinis prefers hosts in small galls. It is not unusual for 90 per cent or more of the one-year galls to be attacked compared to about 20 per cent of the two-year galls. The female has a much shorter ovipositor than T. nigritarsus, more suitable for swollen buds and the smaller artichoke galls and, although she is capable of attacking hosts in galls of all sizes, in the largest she must crawl deep between the outer leaves before she can reach the larva.

Moth caterpillars or their damage are often found in the artichoke galls, attracted by the dense accumulation of leaves. Blastobasis lignea (Blastobasidae) is common, Ditula angustoriana (Tortricidae) less so, and both species are polyphagous, feeding on the leaves of deciduous trees as well as on yew. Both species nearly always kill the host larva, or a parasitoid, as they plough through the centre of the gall–they are thus predators of T. taxi as well as feeding on the leaves of the gall. They cause distinctive damage. B. lignea (Fig. 189) cuts through the bases of the inner leaves causing them to turn brown, then spins them and its own frass into an untidy loose shelter and feeds inside. D. angustoriana eats the inner leaves more neatly and may hollow out the gall; it leaves traces of silk but no frass–this drops to the ground. The caterpillars of both species grow in the galls during the summer and autumn and continue to feed in mild spells during the winter, and they pupate in early summer. B. lignea pupates in the ground in a loose cocoon of silk, soil and debris, while D. angustoriana pupates without a cocoon, in a gall or attached to a leaf or in the ground. The caterpillars, especially of D. angustoriana, are attacked by their own set of parasitoids, mostly ichneumon and braconid wasps (these are not included in the food web, Fig. 184).

FIG 188. Mesopolobus diffinis, clockwise from top left, egg, first instar (scale = 0.25 mm), full-grown larva and female pupa (scale = 1 mm) (after Redfern & Askew, 1998). The egg is covered with fine papillae, the larvae have few hairs and the pupa has no long ovipositor sheath (compare Torymus nigritarsus, Fig. 185).

An uncommon predator of T. taxi is another gall midge, the larva of a Lestodiplosis species. It is bright orange-red and moves actively through the gall, piercing the larva of T. taxi or its parasitoids, or the caterpillars, and sucking out their juices. More common predators are insectivorous birds, particularly tits Parus that peck open the galls, neatly removing T. taxi and any caterpillar present. A caterpillar is a bigger food item than the gall midge larva and a moth-attacked gall is probably more attractive for the tit than a gall containing only the gall midge or a parasitoid. Galls (usually the two-year galls) are more likely to be attacked in winter and spring, when the larvae are reaching their maximum size and when food for the birds may be in short supply. A final inhabitant of the artichoke gall, and a true inquiline, is the gall mite Cecidophyopsis psilaspis, which causes galls in its own right on yew (see Chapter 6, Big Buds). In T. taxi galls, the mites live on the outer surface of the innermost leaves. Usually their numbers are small and they do not distort the leaves as they do in their own big buds, and they do not affect T. taxi after the gall has formed, never penetrating its chamber.

FIG 189. Blastobasis lignea caterpillar and damage in a two-year gall of Taxomyia taxi (after Redfern & Askew, 1998). Scale = 5 mm.

Birch gall midge galls

The communities of insects in birch galls are considerably more complex than in the yew galls. Three gall midges gall birch leaves, Massalongia betulifolia, M. ruber and Anisostephus betulinum, and three species gall the fruits in female catkins, Semudobia betulae, S. tarda and S. skuhravae. The food webs of Massalongia betulifolia and of Semudobia betulae are shown as examples (Figs 190 and 191)–it is remarkable that no species attacks hosts in both leaves and catkins. The gall and lifecycle of M. betulifolia are described in Chapter 9. The S. betulae lifecycle is described briefly here. Females lay eggs in May into a catkin (Fig. 192a), inserting each egg between a fruit and its scale, and three to five days later the hatchling burrows into a soft young fruit. The gall matures quickly (Fig. 192b) and the larva is full-grown by late summer. It overwinters and pupates in early spring, and the adults emerge between March and May. Both cecidomyiids support endo- as well as ectoparasitoids, and eulophids are particularly common, a family that does not occur in the yew galls. Parasitism can be quite high: levels of 40–60 per cent have been recorded for M. betulifolia. Inquilines and predators seem to be uncommon in birch galls, particularly in the leaf galls. The larva of the ubiquitous gall midge Clinodiplosis cilicrus (Fig. 191) feeds in catkins both with and without galls, and sometimes kills the Semudobia larvae. Two gall midges, species of Dasineura (D. interbracta and D. fastidiosa), have been recorded in the Netherlands where they are lethal inquilines of Semudobia, but they have not yet been found in Britain. Predatory Lestodiplosis larvae (a species near L. trifolii) are found in the catkins too, and feed on C. cilicrus as well as Semudobia and Dasineura. Moth caterpillars may destroy part of a leaf or a catkin and with them any galls but have not been studied specifically, and bird predation has been noted only on the leaf midrib galls of Massalongia and the larger galls of Anisostephus betulinum.

FIG 190. Food web of Massalongia betulifolia galls on birch Betula leaves (modified after Redfern & Askew, 1998). No inquilines or predators have been recorded.

FIG 191. Food web of Semudobia betulae galls in female catkins of birch Betula (modified after Redfern & Askew, 1998).

FIG 192. Semudobia betulae in a female birch Betula catkin. (a) Two females laying eggs into a young green catkin still pointing upwards; scale = 5 mm. (b) A mature galled fruit with a round window spot and two ungalled fruits; the scales of the fruits lie behind (scale = 1 mm) (after Redfern & Askew, 1998).

FIG 193. The bottle-shaped cocoon of Massalongia betulifolia containing an adult Synopeas ?nervicola inside the skin of its host, ready to bite its way out through the skin and cocoon (after Redfern & Askew, 1998). Scale = 1 mm.

M. betulifolia and S. betulae both support several endoparasitoids. Synopeas ?nervicola (Platygastridae; the identification of the species is not certain) and Gastrancistrus hamillus (Pteromalidae; Fig. 190) are specific to Massalongia betulifolia, bisexual, with one generation a year and with similar lifecycles. They are koinobionts (see Box 10.1). Their larvae are still small when the host is fully fed and drops out of its gall, and spins a cocoon in the ground. Then the parasitoids grow rapidly and are soon full-grown. They overwinter inside their host’s larval skin inside the cocoon (S. ?nervicola: Fig. 193) and the adults emerge from the soil next spring. Platygaster betularia (Platygastridae) is a koinobiont with a similar lifecycle, and is specific to Semudobia betulae. It lays its egg into the host’s egg and does not hatch until the host larva has hatched and is safely inside its fruit and the gall has formed. Inside the larva, the parasitoid embryo develops an odd V-shaped appendage whose function is obscure: it may help it to absorb nutrients from its host or it may produce substances that immobilise the host’s defences, ensuring its own survival. When the Semudobia larva is fully fed, the parasitoid grows rapidly and kills its host, and by late August is full-grown (Fig. 194). It pupates in the gall, still inside its host’s skin, and the adult emerges in the autumn. The adults, both males and females, remain sheltering inside the galls during the winter, and leave them in late April or early May at the same time as the unparasitised gall midges. P. betularia adults then mate, and lay eggs into the host’s eggs soon after they are deposited into developing catkins. Another platygastrid, Metaclistus phragmitidis, occurs on S. betulae in the Netherlands and probably also occurs in Britain, although it has not been recorded.

FIG 194. The full-grown larva of Platygaster betularia (after Redfern & Askew, 1998). Scale = 1 mm.

The other parasitoids on birch gall midges (Figs 190 and 191) are idiobionts: they kill or paralyse the host when ovipositing so that it cannot develop further. Most species lay their eggs on full-grown larvae or on pupae. Four species are endoparasitic Eulophidae: Quadrastichus brevinervis and Q. ventricosus attack Massalongia betulifolia, and Aprostocetus clavicornis and A. pallipes attack Semudobia betulae. Before laying eggs, females often feed on their hosts via an ovipositor tube, as described above for the parasitoids of Taxomyia taxi. Q. brevinervis is bisexual and adults are active for a long period, from July to autumn. There is one generation a year. The adults mate and the females insert their eggs into the host, piercing the gall from the underside of the leaf. Often, one or more females lay several eggs into one host larva (up to six) but only one will survive to maturity. The long flight period often leads to hyperparasitism: Q. brevinervis can attack the host M. betulifolia, and also its other parasitoids. The full-grown Q. brevinervis larva overwinters in the gall inside its host’s skin, and pupates the following spring, emerging in the summer. Q. ventricosus differs from Q. brevinervis in several ways: it is more common on Massalongia ruber than on M. betulifolia, it has two generations a year with adults in July and September, and is thelytokous (females produce more females parthenogenetically), males being very rare. It is more polyphagous than Q. brevinervis too, attacking Anisostephus betulinum on birch leaves as well as the two Massalongia species, and also Harmandiola tremulae on aspen Populus tremula. The two Aprostocetus endoparasitoids on Semudobia specialise to some extent in that A. clavicornis is commonest on S. betulae, and A. pallipes prefers S. skuhravae and can also attack S. tarda. They are unusual in that, although the larvae are endoparasitic, they leave the host when full-grown and pupate outside its body, like an ectoparasitoid.

The remaining parasitoids in the birch galls (Figs 190 and 191) are probably all ectoparasitic idiobionts: the three species on Massalongia betulifolia are eulophids, and the seven species on Semudobia betulae belong to three families, Pteromalidae, Torymidae and Eulophidae. All of these can parasitise more than one Massalongia or Semudobia species although they tend to specialise on one. Most can hyperparasitise their gall midge host via one of the other parasitoids too, especially when their populations are large. Quadrastichus xanthosoma and Aprostocetus sp. near agrus (its identification is not certain) specialise on M. betulifolia, while Cirrospilus vittatus is polyphagous and M. betulifolia is not its usual host. Q. xanthosoma is thelytokous with two generations a year, adults emerging in June and September, while A. sp. near agrus is bisexual with probably only one generation a year. On Semudobia, Aprostocetus escherichi, A. constrictus, Torymus fuscicornis, Psilonotus adamas and P. hortensia all specialise on S. betulae, while P. achaeus is commoner on S. skuhravae. T. nitidulus attacks all three Semudobia species but is not common on any of them. The biology of A. escherichi and A. constrictus is unknown; they may be endoparasitoids.

Ambrosia gall midge galls

Ambrosia gall communities are not well known, but seem to be less complex than those in the birch galls. Their community structure is similar, with parasitoids and inquilines, though predators are not common. Fig. 195 illustrates the food web of Asphondylia sarothamni on broom Cytisus scoparius and is not complex. A. sarothamni causes two different galls each year, a pod gall and a bud gall (its lifecycle is described in Chapter 9, Ambrosia galls). The bud gall community is simple, with only two ectoparasitoids (a eulophid and a eurytomid) and no true inquilines. In the pod galls, the same two ectoparasitoids occur plus two more, a pteromalid and a torymid, and a cecidomyiid inquiline is also present. No predatory caterpillars or birds are recorded; perhaps they do occur or, perhaps, the ambrosia fungus makes the galls distasteful.

Sigmophora brevicornis (Eulophidae; originally in Aprostocetus) is common in both galls. Its way of life is unusual. It is gregarious, i.e. several individuals survive on one host larva (and gregarious parasitoids are usually endoparasitic) and, after demolishing the host larva, it becomes an inquiline, its larvae feeding on the fungi lining the gall chamber and on gall tissue. In June, a female drills into a pod gall, stings and paralyses the host larva, and lays about ten eggs into the chamber. The hatchlings compete for food and normally fewer than ten survive to pupate–and the size of the subsequent adults, and therefore their fecundity, varies considerably. Development is rapid; the larvae pupate in the gall and adults emerge in late June and July. The first adult to emerge cuts a neat circular hole in the wall of the pod and crawls out, and the rest follow. S. brevicornis can be hyperparasitic, often parasitising itself; mothers are still laying eggs when their older offspring are full-grown larvae and pupae, and will attack them, especially if the pod galls are scarce. In July, the adults mate and produce one or two more quick generations in the pod galls. After the last generation has mated, the males die and the females overwinter, sheltering in the broom bushes. In March or April, surviving females lay batches of eggs into the bud galls, having first paralysed the host larvae, which are by now full-grown. As in the pod galls, the larvae feed on the host larva and then turn to the fungus and the gall tissue, and produce new adults in May, ready to mate and to attack the next generation of pod galls. If the gall midge inquiline Trotteria obtusa (Fig. 195) is present (only in the pod galls) it provides an extra food resource for S. brevicornis. Often there are several T. obtusa individuals in a gall, and S. brevicornis does not usually kill all of them. In the pod galls (but not in bud galls), S. brevicornis larvae are sometimes themselves parasitised by a Torymus ectoparasitoid (a species near T. microstigma), which is thus a hyperparasitoid of Asphondylia sarothamni. The Torymus egg is spiny and its larva is very hairy as is characteristic of the genus. The egg is laid on a full-grown larva or pupa of S. brevicornis in late June or early July, the female piercing the gall with her long ovipositor. Sometimes, the larva feeds on Trotteria obtusa as well as directly on A. sarothamni, and so is partly a predator. The Torymus pupates in the gall in July and the adult emerges later that month by chewing a hole through the gall wall.

FIG 195. Food webs in (a) the bud galls and (b) the pod galls of Asphondylia sarothamni on broom Cytisus scoparius (information from Parnell, 1964).

Two further chalcids are occasional ectoparasitoids of Asphondylia sarothamni. Eurytoma dentata (Eurytomidae) attacks hosts in both bud and pod galls, and also can parasitise Asphondylia ulicis on gorse Ulex europaeus and Asphondylia species on other legumes. It can act as a hyperparasitoid too, via the pupae of Sigmophora brevicornis in the broom pod galls (Fig. 195). E. dentata has two quick generations in the summer on A. sarothamni, the first in the bud galls. Eggs are laid in late May and adults emerge in mid-June; these mate and lay eggs into the pod galls later in June, and their adults emerge in mid-July. The second ectoparasitoid is Pseudocatolaccus thoracicus (Pteromalidae). It is uncommon on A. sarothamni and is only found in the pod galls–it has alternative gall midge hosts. Its eggs and young larvae are found in mid-to late June, the larvae feed on full-grown larvae or pupae of the host and pupate in early July, and adults emerge later the same month. It probably has further generations each year on other gall midges.

Most Trotteria species whose biologies are known are inquilines in the galls of Asphondylia and other gall midges. T. obtusa occurs only in the pod galls of A. sarothamni. Females lay six to eight eggs in the gall cavity in mid-to late June. The larvae hatch and crawl actively around the chamber, feeding on the white hyphae that line it. If the host larva is not fully fed when T. obtusa hatches, it usually dies, crowded out by the inquilines. Occasionally, the host midge is already pupating when the inquilines hatch, and it will then survive and emerge normally–but the T. obtusa larvae will be cramped and will grow only slowly until the host adult has emerged. T. obtusa larvae are pale yellow-orange when young, and light orange with white patches (the effect of the fat bodies showing through the skin) when older. When the host larva dies, or emerges, the fungus mycelium grows rapidly to fill the gall chamber and the T. obtusa larvae tunnel actively through it. By early July, the larvae are full-grown and burrow out of the gall, piercing the wall, which is still soft, with their mouthparts. They drop to the soil and spin cocoons. Some pupate immediately and the adults emerge three weeks later, but most larvae overwinter in their cocoons and pupate in the spring, and adults emerge in early June, ready to mate and lay eggs into the next crop of pod galls.

Most inquilines in gall midge galls do not affect the size and shape of the galls they inhabit or, at least, not markedly. An exception concerns a chalcid inquiline in a complex of closely related Asphondylia galls on Atriplex (Chenopodiaceae) shrubs in southern California. Paragaleopsomyia cecidobroter (Eulophidae) lays a few eggs into the gall chamber, each attached to the wall. Immediately, tissue around each egg starts to proliferate and results in a cluster of ‘endogalls’ that cause bulges in the host’s gall. The larvae hatch and feed on the new gall tissue surrounding them while the outer wall of each endogall becomes hard and brittle. The inquilines pupate in their chambers and the adults chew their way out, leaving numerous neat circular holes over the surface of the Asphondylia gall. Although the inquilines do not feed on the host, it usually dies, crushed by the growth of the endogalls. This example is reminiscent of the effects of some inquilines in rose and oak cynipid galls (see below), but is rare in the galls of cecidomyiids and other non-cynipid gall causers.

SAWFLY GALLS

Sawflies cause a variety of galls: Phyllocolpa cause leaf folds, most Pontania cause beans (a few species cause leaf folds or rolls) and Eupontania cause peas, and Euura cause swollen stems, petioles and buds, and nearly all of them are found on willows. The galls are simple in structure (described in Chapters 4, 8 and 9): the leaf folds are slightly thickened and the Pontania, Eupontania and Euura galls are not much more than a mass of parenchyma tissue. The communities inside the galls of each genus vary considerably. Phyllocolpa and Euura gall communities are simpler than the communities in Pontania and Eupontania galls. Mortality of the sawfly caterpillars varies too, from year to year and geographically; attack by inquilines tends to decrease in the far north, although there are some specialist arctic and alpine species of parasitoids. Phyllocolpa folds have no inquilines but parasitism of the caterpillar can be high, often caused by parasitoids that also attack free-living sawflies, and generalised predators like free-roaming bugs (Heteroptera) and ants can do considerable damage. In Euura stem galls mortality caused by enemies is usually low, while in the Pontania and Eupontania galls it varies from moderate to high. Insects cause most mortality in the galls, although birds, particularly tits (Paridae), can do considerable damage and deer may eat the galls incidentally as they browse willow leaves. When fully grown, Pontania, Eupontania and Phyllocolpa larvae drop out of their galls and spin cocoons in the soil and litter, and there they can suffer considerable predation by small mammals, especially over the winter. Euura sawflies tend to escape this because most pupate inside the gall. In general and as in eriophyoid mites and in aphids, free-living sawfly caterpillars suffer greater parasitism than the gall causers. The gall protects the sawfly from some types of parasitoids (e.g. tachinid flies, which scatter their eggs over the surface of leaves so that the free-living caterpillars ingest the eggs as they eat infested leaves). A particularly simple Euura community, a Eupontania community, and a complex Pontania community are described in more detail.

The stem gall of Euura lasiolepis

Euura lasiolepis is a North American stem-galling species and its community is the best known of its genus (Fig. 196) and is remarkably simple. The larva and pupa of the sawfly are well protected inside the gall. Lathrostizus euurae (Ichneumonidae) is a specialised endoparasitoid. In early summer, the female inserts an egg into a young caterpillar in its first or second instar, drilling through the gall with its long ovipositor and locating the host with the tip. The host is not killed at this stage (the parasitoid is a koinobiont, see Box 10.1): the parasitoid hatches inside the host and grows very little until the following spring, by which time the sawfly caterpillar is nearly full-grown (in instar 5) and has spun its cocoon. There is now sufficient food for the parasitoid and it feeds rapidly, and is soon full-grown inside the shrivelled skin of its host. It bites its way out and spins a cocoon inside the host’s cocoon, and pupates inside. Later, in early summer, the adult bites out of both cocoons and emerges from the gall to start the next generation. L. euurae does not normally kill large numbers of E. lasiolepis (usually about 10 per cent, occasionally more). Although its ovipositor is long, it is very flexible and can only penetrate soft galls before they have become tough and woody. The small host larva is buried in parenchyma tissue and must be located by the tip of the ovipositor–the female moves the tip by rotating on the surface of the gall while the ovipositor is inserted up to its hilt. This technique works well in young soft galls. A week or two after the sawfly egg has hatched, the gall is 4 mm or more in diameter and too hard for the ovipositor to penetrate effectively. So, the ‘window of opportunity’ for parasitism is short and L. euurae must be well synchronised with gall growth to take advantage of it. Although L. euurae is restricted on E. lasiolepis galls, it makes up for this by parasitising other sawfly galls too.

FIG 196. The food web in the stem galls of Euura lasiolepis on arroyo willow Salix lasiolepis in the USA (information from Price, in Shorthouse & Rohfritsch, 1992).

A second parasitoid, an undetermined species of Pteromalus, is a common ectoparasitoid and attacks half-grown E. lasiolepis larvae (in instars 3 and 4) and when the galls are full-sized. This parasitoid has no problem in drilling through the tough gall wall and its target is considerably larger than that of L. euurae. It stings and paralyses the sawfly caterpillar and deposits an egg on it or nearby in the chamber. The larva hatches, bites through the host’s skin, so killing it (it is an idiobiont), and feeds on it from the outside. The chalcid grows rapidly and pupates in the gall without a cocoon, and the adult chews a hole through the gall wall to escape. Although gall toughness is not a problem for the Pteromalus female, gall size is; some galls are too large for her ovipositor to reach the larval chamber, and she is restricted to attacking galls no bigger than about 5 mm in diameter. Large galls (and some are 7.5 mm across) thus provide E. lasiolepis with a refuge from this parasitoid. One moth caterpillar, Batrachedra striolata, is recorded in E. lasiolepis galls (Fig. 196). The larva burrows through the gall feeding on the plant tissue, and will destroy the sawfly and its parasitoids if it comes across them.

Chickadees (Paridae) are particularly adept at extracting the E. lasiolepis larvae, pecking a large hole in the gall to do this. The birds are quick learners and copy others that have found the galls a good source of food. But they are generalist predators; if galls are scarce and other foods are plentiful and more easily accessible, the chickadees will turn to other prey. Elk and deer, and grasshoppers, are sporadic predators, eating the galls and their larvae incidentally while browsing the shoots.

After the sawfly or its parasitoids have emerged, the gall is left with a hole in it, which allows fungi to invade. The fungi infest the old gall and can spread to the shoot beyond it, causing decay and sometimes killing the shoot–but this can benefit the sawfly. The decay prunes and rejuvenates the willow, and encourages buds below the old gall to sprout and to produce new fresh growth. So new vigorous young shoots appear, suitable for E. lasiolepis to infest in future years.

Pontania and Eupontania leaf galls

Eupontania and Pontania gall communities are generally much richer in parasitoids and inquilines than the Euura galls. The galls of two British species are described, the simpler E. collactanea gall community (Fig. 197) and the more complex P. proxima community (Fig. 198). As well as the insects that feed inside the gall, tits peck open the galls and extract the larvae, and deer eat them incidentally while browsing the leaves, as described for Euura lasiolepis. When full-grown, the larvae drop out of the gall and may be eaten by generalised predators in the soil, like ground beetles and small mammals, and these can cause considerable mortality.

FIG 197. Food web for Eupontania collactanea galls on leaves of creeping willow Salix repens in Lancashire, England (from Redfern & Askew, 1998, slightly modified, after Hanapi, unpublished).

FIG 198. Food web for Pontania proxima galls on leaves of crack willow Salix fragilis and white willow S. alba in Britain (from Carleton, described in Redfern & Askew, 1998, slightly modified).

Ichneumon and braconid wasps are common parasitoids in Pontania and Eupontania galls, in contrast to galls of gall midges, tephritids and gall wasps in which chalcid wasps predominate. This is due, partly at least, to the larger size of the sawfly hosts; ichneumons are also common parasitoids of moth caterpillars, with many species similar in size to sawfly larvae. Lathrostizus lugens is an endoparasitoid that attacks P. proxima (Fig. 198), and is the ecological equivalent of L. euurae on Euura lasiolepis (Fig. 196); its lifecycle and biology are probably similar to L. euurae too (see above). L. lugens may itself be parasitised by Mesochorus sp., another ichneumonid (so, a hyperparasitoid of the sawfly). In contrast to free-living sawfly larvae, endoparasitoids of the gall causers are often less common than ectoparasitoids, probably because the host larvae are protected by the gall, which serves as a protection to the parasitoid too.

Scambus vesicarius (Ichneumonidae) and two braconids are common ectoparasitoids of Eupontania collactanea and Pontania proxima, laying eggs on to the surface of the larva or loose into the chamber. S. vesicarius attacks part-grown host larvae (in instars 3–5). It has a long ovipositor with which it drills through the gall wall (Fig. 199). If the wall is particularly thick, it may chew part way through it and then oviposit through the reduced thickness. It stings the caterpillar and paralyses it, and then lays up to seven eggs. The first larva to hatch kills the other eggs and then feeds on the sawfly larva–which provides sufficient food for only one individual. The parasitoid grows rapidly and pupates, and the adult chews its way out, and there are at least two generations a year. S. vesicarius may kill 25 per cent of E. collactanea larvae each year. The two Bracon species, B. picticornis and B. discoideus, parasitise older, nearly full-grown larvae, and B. discoideus is particularly common on P. proxima. Braconids typically grow rapidly, one per host, and when fully fed they spin a papery white or brownish cocoon alongside the remains of the host (Fig. 200) and pupate inside. The adult emerges from the gall after biting a hole through cocoon and gall wall. Other ichneumonids and braconids occur on Pontania proxima from time to time (Fig. 198).

FIG 199. Scambus vesicarius adult female drilling with her ovipositor through the wall of a Pontania gall; the fine ovipositor extends vertically down from the tip of the abdomen between the legs, and the ovipositor sheath is directed upwards at an angle (after Redfern & Askew, 1998).

FIG 200. Cocoon of Bracon discoideus and remains of its host Eupontania collactanea inside the gall (after Redfern & Askew, 1998).

Besides ichneumons and braconids, several chalcid species are found in these sawfly galls, all of them ectoparasitoids that attack half-grown and full-grown host caterpillars. Eulonchetron torymoides parasitises Eupontania collactanea on Salix repens and also is found on E. viminalis and E. pedunculi on other willows. Its female is odd-looking, with an elongated abdomen that tapers into a long ovipositor. It drills through the gall wall, stings and paralyses the host and lays one egg on or beside the body. The larva hatches, feeds on the host caterpillar and pupates without spinning a cocoon, as is usual with chalcids. It also may act as a hyperparasitoid of E. collactanea via the ichneumon Scambus vesicarius. The equivalent of E. torymoides in Pontania proxima galls is the common Pteromalus dolichurus, and other chalcids sometimes occur (Fig. 198). All of them also may act as hyperparasitoids of the host sawfly, attacking other chalcids or even cannibalising their own species, especially when they are common.

Inquilines are common in Pontania and Eupontania galls (Figs 197 and 198). Interestingly, no sawfly species have become inquilines in sawfly galls–in contrast to gall wasps and gall midges, both of which include species that are inquilinous in cynipid and cecidomyiid galls. Eurytoma aciculata, a chalcid, belongs to a large genus that also includes true parasitoids (e.g. on tephritid larvae, see above). E. aciculata attacks young galls of E. collactanea containing a first or second instar sawfly larva. It drills through the gall wall, pierces and kills the young larva, and lays an egg nearby. Eurytomid eggs are characteristic in shape, with a short tail at one end and a long tail at the other. Two types have been found in E. collactanea galls, one white and the other dark brown, suggesting that two species occur in these galls. The Eurytoma larva soon hatches, eats the sawfly host, and then feeds on the gall tissue, which forms the bulk of its food. It, as well as its host, is parasitised by the pteromalid Eulonchetron torymoides (Fig. 197).

The weevil inquiline Curculio salicivorus attacks galls of several sawfly species, both pea and bean galls. It is a small black beetle, less than 3 mm long including its long snout (Fig. 201). It tunnels into the thick wall of a young gall, using the jaws at the tip of its snout, and eats the egg of the host sawfly if it can reach it. It then enlarges the tunnel, or makes a new one, and lays a tiny egg, pushing it deep into the tunnel with its snout (weevils do not possess the long ovipositors characteristic of parasitic wasps). The larva hatches and chews through the gall wall, feeding entirely on plant tissue. When it is fully fed, it pupates in the central chamber that it has largely excavated itself. The weevil larvae are often parasitised. The braconid Diaparsis stramineipes is specific to the weevil (Fig. 198), while Scambus vesicarius and Bracon discoides attack it as well as the host sawfly (Figs 197 and 198). Another weevil, Apion minimum, is an inquiline in these galls in mainland Europe and probably in Britain too, but the larva has not yet been found in the galls.

The tephritid Rhacochlaena toxoneura (formerly Euphranta toxoneura) is an inquiline in galls of Pontania proxima and of several other Pontania and Eupontania species (P. triandrae, P. bridgmanii, E. viminalis and E. pedunculi; Fig. 198). Inquilinism is unusual for a tephritid; other species are normal herbivores, feeding in flower heads and stems of thistles and their relatives, and some cause galls (see Chapter 7). In spring, the female R. toxoneura pierces a young gall with her ovipositor and lays an egg near the host larva, in its first or second instar (Fig. 202). The tephritid maggot is unusually active; it can jump by flexing its body and is a voracious feeder, quickly devouring the caterpillar before starting on the gall tissue. It grows rapidly and forms its puparium in the gall chamber about two weeks after its egg hatched. It is probably the larva that overwinters, falling with the gall in the autumn when the leaf falls, and pupating inside it in its puparium in the spring. When the adult fly emerges, it must push its way out of the puparium and the gall, now partly rotted in leaf litter on the ground, using its ptilinum as an aid (the ptilinum is an inflatable balloon-like structure on its head). Moth caterpillars, e.g. Acleris schalleriana and Archips podana (Tortricidae; Fig. 198), feed on the tissues of the gall and often kill the sawfly, parasitoids and inquilines incidentally. The female moth lays an egg on the surface of the gall or on the leaf nearby–she has no piercing ovipositor–and the young caterpillar chews into the gall.

FIG 201. The weevil Curculio salicivorus ovipositing and feeding in a gall of Eupontania collactanea. (a) The adult excavating a tunnel with its snout and laying an egg (arrowed). (b) Successive larval instars eating through the gall wall, with the full-grown larva in the central chamber (after Redfern & Askew, 1998).

FIG 202. Rhacochlaena toxoneura in Pontania and Eupontania galls, (a) the adult ovipositing (scale = 1 mm) and (b) an egg and young host sawfly larva in a gall (scale = 2 mm) (after Redfern & Askew, 1998).

GALL WASP GALLS

As in other galls, cynipid gall communities contain parasitoids, inquilines and predators and their roles frequently overlap. As in other galls too, most parasitoids are ectoparasitoids (and idiobionts, see Box 10.1), the females paralysing their hosts during oviposition and the hatchlings killing them soon afterwards. The ectoparasitoids are usually not specific: on a particular host plant species, they attack several gall wasps that live in similar galls and appear at the same time of year. In addition, most cynipids are attacked by one, sometimes two, koinobiont endoparasitoids, and these are more host-specific. Inquilines are common in rose (diplolepidine) and oak (cynipine) galls, and in Diastrophus (aylacine) galls in Rubus species in eastern Asia and North America (see Chapter 9 and Box 9.1 for classification of gall wasps). Most inquilines are gall wasps that specialise on one of these tribes, Periclistus in Diplolepis galls, Synergus (and a few other genera) in oak galls, and Synophromorpha in Diastrophus galls. Other aylacine galls seem to have few inquilines–but their communities are not well known. Large complex oak galls, like the oak apple Biorhiza pallida, are often also attacked by a weevil inquiline. The inquilines may share parasitoids with the gall causer but often they attract their own additional set, and these considerably enhance the size of the gall community. Predators are not specific to particular galls. The commonest are moth caterpillars that plough through the galls when soft and are mainly after the gall tissue, and birds like tits and woodpeckers that extract the larvae of any of the species inside. These are not included in the cynipid food webs.

The complexity of cynipid gall communities varies considerably, mainly according to the structure of the gall: small simple galls, like the tiny sexual bud galls of Andricus kollari, have a simple community, while the large complex marble gall (the asexual gall of A. kollari) or the multichambered oak apple (Biorhiza pallida) or the bedeguar gall (Diplolepis rosae) have a much richer community. Small simple galls usually are without inquilines and this reduces the size of the community. Aylacine gall communities are simpler than those in rose and oak galls, also probably due mainly to their lack of inquilines. Food webs are presented for a range of gall wasp galls, aylacine (Fig. 203), diplolepidine (Fig. 204) and cynipine galls (Figs 209–13), and these show a range of increasing complexity, at least in their asexual galls.

Aylacine galls

The gall of Diastrophus rubi on brambles Rubus is the best-known aylacine species in Europe, although its gall community has not been studied in detail. It has been introduced into North America where its community is similar to that of native N. American Diastrophus galls (e.g. D. kincaidii). Most enemies in both continents are related ectoparasitoids, some specific to their Rubus host plants: Eurytoma mayri in Europe and E. brevitergis in N. America; Torymus rubi in Europe, and T. fagopirum and T. solitarius in N. America. T. rubi does turn up in rose galls too. Eupelmus vesicularis is a widespread polyphagous ectoparasitoid, native in both Europe and North America, and attacks both Diastrophus species; it is occasionally found in rose and oak galls, and in some aylacine galls, too. No inquilines are recorded from D. rubi galls in Britain, although cynipid inquilines, species of Synophromorpha, are found in Diastrophus galls in N. America. Probably the bramble gall community is richer in America than in Britain as a result.

The community in galls of Phanacis hypochoeridis, which causes swellings with several chambers in the stems of common cat’s-ear Hypochaeris radicata, has been studied in northeast England (Fig. 203). The gall is soft and succulent at first, becoming hard later in the summer. It contains up to 50 chambers, each with a larva, and these overwinter in the gall and pupate in June. Its food web includes several eurytomid and pteromalid parasitoids, and occasionally a torymid and a eupelmid–most are ectoparasitic and attack several aylacine hosts, although they tend to specialise on related host plants. No inquilines have been recorded in this gall. Eurytoma hypochoeridis and E. cynipsea are common, though often patchy, and each has one generation a year. Stinoplus etearchus is widespread, attacking other Phanacis galls as well as P. hypochoeridis. The other parasitoids are polyphagous and uncommon or rare on this host, with one European species (Thaumatorymus notanisoides) not yet recorded in Britain.

As in most galls, several predators probably feed opportunistically on aylacine galls and may cause considerable mortality. Little is known of these, although tits (Parus species) and black-capped chickadees (Parus atricapillus) feed on Diastrophus galls in Europe and North America, respectively.

FIG 203. The food web of Phanacis hypochoeridis in cat’s-ear Hypochaeris radicata in Europe (information from Ellis, 2002).

The bedeguar gall, caused by Diplolepis rosae (Diplolepidini)

The bedeguar gall, or robin’s pincushion (described in Chapter 9), is the best known of the diplolepidine galls and its community is complex (Fig. 204). The community is remarkably consistent over a large area: of the 16 species recorded from the galls, D. rosae itself and its cynipid inquiline Periclistus brandtii, and four parasitoids (Orthopelma mediator, Torymus bedeguaris, Glyphomerus stigma and Pteromalus bedeguaris) together make up more than 90 per cent of insects reared from the galls throughout Europe.

Periclistus brandtii is sometimes the commonest insect, apart from the gall causer, to emerge from bedeguar galls. It has one generation a year. In July or August, the female pierces the outer chambers of a young gall and lays a group of stalked eggs, attaching each egg to the inner wall of a chamber, and killing the young host larvae in the process. P. brandtii is, therefore, a lethal inquiline, although many D. rosae larvae in chambers deep in the gall or at a distance from the ovipositing female will survive. Soon after oviposition, the nutritive tissue surrounding the host’s chamber de-differentiates–the cells lose their characteristics (see Chapter 1) and revert to parenchyma cells with large vacuoles. This is a response to inquiline oviposition, occurring before the eggs hatch. They hatch seven to ten days later and the hatchlings feed at first on the parenchyma tissue; they are not carnivores and do not feed on the dead host larvae, which by now will have shrivelled. Larval feeding causes the parenchyma cells to multiply around each P. brandtii larva enclosing it in its own chamber, and vascular strands develop linking with the vascular tissue of the gall. Continued feeding stimulates the inner cells of the new chamber to develop into nutritive tissue, with cells that are smaller and more densely packed than typical D. rosae nutritive cells, but are otherwise similar. Then, sclerenchyma cells develop outside the nutritive zone and form a woody shell, typical of the inner gall of cynipid galls (see Fig. 138, Chapter 9). The chambers of P. brandtii are smaller, harder and thicker-walled than those of D. rosae, and occur in clusters fused together in the outer part of the bedeguar (Fig. 205), enlarging and distorting the original gall. So, P. brandtii causes gall tissue in the host plant in a similar way to D. rosae and to other cynipids, but only in an existing bedeguar gall–P. brandtii cannot cause a gall from scratch. The remaining lifecycle of the inquiline is similar to its host’s: the larvae are full-grown by late summer, they overwinter in their chambers and pupate in late spring. The adults, both males and females, emerge in June and July, a few weeks after D. rosae.

FIG 204. The food web of Diplolepis rosae on wild roses (e.g. Rosa canina) in Europe (after Redfern & Askew, 1998, with additional information from Randolph, 2005). Species marked by asterisk may have been recorded in error.

The four common parasitoids are an ichneumon and three chalcid wasps. The ichneumon Orthopelma mediator can be commoner than P. brandtii. It is an endoparasitoid and is characteristic of D. rosae galls, though it sometimes attacks other Diplolepis hosts also. The Ichneumonidae is an unusual family in cynipid galls. O. mediator is the only species that regularly attacks them in Europe–in contrast, for example, to the bean galls of Pontania sawflies on willows in which ichneumons are common (e.g. Fig. 198). O. mediator is very common, in Canada where bedeguars have been introduced as well as in Britain and mainland Europe, and sometimes is so abundant that it kills all of the D. rosae larvae in a gall. It has one generation a year, closely synchronised with its host. The female attacks immature galls and lays an egg inside a very young D. rosae larva. It is a typical koinobiont, the larva feeding very little at first and allowing the host larva to grow normally until it is full-grown in September. Then, the parasitoid grows rapidly, kills the host and is soon full-grown, fitting loosely inside its host’s skin. It breaks out of the host’s skin and overwinters in the chamber, pupating there in the spring, and adults emerge between late April and the end of July. It is sometimes itself parasitised by the bedeguar ectoparasitoids, such as Pteromalus bedeguaris.

FIG 205. Section through a bedeguar gall showing the large chambers of Diplolepis rosae and a cluster of Periclistus brandtii chambers (arrowed; scale = 10 mm); a full-grown P. brandtii larva and its mandible are shown on the right (scale = 0.5 mm) (after Redfern & Askew, 1998).

All the remaining parasitoids of D. rosae (Fig. 204) are ectoparasitic. Glyphomerus stigma, Torymus bedeguaris and Pteromalus bedeguaris are common, Eurytoma rosae and Caenacis inflexa moderately so, while the rest are rare. G. stigma (Fig. 206) parasitises the inquiline P. brandtii as well as D. rosae, and also attacks E. rosae, so it can be hyperparasitic on both of the gall wasps. It eats its host and then becomes inquilinous, feeding on the gall tissue. In Britain it is found in southern England only. It is absent from Scandinavia and most of northern Europe, but is common in Canada, occurring wherever the galls have been introduced. E. rosae (Fig. 207) also becomes inquilinous after demolishing its host larva; it is more often found parasitising P. brandtii than D. rosae and will break through from one chamber to the next, eating several larvae as well as the gall tissue–it is thus a parasitoid, an inquiline and a predator. P. bedeguaris and Torymus bedeguaris are parasitoids of D. rosae and do not attack P. brandtii. Caenacis inflexa (Fig. 208) is a specialist too, only attacking P. brandtii. Although normally primary parasitoids, all of these species can be hyperparasites of D. rosae or P. brandtii via one of the other parasitoids or, sometimes, older larvae of their own species. When hyperparasitic, their food ration is usually smaller, so that the adults developing from them are smaller than usual and produce fewer eggs. Most of these parasitoids have one generation a year, with full-grown larvae overwintering in the gall and adults emerging in June or July. Occasionally there is a partial second emergence in late summer or autumn. These individuals may succeed in laying eggs into galls still attached to rose bushes, although the galls will be very hard and woody, or they may fail to survive. C. inflexa, though, may normally have two generations a year, the overwintered larvae emerging as adults in May and producing offspring that emerge in July.

FIG 206. Glyphomerus stigma: the hairy full-grown larva (scale = 1 mm) with its face enlarged to show the pair of pits (characteristic of this species but of unknown function) (after Redfern & Askew, 1998).

FIG 207. Eurytoma rosae egg and full-grown larva (scale = 1 mm) with a mandible enlarged (after Redfern & Askew, 1998).

The remaining ectoparasitoids in bedeguar galls are rare or sporadic. They are commoner in galls of other cynipids, only occasionally spilling over into bedeguar galls perhaps because their usual host gall is rare that year. Torymus rubi normally attacks Diastrophus rubi galls in bramble stems, Aulogymnus skianeuros and Mesopolobus sericeus are usually restricted to cynipid galls on oaks, and Eupelmus urozonus is very polyphagous, often feeding on Synergus inquilines and on other parasitoids in oak galls. Mesopolobus fasciiventris (not shown in Fig. 204) has been found in bedeguars recently but usually occurs in Cynips galls and Neuroterus spangles on oaks. And other species turn up from time to time. Aprostocetus aurantiacus is probably more common in Diplolepis mayri and in the Diplolepis pea galls, and Aprostocetus eurytomae, reared from D. rosae and other Diplolepis galls in mainland Europe, has been recorded in Britain in D. spinosissimae galls. Very rarely, the weevil Curculio villosus turns up in a bedeguar gall (not included in Fig. 204); this inquiline is a regular inhabitant of Biorhiza pallida oak apples (see below).

FIG 208. Caenacis inflexa egg and full-grown larva (scale = 1 mm) with a mandible enlarged (after Redfern & Askew, 1998).

Communities in other Diplolepis galls in Britain and Europe are less well known. Their galls are less elaborate and usually smaller, often with only one chamber, and their food webs are considerably less complex. The pea gall of D. eglanteriae normally has one chamber but is enlarged and distorted by the inquiline Periclistus caninae, which causes additional chambers in the gall wall. Some of the bedeguar ectoparasitoids also attack D. nervosa and other rose pea galls. In Canada, native Diplolepis galls on native roses are better known; some are single-chambered pea galls on the leaves and others are single-or multichambered swellings in the stems. The structure of Canadian Diplolepis gall communities is similar to those in Europe, although the species are different: there is a Periclistus inquiline, an Orthopelma endoparasitoid and several ectoparasitoids, all belonging to the same genera (Eurytoma, Torymus, Pteromalus and Aprostocetus) and with similar roles in the community as in Britain and Europe.

Oak cynipine galls

Communities in cynipine galls vary, depending on the size and complexity of the galls. Tiny galls that are not much more than the inner gall have no inquilines and, usually, few parasitoids. Large galls with elaborate outer walls may house more than one species of inquiline and their parasitoid community is rich. Figs 209–13 show the enemies associated with the asexual and sexual galls of five oak gall wasps (the structure of their galls and the lifecycles of the gall causers are described in Chapter 9). In all of them, there is overlap between species; inquilines and parasitoids attack similar gall species, though they are nearly always restricted to one host plant. Conversely, overlap between the sexual and asexual galls of one gall wasp is very unusual; their communities are distinct. This is not surprising: their season of development, position on the tree, size, shape and structure, rate of growth and, sometimes, even species of tree, are usually so different that most inquilines and parasitoids are unlikely to be able to cope with both. Usually, the sexual generation of the galls appears in spring, grows rapidly and is small and simple, and has few parasitoids and no inquilines. Most asexual galls develop relatively slowly in summer, are large with a complex structure, and have rich parasitoid communities enhanced by inquilines. Some inquilines and parasitoids, though, can cope with both small spring galls and larger summer/autumn galls, but not of the same cynipid. They have two generations a year that specialise on spring or summer galls belonging to different species. The morphology of the second-generation adults is sometimes modified to cope with the larger galls–in some cases, females are larger and develop longer ovipositors in their second generation.

FIG 209. The food web of Neuroterus quercusbaccarum on oaks (Quercus robur or Q. petraea) in (a) the asexual common spangle gall, and (b) the sexual currant gall (modified from Askew, 1961c; with additional information from Williams, 2005).

FIG 210. The food web of the asexual red pea gall of Cynips divisa on oaks (Quercus robur or Q. petraea) in Britain (modified from Askew, 1961c; Gilbert et al., in Williams, 1994; with additional information from Williams, 2005). No parasitoids or inquilines are known from the tiny sexual red wart galls.

FIG 211. The food web of Biorhiza pallida on oaks (Quercus robur or Q. petraea) in (a) the sexual oak apple, and (b) the asexual root gall (modified after Redfern & Askew, 1998; with additional information from Williams, 2005).

FIG 212. The food web of Andricus quercuscalicis in Europe, in (a) the asexual knopper gall on pedunculate oak Quercus robur, and (b) the sexual catkin gall on Turkey oak Q. cerris (information from Schönrogge et al., in Williams, 1994; Csóka et al., in Raman et al., 2005; Williams, 2005).

In addition to the parasitoids and specific inquilines, there are general predator-herbivores and opportunists that attack a variety of oak wasp galls, like moth caterpillars and fly maggots (and these are not included in the food webs). The caterpillars are attracted by the succulent gall tissue, but also devour any larvae they come across. The fly maggots are true predators that explore any small crevice for prey, and probably are not restricted to searching for food in galls. In addition, detritus may collect and fungi grow in cracks and crevices and amongst hairs and spines of the more elaborate galls, and attract opportunists, like psocids, springtails and mites. These opportunists are similar to the ‘successori’ that colonise a gall after the causer and its enemies have left. So, a single gall can yield large numbers of individual insects: e.g. maybe 50 from a marble gall and a 100 or more from an oak apple.

FIG 213. The food web of Andricus kollari in (a) the asexual marble gall on oaks (Quercus robur or Q. petraea), and (b) the sexual bud gall on Turkey oak Q. cerris (modified after Redfern & Askew, 1998; with additional information from Williams, 2005).

Synergus inquilines are characteristic of oak wasp galls and are largely restricted to them. They are cynipid wasps and are the ecological equivalents of Periclistus in Diplolepis galls; like them, they modify an existing gall by inducing the plant to produce more chambers lined with nutritive cells. Some Synergus species attack several host galls on oak, including some short-lived sexual galls in spring as well as woody asexual galls in late summer and autumn. An example is S. gallaepomiformis, which has been recorded from about 20 kinds of gall wasp gall in Britain. It has two generations a year like its hosts and attacks different host species in each generation.

Some Synergus species are lethal inquilines: they kill the young host larva by crowding it out or crushing or starving it, or the adult female pierces it when she lays her eggs. S. reinhardi in marble galls of Andricus kollari (Fig. 213a) constructs chambers in the centre of the gall (Fig. 214), which crush the host larva. The inquilines are protected by the thick wall of the mature marble gall and are then vulnerable only to parasitoids with long ovipositors. But, if only one or two S. reinhardi chambers are present, the marble gall remains smaller than usual, less than 5 mm across, and is then vulnerable to a wider range of parasitoids. S. nervosus has a similar effect in the asexual galls of Cynips divisa (Fig. 210). It attacks early in development of the gall and kills the host larva, and the gall often remains small and stunted. But, if the attacked pea gall is in the ‘best’ spot on a leaf, i.e. on a side vein nearest to the midrib, then the gall can grow larger than normal and the inquilines will be relatively well protected from parasitoids. S. nervosus competes with itself for the gall: the female lays several eggs into the gall chamber and the hatchlings ‘fight’ until only one remains. Most surviving larvae overwinter as full-grown larvae and pupate in spring; but some adults emerge the previous summer, and may attack other galls. In Britain, S. reinhardi is restricted to the asexual galls of A. kollari and its close relatives, the cola nut of A. lignicolus, the ram’s horn gall of A. aries and the A. corruptrix asexual gall. S. nervosus can survive in a wide variety of mainly asexual generation galls, e.g. Andricus, Neuroterus and Trigonaspis as well as Cynips (Figs 209a & b, 210 and 212a).

FIG 214. Section through a marble gall of Andricus kollari with central chambers of Synergus reinhardi (scale = 10 mm) (after Redfern & Askew, 1998).

Other Synergus inquilines live in the gall wall well away from the host larva and do not kill the host. S. umbraculus forms chambers in the outer wall of several large galls, e.g. marble galls of A. kollari (Figs 213a and 215), knopper galls of A. quercuscalicis (Fig. 212a) and oak apples of Biorhiza pallida (Fig. 211)–and 20 inquiline chambers may be found in one marble gall. Their chambers are nearer to the edge of the galls than the A. kollari host or S. reinhardi larvae in the centre and are more likely to be parasitised by ectoparasitoids that attack the full-grown gall. Although Fig. 213a shows the variety of parasitic species on S. umbraculus and S. reinhardi to be much the same, the numbers of S. umbraculus killed are usually higher. The polyphagous Eurytoma brunniventris* and Mesopolobus sericeus, for example, are common on S. umbraculus, occasionally attack S. reinhardi and are not found on A. kollari, whose larva is furthest away from the surface of the gall. S. pallicornis forms up to six small chambers in the wall of red pea galls of Cynips divisa (Fig. 210) and also attacks asexual galls of C. quercusfolii and A. kollari and its close relatives. In the red pea galls, S. pallicornis is more likely to attack galls already containing S. nervosus and galls with both inquilines are often larger than those without. S. nervosus benefits from this association; its larvae are bigger and heavier when S. pallicornis is present and this, no doubt, leads to larger females with greater fecundity. S. pallicornis does not seem to benefit in return, though. Synergus individuals living in the outer walls of oak galls have to contend with high concentrations of tannins and other toxins, higher than near the centre of the gall, and these deter many general herbivores. The inquilines overcome this toxicity by secreting powerful enzymes that break down the tannins. The gall is thus made more palatable, allowing the inquiline larvae to survive.

A few inquilines of oak gall wasps are not cynipids. A weevil, Curculio villosus (Coleoptera: Curculionidae), is common in the oak apples of Biorhiza pallida (Fig. 211) and the larvae live almost nowhere else (very rarely they turn up in bedeguar galls); they are more common than S. gallaepomiformis and S. umbraculus, the other inquilines in British oak apples. The weevil larvae feed on the soft spongy gall tissue and on larvae of the host and parasitoids, and can inflict high mortality–and so are also predators. The adult (Fig. 216) has a very long snout, which it uses to excavate a tunnel in the gall and to push its egg deep inside. Apart from C. villosus, weevil inquilines are unusual in cynipid galls. Two gall midge larvae (Cecidomyiidae) are inquilines specific to spangle galls: Parallelodiplosis galliperda on the common spangle Neuroterus quercusbaccarum (Fig. 209a), and Xenodiplosis laeviusculi on the smooth spangle N. albipes. In both, a larva lives under the spangle gall between it and the leaf surface; it feeds on leaf cells and on gall tissue and may cause the gall to become thicker and more domed, and sometimes the host larva is killed.

FIG 215. Section through a marble gall of Andricus kollari showing the smaller chambers of Synergus umbraculus in the gall wall (scale = 10 mm) (after Redfern & Askew, 1998).

Non-specific herbivores are sometimes common in oak cynipid galls (but are not included in the food web diagrams). Caterpillars of several oak-leaf-feeding tortricid and noctuid moths feed opportunistically on gall tissue and often eat cynipid and parasitoid larvae when they encounter them. Pammene amygdalana and P. argyrana (Tortricidae) may be more restricted to feeding on galls; they attack the asexual galls and larvae of many oak gall wasps, including Andricus kollari and A. quercuscalicis, but not the sexual galls–perhaps these are too small to provide an adequate meal or the caterpillars may not live on Turkey oak. There are advantages for the moth if the gall causer and inquilines are eaten: the caterpillar gets extra protein, which supplements the low nutrient value of the outer gall tissues and, if the galler dies young, the gall stops growing and does not become so woody or toxic, and so remains more palatable for the caterpillar. As galls mature and age, particularly the larger ones, the outer tissues decay and fungi may colonise, and can attract detrital feeders. The ubiquitous gall midge Clinodiplosis cilicrus is often found in oak apples of Biorhiza pallida and between the outer scales of the oak artichoke Andricus foecundatrix, but does not affect the gall causers.

FIG 216. Curculio villosus adult (scale = 1 mm) (after Redfern & Askew, 1998).

In addition to the inquilines, the parasitoid community in oak cynipid galls is large and diverse. Most parasitoids are not specific to a single oak gall species but attack the host or its cynipid inquilines or other parasitoids in galls that are similar in structure, in position on the host plant and in their season of growth. Often the gall causer is attacked by one or two endoparasitoids; these are usually not common and there are not nearly as many species as ectoparasitoids, nor such a variety of life styles. Some ectoparasitoids, like most of the endoparasitoids, only attack the gall causer–these are the larger species that require a larger food ration than an inquiline or another parasitoid can provide. Other species only attack Synergus inquilines, or their parasitoids, in the outer gall; their ovipositors are not long enough to reach hosts in the centre of large galls. Some species are less restricted, and attack the gall causer, the inquilines and other parasitoids of either, including older individuals of their own species–so, they can be primary parasitoids, hyperparasitoids and cannibals, sometimes all in the same gall (most hyperparasitic links are not shown in the food webs). A few species are very polyphagous, attacking galls on roses as well as on oaks and sometimes parasitising insects in other orders (e.g. fly maggots and moth caterpillars in oak leaf mines). As the degree of polyphagy and the number of types of gall attacked increases, so the number of generations a species has often increases, perhaps to three, four or five a year. Polyphagous species reproduce sexually, and males are common. A few parasitoid species are both ectoparasitoids and inquilines, but at different stages of their larval life. Some feed on and kill the larvae before starting on the gall tissue and others eat gall tissue first, allowing the gall and the host larva to grow before they kill and eat the larva.

An oak wood with a large variety of cynipid galls has a rich and varied parasitoid fauna coexisting in the community. Coexistence is possible because of slight differences in the way of life of each parasitoid species, even though many search for the same galls and must compete to some extent–their choice of particular galls, their flight times, length of ovipositor and female behaviour, number of generations a year, feeding behaviour in the gall, etc., result in sufficient difference to allow a large variety of parasitic species to live together. Descriptions of selected parasitoids follow, which illustrate the variety of their life styles.

Most endoparasitoids of oak gall wasps belong to two chalcid families: Eurytomidae and Eulophidae. They are more host-gall-specific than most ectoparasitoids, they usually attack the host larva early in its development and they are well synchronised with the host’s lifecycle, and often have strongly female-biased sex ratios (some are thelytokous). The eurytomid Sycophila variegata parasitises oak apples of Biorhiza pallida and marble galls of Andricus kollari and, in Britain, is known from only two other galls, the agamic galls of A. corruptrix and the sexual galls of A. grossulariae. Its relative, S. biguttata, attacks a wider range of hosts: the pea gall of Cynips divisa, the knopper gall of A. quercuscalicis, the marble gall of A. kollari, and the asexual galls of five other Andricus species, two Cynips species and Aphelonyx cerricola, a recent introduction to Britain. Both Sycophila species have a similar life history with one generation a year. They are restricted to the gall causer (or almost so, in the case of S. biguttata, which has occasionally been reared from inquilines in the outer gall of A. quercuscalicis; Fig. 212a). Both species are koinobionts. The female attacks early when the gall and the host larva are young and small. She pierces the gall with her rather short ovipositor and inserts an egg into the host larva; the egg remains dormant, not hatching until the host larva is fully fed, perhaps three months later. By this time, the host is large and provides the parasitoid with sufficient food, and the gall is full-sized and hard, vulnerable only to parasitoids with long tough ovipositors, like species of Torymus and Megastigmus; and so the endoparasitoid is partially protected from hyperparasitic ectoparasitoids. The Sycophila larvae grow rapidly in late summer and autumn, and the full-grown larvae overwinter. They pupate in the spring and adults emerge in summer, S. variegata in June and S. biguttata over a longer period, from early to late summer.

Endoparasitic eulophids are species of Pediobius and Baryscapus. Two Pediobius species specialise in Neuroterus spangle galls (although P. lysis has not been found in the common spangle, Fig. 209a), and P. clita also attacks the tiny catkin galls of Andricus quercuscalicis (Fig. 212b). Baryscapus diaphantus is occasionally found on Biorhiza pallida in oak apples and B. berhidanus attacks A. quercuscalicis in the inner gall of knopper galls, though it is not yet recorded in Britain. Baryscapus species are gregarious, i.e. several parasitoid larvae develop inside each host larva, which is unusual on oak cynipid hosts.

A few ectoparasitoids of oak cynipids have some characteristics of the endoparasitoids. Torymus cyaneus is restricted to asexual Cynips galls and attacks the host larva only (Fig. 210). The female attacks galls early when they are small but does not paralyse or kill the young Cynips larva. The parasitoid hatches and feeds first on gall tissue, so allowing the gall and the host larva to grow, and only kills and eats the host later when it is sufficiently large to provide a decent meal. By this time, too, the gall is tough, which helps to deter later-attacking ectoparasitoids. So T. cyaneus is an inquiline first before becoming parasitic. The eulophid genus Aulogymnus is similar, attacking early and delaying killing the host larva until the gall is full-sized, although it does not feed on the gall tissue. It causes the gall to become harder and thicker-walled than normal, so giving greater protection to its own larvae. Some species are thelytokous with males very rare and most species specialise on relatively few hosts. A. gallarum attacks Neuroterus spangle galls, the currant galls of N. quercusbaccarum (Fig. 209a & b), and the sexual galls of Andricus curvator, A. quercusramuli and A. quercuscalicis (Fig. 212b), which are on the trees at the same time of year. Ormocerus vernalis (Pteromalidae) is also thelytokous, without males, and with a restricted host range: it is known only from the sexual galls of Neuroterus albipes and, in mainland Europe, from the catkin galls of Andricus quercuscalicis (Fig. 212b).

Eurytoma brunniventris is an unusual ectoparasitoid, with a different strategy from the endoparasitoids and the more specific ectoparasitoids. It also attacks the gall early, but the ovipositing female kills the young host larva, and so the gall stops growing and remains small and thin-walled. The hatchling feeds on the dead host and on any inquiline and parasitoid larvae it can find, and then becomes inquilinous; if there are no other larvae present, it can complete its development on gall tissue alone. It attacks a wide range of galls: asexual galls of Andricus, Cynips, Neuroterus and Aphelonyx and sexual galls of Andricus, Biorhiza, Neuroterus and Callirhytis–but not the tiny Andricus bud galls on Turkey oak. Its short ovipositor restricts it to the gall causer and inquilines in small and young galls; in large galls, it can only reach larvae near the surface of multichambered galls or inquilines in the gall wall. In Cynips divisa pea galls (Fig. 210), E. brunniventris is usually found in the main chamber, eating the host larva or the inquiline Synergus nervosus as well as the gall tissue. Apparently it does not eat S. pallicornis, even though its chambers in the gall wall are within reach. The relationship between E. brunniventris and S. pallicornis seems to be competitive, both feeding on the gall tissue. S. pallicornis is the superior competitor, taking the lion’s share of the food and, although the E. brunniventris larvae survive, they are smaller when S. pallicornis is present. E. brunniventris has a long flight period, from spring to autumn, and probably has at least two generations a year, attacking full-grown galls as well as young ones. In young galls, it is heavily parasitised by polyphagous ectoparasitoids because these galls remain small and thin-walled. It is safer in larger galls but, in these, oviposition is a slower process–galls of Cynips divisa take females less than a minute to drill into when attacked early in their development but more than half an hour when full-sized. During this time the female is vulnerable to ants and other free-roaming predators.

Eupelmus urozonus (Eupelmidae) is a true ectoparasitoid and is very polyphagous: in Britain, it is known from 16 oak cynipid galls with varied structures and occurring on buds, leaves and stems (including those in Figs 209–13), and it also attacks galls of Diplolepis rosae on roses (Fig. 204) and Diastrophus rubi on brambles. It attacks the gall causer or its cynipid inquilines or their parasitoids and is common, with several often emerging from one gall. It has two generations a year and males and females are equally common. The male is smaller than the female and he mates while clinging upside-down beneath her abdomen with his head pointing backwards. Copulation is brief, but the courtship that leads up to it is long and elaborate. In May or June, females lay eggs into a variety of spring galls, e.g. the sexual galls of Andricus curvator and Biorhiza pallida, and asexual galls of A. paradoxus, and these produce adults in July or August. These second-generation females lay eggs into the asexual galls of A. kollari, A. foecundatrix and many others, overwinter as full-grown larvae and pupate in the spring, with adults emerging in May or June. The behaviour of females before they lay eggs may be typical of parasitoids in general. Females investigate each gall thoroughly by drumming over its surface with their antennae before drilling into it, and may make five or six trial drillings into each gall, each lasting ten seconds or so. In E. urozonus, if the female accepts the gall, she will lay an egg during a drilling that lasts at least 20 seconds. Inside the gall, and unusual behaviour for a parasitoid, the egg is bound to the gall wall with silk threads (Fig. 217)–the silk is produced from the tip of the ovipositor. Usually only one egg is laid per chamber; if more only one hatchling will survive. Unusually for a polyphagous parasitoid, the host larva is not paralysed or killed by the ovipositing female; perhaps this is unnecessary because the silken tent protects the egg from damage by the host larva. As soon as it hatches, the parasitoid larva kills the host. E. urozonus larvae are hairy, though less so than Torymus larvae, and have strong mandibles.

FIG 217. The egg of Eupelmus urozonus (Eupelmidae) protected in a silk tent (after Askew, 1961a).

In contrast to E. urozonus, Torymus affinis (Torymidae) is a specific ectoparasitoid, known only from oak apples of Biorhiza pallida (Fig. 211a) where it is common. It has one generation a year. Females lay eggs into growing oak apples in early May, one per chamber without paralysing the host larvae, and these survive for several days until the gall is full-sized. Then the parasitoid larvae kill their hosts, feed and grow and overwinter fully fed. They pupate in early spring and males and females emerge in April. T. affinis (Fig. 218) is an attractive insect, brilliant metallic green with golden highlights, the female with a long slender ovipositor twice the length of her body. The egg is white and lemon-shaped, the larva hairy, and the pupa of the female carries the ovipositor curved along its back (Fig. 219).

Galls that are similar in size and structure and that occur in a similar position on a host plant and at the same time of year tend to have similar sets of parasitoids and inquilines. But there are significant differences, and these allow the members of the community to live together. The asexual galls of Cynips divisa, C. longiventris and Neuroterus anthracinus provide an example. The galls are small, hard and globular, and are attached to veins on the underside of leaves throughout the summer. C. divisa is the best known of the three (Fig. 210). The C. longiventris community is similar, except for lacking an inquiline (Synergus albipes). The oyster gall (N. anthracinus) community is slightly less species-rich with fewer parasitoids, but includes the three inquilines. In C. divisa, the enemies attack in a particular sequence. First, Eurytoma brunniventris and, second, Synergus nervosus larvae appear in the galls, within a week of their appearance. A few days later, two Torymus species appear. T. auratus paralyses the host or S. nervosus larva before laying an egg and the gall stops growing at only 2–3 mm diameter. T. cyaneus, described above, allows the host larva to survive for a while and its larva feeds on the gall tissue until the gall reaches its full size of 4–5 mm diameter. So, T. auratus is vulnerable to later parasitoids while T. cyaneus is more protected by the thicker and harder gall wall. Then two Mesopolobus species, M. fasciiventris and M. sericeus, attack in July, mainly targeting the Synergus inquilines and E. brunniventris, but also the host C. divisa if any have survived to this stage. At the end of August and through September and October, second-generation females of T. auratus and T. geranii (its first generation attacks several oak cynipid galls, but not C. divisa) lay eggs on surviving host, inquiline and any full-grown parasitoid larvae, and then the third generation of M. fasciiventris and M. sericeus attack these Torymus species, and any other larvae still alive. As a result, in some years, very few C. divisa survive to cause galls the following year. But survival varies tremendously depending on the size of the various inquiline and parasitoid populations in that year and on the abundance of other galls including C. longiventris and N. anthracinus that they can attack. Many of these galls drop to the ground during September and October, where they have more chance of avoiding the later parasitoids, and relatively few are eaten by tits and small mammals–and so, on the ground, surviving host larvae are relatively safe.

FIG 218. Torymus affinis (Torymidae), adult female. (Robin Williams)

FIG 219. Torymus affinis egg (scale = 0.5 mm), full-grown larva and female pupa (scale = 1 mm) (after Redfern & Askew, 1998).

A third Cynips species, C. quercusfolii, causes cherry galls (the asexual galls) on the underside of oak leaves from June to October and partially shares a community with the pea galls. Cherry galls are much larger with very thick walls (about 4 mm thick) and they grow very rapidly early in the summer. As a result, they escape some of the pea gall enemies. Cherry galls are attacked by the same three Synergus inquilines (see Fig. 210) but S. nervosus, which kills the host larva in the centre of the gall, is quite rare and S. pallicornis, which does not, is more common. S. nervosus is restricted to attacking the gall for a brief period early in early summer when the gall is small, while S. pallicornis, forming its chambers in the thick gall wall, is not so restricted. A major enemy of C. quercusfolii larvae is the endoparasitoid Sycophila biguttata and by late summer and autumn most of these, as well as many of the C. quercusfolii hosts, have been parasitised by Torymus auratus, its long ovipositor able to reach the centre of the mature gall. So, in future years, the S. biguttata population will be smaller than it might otherwise have been, which benefits C. quercusfolii in the long run.

The sexual galls of Neuroterus albipes, N. numismalis and N. quercusbaccarum form another group that occur together on oak leaves in spring, and that have some enemies in common but significant differences too (Fig. 209b shows the food web of the currant galls). Four parasitoids occur in all three Neuroterus galls: Aulogymnus arsames, Eurytoma brunniventris, Torymus flavipes and Mesopolobus tibialis. The small thin-walled blister and egg-shaped galls of N. numismalis and N. albipes share three more species, Aprostocetus aethiops, M. fuscipes and M. fasciiventris, which do not occur in the larger, thick-walled currant galls of N. quercusbaccarum. The currant galls share an inquiline, Synergus albipes, and the polyphagous M. sericeus with N. numismalis blister galls, but currant galls otherwise are different, with four additional inquilines and three more parasitoids. Currant galls are common on oak catkins as well as on the young leaves, and usually more host larvae survive in the catkin galls because the inquilines and parasitoids are less common and there are fewer species; the catkin galls are missing Eurytoma brunniventris and Mesopolobus sericeus, both of which are common in the leaf galls. But leaves are a commoner resource than catkins, which may partially redress the balance.

Like galls of other groups, cynipid galls are attacked by a variety of opportunistic enemies that target the succulent tissue of the gall or feed on the larvae it contains. Moth and sawfly caterpillars can cause major mortality of gall causers in young galls. Older galls are harder and with higher concentrations of tannins, and are less vulnerable. Predatory insects, birds and mice chew or peck holes in the galls and extract any larvae they find. Hornets and common wasps (Vespa and Vespula species) open soft galls, like oak apples and cherry galls when young, while great tits Parus major and nuthatches Sitta europaea regularly peck open large woody galls as well as the softer galls of spring generations. Long-tailed tits Aegithalos caudatus are regular predators of C. quercusfolii in cherry galls and, in some years, they destroy nearly all the galls on particular branches of a tree. Woodpeckers and nuthatches can recognise acorns containing Callirhytis erythrocephala, indistinguishable from ungalled acorns to the human eye. Woodmice Apodemus spp. gnaw holes and extract the larvae from woody galls fallen to the ground and, in mainland Europe, wild boar Sus scrofa eat them incidentally while rooting for acorns–the extremely tough walls of Callirhytis galls inside the acorn sometimes enable the larvae inside to survive the passage through the boar’s gut! Deer (Cervidae) eat galls on the tree as they browse the leaves and eat acorns on the ground too, but avoid those with knopper galls of Andricus quercuscalicis. Some woody galls, e.g. marble galls of A. kollari, remain on the tree long after the causer and its enemies have left, and provide nest sites for ants, solitary bees and sphecid wasps, which come and go through an exit hole. Finally, fungi often colonise old oak galls and provide a food source for successori. Some species are more specific and colonise earlier, when the gall is young. An anamorphic fungus, Apiognomonia errabunda (formerly Discula quercina; Valsaceae: Ascomycetes), can kill a high proportion of Andricus species in laboratory conditions, but probably has little effect in the wild. (An anamorphic fungus has no known sexual spores so that its classification is uncertain.) The high tannin content of the outer wall of many oak galls may deter fungal attack, and many cynipid galls have a higher tannin concentration than the nearby ungalled oak leaves.

A synthesis: some general questions arising from studies of interactions in galls

The communities in galls present a more or less intricate array of enemies, all interacting and affected by one another to a greater or lesser extent, an extent that varies over time and from place to place. Discovering the impact of one species on another over a period of time is complex enough, and expanding this outwards to show how the intricacies of one gall affect others coexisting on the same host plant is even more so–particularly for cynipid galls on oak trees. Construction of food webs, as presented here, makes a start at unravelling such intricacies. The next step would be to construct ‘quantitative’ food webs: to show the abundance of each species in a web and how frequently its links with other species occur. Extending such information on one gall to other galls with inquilines and parasitoids in common, and eventually to all the galls dependent on the host plant, would reveal invaluable information about the structure and workings of the oak gall community. In temperate areas of the world, e.g. Britain, Europe and North America, oak trees often dominate the forests and the oak gall community is richer than any other. In other areas, other plants are dominant and their gall communities are rich, such as those on eucalypts and acacias in Australia. Galls of other trees and, particularly, of small herbaceous plants are fewer in number and their interactions are probably less complicated, but no less interesting. The field is wide open for studies of this kind.

Other points arise from the detailed studies of communities in galls and have a wider relevance to ecology or to evolution; these raise questions that are discussed further below or in other chapters. The gall provides food for a highly specialised herbivore–through the provision of nutrient tissue and an increased vascular supply, the gall and its larva become a nutrient sink, ensuring that sufficient resources pass to the larva (and thence to parasitoids and inquilines) until the host is fully fed (see Chapter 1). How this occurs is a question under active investigation in research centres in Britain, Europe and elsewhere. The size and complexity of the community are related to the structure and size of the galls–though not exactly: some simple galls have surprisingly large and complex communities. Why does the galling herbivore cause such an elaborate structure to develop when other herbivores, sometimes related, browse in the open or mine in stems or leaves, and survive without inducing extra tissue? Answers may involve avoidance of plant defences (such as tannins and other toxins), reduction of competition from other herbivores on the same plant, or protection from enemies or adverse weather. Or, answers may relate to competition between the parasitoids and inquilines of the gall causer, at the next trophic level up.

Protection from enemies invokes the idea of a coevolutionary ‘arms race’: an innovation from one side (a parasitoid or inquiline) leads to a response from the other (the host), and vice versa. So, as galls become thicker-walled, say, or hairier or sticky (see below for a list of characteristics), so the enemies must evolve adaptations to cope, or depart and find alternative hosts elsewhere. Unlike a true arms race, which leads to deadlier and more numerous weapons and, perhaps, annihilation of the other side, the parasitoids and inquilines should not become so efficient that they jeopardise the future of the gall causer, perhaps their only source of food. A better analogy is the ‘Red Queen Hypothesis’ (Van Valen, 1973)–the Red Queen in Lewis Carroll’s Alice Through the Looking-Glass says to Alice ‘Now here, you see, it takes all the running you can do, to keep in the same place.’ In other words, innovations from each side are countered by the other, with each continually striving to keep up.

Gall-causing has evolved independently many times, in unrelated groups of organisms, suggesting that it must provide some advantages. But within most families of insects and mites, it occurs only in a minority of species; most herbivores survive without inducing a swelling in the plant. The gall midges (Cecidomyiidae) and gall wasps (Cynipidae) are exceptions, with a large proportion of both families causing galls. Even in these, though, one of a pair of closely related species may cause a gall while the other does not. These evolutionary questions are discussed further in Chapter 11.

Another question involves variation of the gall community across its geographical range. In galls of some species, the community is remarkably constant wherever the gall occurs. In others, the community is rich in some areas and less so elsewhere. This may be due to a natural expansion in the range of a species or of its host plant, with the enemies taking time to catch up. Or humans may intentionally or accidentally introduce a gall to a new area, without its normal complement of enemies. Distribution of species is discussed further in Chapters 12 and 14.

A final question concerns the population dynamics of species. All species fluctuate in abundance over time and numbers vary from place to place. Do enemies affect the numbers of a gall-causing population (top-down factors; see Chapter 10 and its footnote)? Or are enemies unimportant compared to the effect of the plant and its environment (bottom-up)? Some examples are discussed in more detail below.

Why are there so many shapes and sizes of galls on a single plant species? In Britain and Europe, this question is often asked of cynipid galls on oaks but also applies to other insect groups and other host plants, such as roses, willows and poplars. In North America, gall diversity on oaks is higher than in Europe, related to the high number of native oak species, and their gall communities are probably highly complex (but are not well known). In other parts of the world, common trees are eucalypts, acacias, pistacias, etc. and these have large varieties of galls. Unlike cynipids on other plants, most oak gall wasps have alternating sexual and asexual generations with each causing a gall of a different size and shape, and this produces an additional complication (alternating generations occur in the sycamore gall wasps too, but in no other cynipid group). The answer to the question may involve pressure from enemies of the gall causer, and may be to do with competition between the parasitoids and inquilines, each species carving out a niche that is sufficiently different from others to allow it to survive. The answer may also involve interactions that occurred in the past but are now redundant. A particular structure that evolved to protect a particular parasitoid would never be lost even if the parasitoid had since disappeared and the structure was no longer useful. Galls are the product of the host plant and so are not costly to the gall causer–so there is no pressure for them to be lost during later evolution. The following bullet points refer particularly to oak cynipid galls in Britain and Europe, but many are also relevant to the galls of other groups on other plants.

Gall wall thickness–the host larva is attacked by fewer parasitoid species in thick-walled galls than in thin-walled galls, because the parasitoids are restricted by the length of the ovipositor. Apart from oak galls, this also occurs in tephritid galls in thistles (and, in North America, in goldenrod): parasitoid species with short ovipositors attack smaller, younger galls while those with longer implements can attack hosts deeper within the gall. Torymus flavipes in oak galls does both: females of its two generations have ovipositors of different lengths and can attack both thin-and thick-walled oak galls.

Gall toughness–drilling by parasitoids is more difficult and takes longer in tough than in soft galls, exposing the ovipositing female to predators for a longer time. Some parasitoids can cope with tough galls better than others: the tip of the ovipositor of Torymus auratus, for example, is hardened with a high concentration of manganese. The tough outer wall of many cynipid galls probably deters predatory moth caterpillars, too.

Number of gall chambers–many-chambered galls are larger than single-chambered and host larvae in the central chambers are more protected than those near the periphery. Inquilines that form chambers in the gall wall add further chambers, which have a similar effect of protecting the gall causer deep in the centre (unless, of course, the inquiline kills its host). Urophora (Tephritidae) larvae in multichambered thistle galls are protected in the same way as larvae of Biorhiza pallida in oak apples and Diplolepis rosae in bedeguar galls.

Sticky coatings of resin–stickiness gums up the delicate valves of the parasitoid’s ovipositor, which then require thorough cleaning and so the female lays fewer eggs in the time she has available. While cleaning their ovipositors, females are exposed to predators, and they sometimes get stuck fast by the resin while ovipositing.

Coatings of sugary nectar–nectar is produced on the surface of some Andricus, Disholcaspis and Dryocosmus galls on oaks (though not in Britain). The nectar attracts ants, which drive away any parasitoids and inquilines attempting to lay eggs. This is an unexpected adaptation because oak trees do not otherwise produce nectar (their flowers are wind-pollinated).

Hairiness and spines–dense hairs probably make oviposition by insect enemies more difficult while stout spines may deter vertebrate browsers like deer.

Air spaces between the inner and outer gall–these occur in several cynipid galls on oaks; they make egg-laying more difficult because the ovipositor is unsupported as it crosses the air space, reducing the force it can exert at its tip without buckling. The hardness of the shell of the inner gall is often greater than the outer gall, too, which adds to this problem. Some inner galls are loose balls or are slung from the outer gall by a thin thread; they move when touched by the tip of the ovipositor, making it difficult for the tip to pierce them.

Parasitoids and inquilines are also affected by non-structural traits of galls: e.g. the family to which the host plant belongs, the location of the gall on the plant and the time needed for gall development, particularly the ‘window’ during which the gall is vulnerable to attack. Successful parasitoids adapt in different ways, becoming more or less specialised to cope with all of these various traits, and the big variety of adaptations helps to explain why the numbers of galls and the richness of the enemy community is so great–especially in a host plant like an oak tree that provides a large variety of microhabitats. The gall does seem to protect the causer from its enemies in some groups, but not in others. In eriophyoid mites, aphids and sawflies, which all cause relatively simple galls, free-living species have more enemies than the gallers. But in thrips, tephritids, chloropids and cecidomyiids, the gallers have more enemy species than the free-livers. Cynipids are all associated with galls, and have the greatest variety of enemy species; there are no free-living species in the family, though, with which to make comparisons. In many galling groups, therefore, competition between enemies for the gall causer may be more relevant than protection for the galler.

Moving on to questions that involve population dynamics. Why is a species common or rare in a particular area? Why does its abundance fluctuate so much, or so little, from year to year? Are top-down forces more important than bottom-up? Two case studies indicate that both occur.

The North American sawfly, Euura lasiolepis, is affected by drought. It causes stem galls in shoots of its host plant, the arroyo willow Salix lasiolepis (described in Chapter 8; see Price & Hunter, 2005). Variable winter rain and snowfall are responsible for most of the change in sawfly populations from year to year: 52 per cent of the change in the wetter sites and 73 per cent in the dry sites. When winter rain and snow are relatively plentiful, the willows grow well and produce plenty of long shoots in the following spring, and sawfly females favour these shoots when searching for egg-laying sites. The females lay more eggs on young, vigorously growing shoots that will grow long (receptors in the ovipositor recognise these shoots and are attracted by their high concentration of the chemical tremulacin, a phenolic glycoside), more and larger galls develop, and more of the subsequent larvae survive to become adults and to produce the next generation. In drought years especially in dry areas, willow growth is poor and sawfly numbers decline, producing poor crops of galls in the next few years. The effect of sawfly enemies is negligible and, if anything, reinforces the effects of climate. The commoner parasitoid, Pteromalus sp. (see the food web, Fig. 196), has less effect on sawflies in ‘good’, i.e. wet, years than in drought years because it cannot reach the host inside large galls; thus, it cannot take advantage of the high number of galls in wetter sites or in wetter years. Fluctuation in numbers of this sawfly, therefore, is almost entirely due to bottom-up effects that affect the growth of the host plant on which the sawfly depends.

The yew gall midge Taxomyia taxi (described in Chapter 6, Rosettes and Artichokes) in southern England provides a contrasting scenario in which the top-down effect of parasitoids on population numbers is much more significant than the bottom-up growth of the host plant (Taxus baccata; Redfern & Hunter, 2005). The gall midge, which causes galls in the leaf and flower buds, fluctuates in numbers over the long term, with peaks and troughs at intervals of 14–20 years (Fig. 220 shows an example of the typical pattern). A simulation model, constructed from the rate of growth of the tree and from the numbers of the host and its parasitoids in the galls over 35 years, confirmed that the effect of one parasitoid overrode the effect of tree growth. Torymus nigritarsus fluctuates in a similar fashion to its host but with its peaks and troughs usually delayed by a few years. Each species regulates the numbers of the other but only after a delay: egg-laying, growth and survival of the subsequent larvae, and the performance of the next and subsequent generations of adults all take time, and this means that the response of the parasitoid to its host, and vice versa, cannot be immediate. It is these time delays that cause the fluctuations in numbers of both host and parasitoid. In contrast, the second parasitoid, Mesopolobus diffinis, has little effect on its host’s numbers; it followed the fluctuations of T. taxi very closely and was dependent upon it. The growth of the yew tree had negligible effects.

FIG 220. Fluctuation in density of Taxomyia taxi and its two parasitoids in artichoke galls (the two-year lifecycle) in even-year generations on a single tree at Kingley Vale, West Sussex (modified from Redfern & Hunter, 2005). Between 1976 and 1980, T. nigritarsus was too rare to appear in samples. M. diffinis = Mesopolobus diffinis; T. nigritarsus = Torymus nigritarsus.

The population numbers of gall insects, as for all organisms, are regulated in different ways, by bottom-up or top-down influences or by a mixture of the two, but always imprecisely. Naturalists often comment upon, say, the glut of common spangle galls (caused by the gall wasp Neuroterus quercusbaccarum) or of pocket plums (caused by the fungus Taphrina pruni) in some years and their dearth in others. Fluctuations in abundance are to be expected, with occasional higher peaks or lower troughs occurring, all due to the intricate web of interactions between the host plant, its environment, the gall causer and others on its host plant, and the various enemies of the gall causer that make up the subcommunity in the gall.

CHAPTER 11

Evolution of Galls

GALL-CAUSING HAS EVOLVED independently many times. That this is so is clear from the huge range of organisms in which it is found: bacteria, fungi, nematodes and mites and insects. Amongst insects, the occurrence of gall-causing species in many different groups shows the same thing. Furthermore, it has evolved in groups with different original habits. Many gall insects, particularly thrips, aphids, scale insects, tephritid flies, beetles, moths and sawflies, have evolved from herbivorous ancestors that fed on spores, pollen, sap, foliage, stems or roots. Gall midges and gall mites evolved from detritivores or from species that ate fungi or bacteria on detritus, and gall wasps, fig wasps and the few chalcid and braconid gall-causing wasps are derived from parasitoids. The evolutionary history of insects, and of gall-causing groups amongst them, is much better known than that of other gall causers and this chapter concentrates on them.

TRACING EVOLUTION–FOSSILS AND THE OLDEST GALLS

The obvious way to trace evolution is through the fossil record. As with most other animals and plants, however, the record is far from complete. Also, a fossil gall may not indicate what caused it, and it may be impossible to tell whether a fossil adult insect was or was not a galler.

FIG 221. The oldest fossil gall: the Pteriditorichnos stipitopterii petiole gall of the tree-fern Psaronius chasei (Marattiaceae). (a) The host plant P. chasei (centre) in its coal-swamp forest (original colour drawing by Mary Parrish in Labandeira & Phillips, 2002, with permission). (b) The upper part of the trunk of the host plant P. chasei showing one complete frond and the bases of several petioles; ungalled and galled sections of petiole bases enlarged (original by Conrad Labandeira, with permission). (c) A photomicrograph of the gall in section, showing the exit hole (lower left). (d) The section in (c) interpreted (original by Conrad Labandeira) (b, c & d from Labandeira & Phillips, 1996, 2002, with permission).

The oldest fossil gall known (Fig. 221) comes from the late Carboniferous of Illinois, USA, and is about 302 million years old (Labandeira & Phillips, 1996, 2002; see Table 11.1 for a summary of the geological time scale). The gall was found at the base of the petioles of a tree-fern, Psaronius chasei (family Marattiaceae), in a coal deposit containing tree-ferns, seed-ferns, lycopods and sphenopsids, and is older than the first conifers (Fig. 221a is a reconstruction of this tropical swamp forest). The tree-fern was about 7 m tall, with broad tripinnate fronds, and with swellings in the base of the petioles (rhachises) of the fronds (Fig. 221b).* The swellings, or galls, on average are 3.6 to 4.9 cm long (up to 24 cm long) and 1.0 to 2.5 cm wide and have been given the name Pteriditorichnos stipitopteris, not yet related to any insect. The structure of the gall is well preserved (Fig. 221c & d). It contains a central cavity excavated in parenchyma tissue, surrounded by a distinct layer of cells that are either enlarged or are smaller and grouped in patches, similar to callus tissue of simple modern-day galls. The lumen of the gall is packed with cylindrical frass pellets of several distinct sizes, suggesting that four instars of a larva developed inside before escaping through the exit hole that appears at one side of the gall. Because the larva has left the gall, the insect order cannot be certainly identified but the frass is typical of the chewing larva of a winged insect, like a beetle or sawfly (probably not a moth caterpillar as Lepidoptera are thought not to have evolved until the Triassic, 50 million years later). Similar galls are caused by sawflies in fern petioles today. This gall, 302 million years old, suggests that sawflies and other endopterygote insects (those with a pupal stage in their lifecycle) evolved earlier than previously thought (the first fossils of adult sawflies are known from the Permian, 290 million years ago). The same fossil tree-ferns show evidence of damage by piercing and sucking insects, probably aphid-like Hemiptera, but this damage is not associated with gall-like swellings. Other galls of similar age (Permian and Carboniferous) are harder to interpret. Some are swellings on the aerial roots of the tree-fern Lepidodendron and may be fungal in origin, although no fungal tissue is evident inside the swellings. They resemble galls caused by living Urophlyctis (Chytridiales) species, but they may be insect galls that are not well enough preserved to show their contents.

BOX 11.1. Classification and cladistics

Classification involves more than just naming plants and animals. The similarities and differences between species, and between groups of species, are used to place them in a hierarchy of groups. Hence, we talk about insects (class Insecta) within which there are orders (e.g. beetles, order Coleoptera), within which there are families (e.g. some weevils, family Apionidae), within which there are genera, within which there are species. In large orders and families and where knowledge is good, these groups may be subdivided (orders into suborders, families into subfamilies and, sometimes, superfamilies, etc.). Tribes and subtribes may be recognised between subfamilies and genera.

In principle, this classification should be ‘phylogenetic’: all members of a group should be more closely related to each other than to anything else; they should have a unique common ancestor, and all known descendants of that ancestor should be included. A group that meets these conditions is said to be ‘monophyletic’ (see Fig. 222).

In practice, since we do not have a complete fossil record, it can be hard to tell if what looks like a natural group really meets these conditions. As more species are discovered, and known ones are looked at more closely, opinions, and therefore classifications, change. There are two kinds of evolutionary change that make things particularly difficult, ‘convergence’ and ‘parallelism’. In convergence, two or more species with very different origins may evolve the same character(s) independently as a response to similar selection pressures (think of the leathery wings of pterosaurs and bats as an example). Their similarity in this respect does not reflect common ancestry. Parallelism is really a special case of convergence; amongst a group of related and rather similar species, several different lines of descent may follow the same trend (e.g. for increasing gall complexity).

FIG 222. A cladogram illustrating groups of hypothetical species 1–8 that are monophyletic at different levels, paraphyletic and polyphyletic.

In addition, some ancestral characters change so much, in different ways, that we do not at once recognise their common origin (fish scales and mammal teeth are derived from the same tissue source, for example). Hence, biologists distinguish between ‘homologous’ structures that have a common origin even if their functions are different, and ‘analogous’ structures that may have a similar function but are derived independently.

It is often the case that a classification has to be revised, because some groups are found to be either ‘paraphyletic’ (they have a single common ancestor but exclude some species that share that ancestor), or ‘polyphyletic’ (some members of the group have closer relatives outside it than within it). Historically, many cases of convergence and parallelism were detected by looking at the fossil record, and by studying embryology: adult features that are very different can be seen to originate from the same tissues during development of the embryo. However, there are many characters for which conclusive evidence is lacking, and classifications were a matter of deciding, in the event of a conflict of evidence, which characters were to be regarded as ancestral (homologous) and which convergent (analogous). Even faced with the same evidence, opinions on this have differed!

FIG 223. A cladogram for three hypothetical species of insect with primitive and derived states for two characters: mandibles and number of pairs of wings. The primitive character states are simple mandibles and two pairs of wings, and derived states are complex mandibles and one pair of wings. Species 1 and 2 share the primitive mandible character (symplesiomorphy, the blue bar), while Species 2 and 3 share the derived wing character (synapomorphy, the red bar); Species 3 has both derived characters. (Modified after Skelton, 1993.)

Two recent developments have helped to make the process of classification more rigorous, though they have not eliminated differences of opinion completely. ‘Cladistics’ is the name for a rigorous way of determining the status of characters, and for reaching the most ‘parsimonious’ conclusion about evolutionary change, i.e. the one involving the smallest number of independent evolutionary events (changes in the state of a character). The crucial distinction in cladistics is between a shared derived character (a ‘synapomorphy’ in cladistic terminology) possessed by two or more species or larger groups but not found in any of the others being considered, and a shared ancestral character of the group from which those possessing the derived character have diverged (a ‘symplesiomorphy’ see Fig. 223). Synapomorphies, correctly identified, enable subgroups to be defined within the group considered; symplesiomorphies, on the other hand, are no use in this respect, because they represent the ancestral condition for the group as a whole.

This is not the place to provide details of how the characters are assigned (there is a good introductory account in Skelton, 1993). Almost always, at least when many characters are considered, there are several possible phylogenies available to explain the observed set of characters considered. The cladistic approach is to choose the phylogeny that requires the smallest number of changes (note that the simplest solution is not necessarily the right one, but it ensures that the same criteria are being used in each case). Even then, there will often be convergences, cases where the same character has evolved more than once. When large numbers of characters are used, the analysis uses computer programmes that can provide information on the reliability or confidence with which each subgroup is distinguished. This information is not presented in this chapter, but it can be found in the original sources for the cladograms.

The results of a cladistic analysis are presented as a ‘cladogram’, expressing the relationships within the group considered as a branching tree. There are many cladograms in this chapter. It is important to note the difference between these and a conventional family tree, in which actual ancestors are given. Actual species (or larger groups) are always at the tips of the branches; you can work out what characters the presumed ancestor of a subgroup (or of the whole group) might have had, but the ancestors themselves are not represented. In principle, a cladogram should be a set of bifurcations (one branch should split into just two at any point), but where the analysis cannot distinguish between possibilities, several branches may diverge from the same point.

Although the terms ‘primitive’ and ‘advanced’ can be regarded as rather loaded (everything alive now is, in an important sense, equally advanced), they make sense when interpreted as having more (primitive) or fewer (advanced) symplesiomorphic characters, i.e. those characters present in the presumed ancestor of the group as a whole. The terms are used in this sense in this chapter.

The second important development is of ‘molecular taxonomy’. Now that it is possible to extract and sequence the genetic material, DNA, from organisms, it is possible to compare whole sequences of DNA from different species and note the similarities and differences. This has greatly increased the number of ‘characters’

available for analysis, and certainly needs the complex computer programmes that are used in cladistics. Usually, and especially for larger groups, molecular taxonomy has confirmed the conclusions based on morphological characters, but it has sometimes revealed cases of convergence and parallelism within families and genera that were not previously recognised. A strand of DNA consists of a series of ‘letters’ (bases) that pair with bases of its partner strand (each molecule of DNA consists of two strands wound into the famous ‘double helix’) and single changes in the sequence of these base pairs are relatively simple and they can happen rather often, and independently of one another. Hence a parsimonious approach is really necessary: often there are thousands of possible evolutionary pathways that could give rise to the pattern we see today.

The important thing to remember is that the cladograms presented here, some of which use molecular data, are representations of the most probable course of evolution in the groups concerned, as far as we know at present. Most of them will stand the test of time but classifications, and our understanding of the course of evolution, will be modified as new information becomes available and is analysed. What the cladograms do show, however, is that there are many cases of convergence and parallelism in the evolution of galls and their causers.

These very ancient galls existed before the evolution of flowering plants. The evolution of insect galls seems to have blossomed during the Cretaceous period (146–65 million years ago) when flowering plants were diversifying rapidly and when species of herbivorous insects in general were multiplying. By the Cretaceous, most families of insects and of flowering plants that exist today had appeared. Thus, although galling is ancient, galls were probably not common until flowering plants and insects diversified in the Cretaceous and later. Fossils of the most complex galls, caused by gall wasps (Cynipidae) and gall midges (Cecidomyiidae), are not known until the Tertiary: the oldest fossil gall midge is from the Oligocene, perhaps 30 million years old, and fossil gall wasp galls have not been found before the late Tertiary, although the oldest adult fossil is about 45 million years old (Eocene).

TRACING EVOLUTION–THE RELATIONSHIPS OF LIVING SPECIES

Although the fossil record can tell us about the minimum ages of some groups of insects, it is far too sparse to let us work out details of relationships, especially amongst closely related groups (species within a family, for example). In practice, most of our knowledge about evolutionary change comes from considering the relationships between living species. This is done using the similarities and differences between groups and species. Thus the classification of insects, and of other organisms, is intended to reflect evolutionary relationships–to be phylogenetic. In practice, different characters may suggest different relationships. Since there can only be one actual history, some characters held in common between, say, two species will be the result not of common ancestry but of convergent evolution: the same character has evolved more than once, independently (this is immediately obvious in the case of gall-formation itself).

Increasingly, solving this problem and tracing relationships has been done by the method of ‘cladistics’ (see Box 11.1). The aim is to ensure that all of the species (and genera, tribes, families, etc.) in a named higher group are monophyletic: sharing a common ancestor shared by no others. Within a named group, cladistics should also indicate which members are the most closely related. The end product of a cladistic analysis is a phylogeny expressed in a cladogram and it shows the most likely pattern of evolution in the groups considered. But cladograms are not fool-proof; later work, especially using DNA characters, may upset older conclusions, and the naming of species and higher groups does not always keep pace: some names are technically illegitimate because they represent paraphyletic or polyphyletic groups (these terms are explained in Box 11.1).

A number of insect groups in which gall causers are found have been subjected to cladistic analysis, using morphology or DNA or both. Some of these are used below to answer some of the major evolutionary questions raised by the study of galling.

THE EVOLUTION OF GALLING

The repeated evolution of a way of life is evidence that there must be particular advantages associated with it. But since not all, or even a majority, of insect herbivores cause galls, there must be some limitations as well. Although galls are composed entirely of plant tissue, their shape and structure seem to be dictated by the insect almost completely overriding the plant, and the insect gains all the benefits the gall provides. All advantage passes to the galler, which acts as a parasitic herbivore; the only exception in the insects is the unusual mutualism between figs and pollinating fig wasps (and there are a few examples of true mutualism in bacteria and fungi and their host plants). Romanes, in 1889, first suggested that galls evolved as adaptations of the insect rather than of the host plant. And, a 100 years later, Dawkins (1982) proposed that plant galls should be regarded as ‘the extended phenotype’ of the galling insect, controlled by the genes of the insect rather than the plant’s.†

Why do larvae of some species cause galls and not others? There are other herbivore lifestyles, like leaf mining, stem boring and free-living grazing and all, including galling, have benefits as well as disadvantages. Three suggestions may explain why galls evolved and why their shape and structure are so variable.

(1) Improved nutrition for the gall causer. Galls and their contained larvae are nutrient sinks: nutrients are drawn from elsewhere in the plant, diverted from buds, leaves, seeds or roots, and end up in the gall to be utilised by the galler. So the galler acquires a high-quality and guaranteed food supply. Enhanced nutrition is a general advantage of gall induction. It may also help to explain the variety of galls in some insect groups (particularly galls containing many individuals, as in aphids and thrips).

(2) Protection from a harsh environment. Insects enclosed in galls are protected from extreme climates: desiccation is likely to be a serious problem in dry climates or in areas subject to frequent fires. Gall structure in some groups may provide further protection, e.g. with thick woody or corky layers or coverings of waxes or resins, and spherical galls will lose less water than flatter galls of similar size.

(3) Protection from enemies. Although galls do provide protection from many general predators, they are targeted by specialists, especially parasitoids and inquilines. Gall-causing sawfly larvae and aphids, and gall mites, suffer less mortality than their free-living relatives but, in most other insects, enemy attack is greater on the gallers than the non-gallers. Because mortality of the gallers is often high, there should be strong evolutionary selection for increased protection, and there is compelling evidence, though not absolute proof, that some gall structures have evolved to protect the gall causer (e.g. in gall wasps, discussed below).

Any of these factors could enhance the survival and increase the number of offspring produced by the gall insect, and promote the evolution of galling. Which factors are the most important may vary between groups.

Although the genome of the insect is the main determiner of the size and structure of the gall, the plant and the environment around it can cause modifications: galls developing late in the season, or on older plants, or in unusually dry years, for example, are often smaller than early galls on young vigorous plants when there is no shortage of water. And the galling insect may not survive in small, underdeveloped galls.

Amongst the possible disadvantages of galling, those insects that cause galls tend to be very specialised, adapted to one type (often one species) of host plant. It is sometimes thought that galling, because it is such a specialised lifestyle, may be an evolutionary dead end. Evidence suggests this is not so; gall causers can increase their range and spread on to new host species and evolve into new species, and gall-causing can be lost and new species appear with new life strategies.

Modes of evolution in gall-causing insects

Apart from tracing the patterns of evolution from non-gallers to gallers (and sometimes back again), there are many other evolutionary questions to be asked about galling insects. Many gall-causing groups contain very closely related galling species. How did such speciation happen? Evolution of new gall species occurs in two main ways. First, a barrier may develop across the range of an ancestral species, dividing its distribution geographically and preventing individuals on each side from mixing together. Each subpopulation becomes isolated in environments that are different and will develop and evolve independently. Eventually they will become different species, unable to interbreed even if they did happen to meet. This is ‘allopatric’ speciation. Natural barriers develop over geological time, e.g. the building of mountain ranges or the separating of landmasses by plate tectonics.‡ Isolation of populations and their separate evolution can occur over the shorter term too. A species may accidentally or intentionally be introduced to a new country; if it can survive in its new environment, it will be isolated from its parent population and eventually will evolve into a new species.

Second, new species can arise ‘sympatrically’. An ancestral species may produce offspring that remain in the same geographical area but that can no longer mate successfully with the parent population. This can occur by host-shifting, the offspring moving to a new host species, or by organ-shifting, moving to a new organ on the same host (e.g. from bud to fruit, or leaf to root). A female makes a ‘mistake’ and lays an egg on the wrong host or in the wrong place on the host. If its larva manages to survive, become adult and reproduce, the resulting offspring may eventually become sufficiently different from the parent to be unable to breed with it, or they may be compatible but never meet because, say, their adults are active at different times. Organ-shifting may have occurred more often in the evolutionary history of gall insects as it involves less change for the new species than host-shifting, because the host plant remains the same. At present, allopatric speciation is the most generally recognised mode of evolution in animals and plants. But gall-causing insects, and some other insect herbivores, provide some of the best examples of the alternative sympatric mode and, as more information accumulates, this may prove to be equally common.

There can be big advantages in moving to a new organ or host plant. In the new place, food may be of better quality or last for a longer period. Or the new species may escape from old enemies or be better protected by the new gall. Or it may be better protected from a harsh environment. Or there may be more space and less competition in the new place. Similar advantages may also have lead to the evolution of alternation of generations (in gall aphids, gall wasps and a few gall midges).

Gall insects cannot survive without their host plants and cannot evolve independently of them. Did insects coevolve with plants, so that each in turn drove the evolution of the other (an evolutionary ‘arms race’ or ‘Red Queen’ scenario; explained in Chapter 10)? Or did the insects follow the plants so that, as new plant species evolved or even long after they had appeared, the gall insect shifted on to a new host species? It seems that both coevolution and host-shifting have occurred, with coevolution being much the rarer–the famous interaction between figs and their pollinating fig wasps (described in Chapter 7) provides the main example of coevolution.

Beyond speciation, there are other questions. Can we trace and explain changes in gall complexity and position? Can we account for changes in life histories and for the appearance of social organisation? In the rest of the chapter, particular well-studied case histories are used to explore these questions.

Aphids: complex lifecycles, the evolution of social organisation, and a unique inquiline

In aphids (defined broadly to include all in the superfamily Aphidoidea, see Table 4.4 in Box 4.5, Chapter 4), galling occurs in the more ancient families and subfamilies: in the Adelgidae and Phylloxeridae, in the Eriosomatinae (= Pemphiginae), Hormaphidinae and Melaphidinae of the Aphididae, and sporadically in more recent subfamilies of the Aphididae. Although the Aphidoidea is an ancient group of insects with a history of more than 200 million years, there is no evidence of aphid galls in the fossil record until the Cretaceous period. But fossils are only found occasionally; adelgid galls on conifers probably evolved much earlier. Adelgidae and Phylloxeridae split off from other aphids early and, in the Aphididae, the basal subfamilies Eriosomatinae, Hormaphidinae and Melaphidinae are considered to be primitive offshoots from the main aphid line. These three subfamilies are monophyletic: all three branches are related and evolved from one ancestral species (see Box 11.1). It seems that most of the gall-causing tribes in the Aphididae (Box 4.5) differentiated about 80 million years ago when aphids shifted from conifers to flowering plants. Gall-causing aphids are, therefore, relatively ancient and galling has been lost in the more recent subfamilies and tribes. Adelgidae remain restricted to conifers and Phylloxeridae gall more primitive flowering plant families: the willows Salicaceae, oaks and beech Fagaceae, pecan Juglandaceae and the grapevine Vitaceae, which are all woody plants. The main galling subfamilies in the Aphididae also cause their galls on the less highly evolved woody flowering plants. The best-known tribe is the Fordini (see below), which galls pistacias (Anacardiaceae). These are Mediterranean insects, and aphids in general are characteristic of the northern hemisphere and temperate climates.

Like oak gall wasps (described later) aphids can have complex lifecycles. Aphid lifecycles include egg-laying and viviparity (live births), cycling between sexual and parthenogenetic generations, and alternation between two host plant species (heteroecy; see Box 4.5 Chapter 4), and these complications occur in both galling and non-galling species. The evolution of the gall-causing aphids in general is considered first, and then the pistacia aphids (tribe Fordini) are described.

Cyclical parthenogenesis with two host plants seems to have appeared very early in the evolution of aphids, perhaps before 200 million years ago. The original ancestral aphid lifecycle was probably fairly simple: all females laid eggs, all were winged, males were common and all reproduction was sexual, and all individuals fed on one woody host plant species (they were primitively autoecious). Parthenogenesis and loss of wings may then have appeared, allowing a more rapid multiplication of females in summer when temperatures were favourable and food abundant. Early in aphid evolutionary history, woody plants would have been the most abundant food plants available: tree-ferns, early gymnosperms and conifers. The most ancient aphids, the adelgids, are still confined to conifers. Later, the first families of flowering plants appeared, also woody plants. The quality and quantity of sap in trees is relatively high in spring and early summer when the new leaves are growing but both decline during mid-and late summer. Perhaps galling, and the pineapple galls of Adelgidae on conifers, evolved as a response to deteriorating food supply so enhancing the quality of food for summer individuals.

Heteroecy may have been another solution to the problem of deteriorating food quality. Alternation particularly to herbaceous plants enhances food supply because these produce new leaves throughout the summer. Heteroecy occurs in several aphid lineages that are not closely related and it must have arisen independently at least three times: once when the Adelgidae evolved and at least twice in the Aphididae. The elm and poplar gall aphids switch to herbaceous hosts as food quality declines in summer and return to the tree to overwinter and to make use of the early spring growth, and this enhances reproductive output. The pistacia aphids alternate to their herbaceous hosts (grasses) in the autumn when food quality deteriorates as the trees lose their leaves, rather than in the summer. Pistacias are Mediterranean trees and are common in areas with very hot dry summers; in Israel, for example, the mild wet winter is a more favourable period for the growth of the grasses. Adelgids, which evolved before flowering plants appeared, alternate to a second tree, a conifer like the primary host, and so this advantage of heteroecy does not seem to apply to them. Whatever the cause of heteroecy, it may have been the step required for the early aphids, originally restricted to ancient primary host trees, to move on to new herbaceous hosts and to evolve into new species. Viviparity is also characteristic of aphids and evolved after the Adelgidae and Phylloxeridae had appeared–aphids in these two families lay eggs. Viviparity, along with parthenogenesis, telescopes the generations reducing the time required for each so that, when there is plenty of food and environmental conditions are good, reproduction rates can be very high.

Heteroecy in more recent aphid groups is uncommon–only 10 per cent of all living aphids alternates between two host species. Most aphids are secondarily autoecious, evolved from heteroecious ancestors and having lost either the primary or the secondary host. The loss of a host plant could have occurred in a number of ways:

By the loss of the primary host through the transfer of the sexual females and males, the egg and fundatrix to the secondary herbaceous host (the fundatrix is often the least adaptable morph but, in this scenario, it would have to be able to survive on the secondary host).

By eliminating migrants to the secondary host so that this host is lost and all generations remain on the primary host.

By eliminating the migrants returning to the primary host so that this host and the sexual and fundatrix generations are lost; and therefore the loss of sexual reproduction.

Cryptomyzus species (subfamily Aphidinae) illustrate several types of lifecycle. Some species are heteroecious, alternating between galls on a woody primary host (Ribes, currants) and secondary herbaceous hosts (mints, Lamiaceae) without galls. Some species remain all year on Ribes causing galls, having lost the secondary host. And others have lost the primary host and remain all year on mints without causing galls. The two Pemphigus species on poplars have different lifecycles too: P. bursarius has the full holocycle alternating to lettuces or other composites in the summer, while P. spyrothecae has lost the secondary hosts (their lifecycles are described in Chapter 5). In P. spyrothecae, the offspring of the fundatrix remain inside the gall and produce more offspring instead of migrating to the secondary host. Compared to P. bursarius, P. spyrothecae has fewer summer generations and so a reduced reproductive output, but avoidance of losses during migration to and from the secondary hosts probably compensates for this.

Loss of the sexual phase (anholocycly) has arisen independently in most aphid subfamilies and in both heteroecious and autoecious species. It is the result of a genetic change but can also be due to altered environmental conditions, like a change in day length. Tropical aphids, subjected to more or less equal day and night length throughout the year, are mostly derived from northern species and most are anholocyclic. Loss of sex resulting from loss of the primary host is common where year-round growth of the secondary hosts is possible, and where a cold-hardy stage is not needed–the egg, the result of sexual reproduction, tolerates cold weather better than the nymphs or adults. Some normally heteroecious species are anholocyclic in parts of their geographical range. Pemphigus bursarius, normally alternating between poplars and lettuces, sometimes survives permanently on lettuce roots. Forda formicaria and F. marginata are found on the roots of grasses in North America, Central Europe and New Zealand, well outside the range of their pistacia primary hosts. Some of these populations may be relicts that have survived after the Pliocene or Pleistocene glaciations eliminated their primary hosts in more northern areas. It seems that very few anholocyclic species have diverged much from their sexual ancestors, but loss of sexual reproduction may have occurred too recently for there to have been sufficient time for much diversification to have occurred.

Aphids of the tribe Fordini (Aphididae, subfamily Eriosomatinae) cause galls on the leaves of Pistacia: in the margin, the leaf blade, the veins or the petiole. In Israel where they have been studied intensively, there are three host species, Pistacia palaestina, P. atlantica and P. lentiscus, with overlapping distributions and each aphid species is restricted to one of these as its primary host. About 16 species occur in Israel, all of them causing pouch galls with a similar structure but varying dramatically in size (see Chapter 5). The simplest are pea-shaped, about 5 mm across and attached to the mid-vein of the leaflet, and are unsealed. Margin galls are elongate, about 20 mm long in the leaflet margin, and are also unsealed. Bag-like galls on the upper surface of a leaflet are larger and occupy part or most of the leaf surface, and have a permanent opening near the mid-vein. The largest galls, with volumes of about 4 cm³, hang down from the underside of the mid-vein or arise from buds, and are spherical, banana-or cauliflower-shaped. They are sealed until ripe, when they split open to release the aphids. Bud galls arise from a very young leaflet still inside the leaf bud and eventually take over the whole bud. Most pistacia aphids cause just one type of gall and have a two-year cycle with two host plants (the holocycle). Four species cause two types of gall: the fundatrix gall is a small pea in the mid-vein of the leaflet and her offspring cause larger margin galls. Slavum wertheimae is unusual with only one host, P. atlantica, and has a lifecycle of one year.

FIG 224. Cladogram for Israeli pistacia aphids (Fordini) indicating the type of gall produced; the host Pistacia for each species is shown in the right-hand column (based on DNA analysis; modified after Inbar et al., 2004). Horizontal dashed lines separate the three groups in the tribe. Two new Geoica species and sp. A and sp. B in the Forda-group have recently been recognised but have not yet been named. The line symbols indicate the type of gall produced.

DNA analysis indicates that the Fordini of Israel all evolved from a single gall-causing ancestor (they are monophyletic; Fig. 224). Anatomical character analysis supports this. The tribe divides into two clades: the Forda–group (the six species between the dashed horizontal lines) and the Baizongia–group (the seven species above the upper line). Smynthurodes betae, previously thought to be closely related to the Forda species, is ancestral to all the other species–it evolved before the split that divided the Forda– and Baizongia–groups. The clusters of species in the cladogram do not correspond to the host plants. Each aphid galls only one host species and several less closely related aphids gall the same host. This suggests that aphids have shifted between Pistacia host species several times, each time evolving into a new species. The winged females, the sexuparae, ‘choose’ the primary host when they migrate from the roots of grasses, the secondary hosts; they can recognise Pistacia trees and avoid landing on other trees, but they do not always recognise the correct pistacia species. They give birth to sexual females and males on whichever tree they have alighted upon, and these mate and produce the egg successfully. The problems arise when the fundatrix hatches from the egg–she may be able to start a gall but on the ‘wrong’ host it usually does not mature and her offspring inside do not survive. The fundatrix is the least adaptable morph. Occasionally, though, the fundatrix does succeed and the opportunity is there for a host shift; perhaps, as in the willow sawflies (described later), host-shifting is not uncommon in pistacia aphids. The step is probably easier via a hybrid between the original and the new host and hybrids often carry galls characteristic of both parental species. Shifts between P. atlantica and P. palaestina have occurred twice in the Forda–group and at least twice in the Baizongia–group. But only one species, Aploneura lentisci, galls P. lentiscus; shifting to this host, which is not as closely related to P. atlantica and P. palaestina as they are to each other, is more difficult. Perhaps this is because P. lentiscus produces large quantities of resin and is chemically different from the other two trees.

The types of galls, whether pea, margin, bag, spherical or bud galls, cluster more closely than associations with the host plants (Fig. 224): closely related species cause similar galls but on different hosts, indicating that the aphid rather than the plant controls the shape and the position of the gall (the gall is the extended phenotype of the aphid). Sealed galls evolved only once, in the Baizongia–group. The ancestral pistacia aphid probably induced a small pea-shaped gall on the mid-vein of the leaflet. The two–gall lifecycle, with the pea gall of the fundatrix and the margin gall of her offspring, seems to have appeared early, before the Forda–and Baizongia–groups split apart. It occurs in S. betae and is retained in the three Forda species but has been lost from the Baizongia–group. The bag galls of Asiphonella dactylonii and Aploneura lentisci probably developed from simple peas, with spherical galls developing from bag galls. The bud galls, which grow large enough to take over the whole leaflet, probably also developed from very young pea-shaped galls but along a different pathway. The strength of the nutrient sink induced by each gall increases along this sequence (i.e. from pea ‹ margin‹ bag ‹ spherical ‹ bud galls) and is reflected in the reproductive output of aphids within the gall. The smallest galls draw nutrients from adjacent parts of the leaflet only and support about 20 aphids. The increasing sizes of the margin, bag, spherical and bud galls affect increasing proportions of the host plant: the whole leaflet, the entire leaf with all of its leaflets, to all of the leaves on a shoot and even on neighbouring shoots. The numbers of aphids in large galls of Baizongia pistaciae and Slavum wertheimae can exceed 10,000. Evolution in aphid galls has involved an increasing area of feeding surface, and this has enabled the aphid colony to increase dramatically in size.

Some aphid galls are very large and enclose large numbers of individuals. The gall is initiated by the fundatrix, a first instar female nymph (see Chapter 4 and Box 4.5). When mature, the fundatrix produces large numbers of offspring that live and feed together in the gall. The gall enlarges as numbers increase; one of the main reasons for increase in gall size is to provide a greater area of feeding surface inside. If many related individuals live together in a nest or other confined space, cooperation and apparently altruistic social behaviour can evolve. Particular individuals carry out tasks that benefit others in the colony, such as getting rid of wastes or defending the gall. These individuals are often sterile or nearly so, unable to produce offspring or producing fewer than their sisters. The ‘altruism’ is less selfless than it appears: the individuals are identical clones of the fundatrix, so a sterile ‘aunty’ is related as closely to her sister’s offspring as to her own if she had any. Some individuals become soldiers and defend the gall from intruders and usurpers. Others carry out housekeeping duties, removing dead bodies and cast skins, and the aphids’ copious honeydew that would otherwise foul the colony. Some soldiers carry out both jobs.

In species with soldiers, these are tougher and have weapons not found in the non-soldiers. Nearly all aphid species that have soldiers are gall causers, although not all galling aphids have soldiers. In two aphid subfamilies, the Eriosomatinae and the Hormaphidinae (Box 4.5), both with many gall causers, social behaviour has evolved several times: in the elm aphids (Eriosomatini), in poplar aphids (Pemphigini), in the Cerataphidini and in the Nipponaphidini (Table 11.2), but it has not evolved in the Fordini. First or second instars develop into soldiers. They are active individuals with tough skins and with an array of weapons, such as spiny legs and horned heads as well as stabbing stylet mouthparts. They congregate around the entrance to the gall and readily attack enemies at the entrance or on the galls’s outer surface and, sometimes, inside it (see Chapter 10 for some examples). The more vulnerable morphs, those that reproduce or are migrants, tend to be found in the safer depths of the gall. Some individuals that are sterile never grow up into adults (and may not feed). Others are soldiers only when young and grow into normal breeding adults. Some species are ‘dimorphic’: they produce two types of first instar nymphs, soldiers and non-soldiers. Gall-cleaning, the removal of honeydew, cast skins and dead aphids, is essential for the survival of the aphids and may have been a precursor of the defensive behaviour of soldiers, particularly in galls that tightly enclose their aphids. Fighting between first instar fundatrices for the best galling sites may also have been a precursor, with the offspring of the fundatrix developing her weapons and kicking and shoving behaviour.

TABLE 11.2. Gall-causing aphids with and without soldiers (modified after Foster & Northcott, in Williams, 1994, using the classification of Blackman & Eastop, 1994, as in Table 4.4, Box 4.5)

Because of the excess of sugar sap in their diet, most aphids produce waste honeydew that needs to be disposed of, particularly important in the enclosed space of a gall–aphids could drown in their own waste. All gall-causing aphids produce wax to parcel and transport honeydew and it is clear that wax is essential to the survival of aphids in galls. Special cells concentrated in the upper epidermis of the abdomen and around the anus produce the wax so that, as each drop of honeydew emerges from the anus, it is coated with wax strands and further covered with wax powder as the drop rolls around in the gall. So the drop becomes spherical and non-wetting and can easily be pushed out of the gall. In Pemphigus spyrothecae galls on poplar leaf petioles, ‘marbles’ of honeydew are rolled out of the gall by pushing and shoving by soldiers (see Chapter 5) and, in a large colony, up to 1 cm³ can be removed per day. If removal is prevented, 90 per cent of the aphids will die in a few days. Probably the ancestor of aphids, very early in their evolution, produced wax and this may have allowed galls to evolve. Wax production is common in free-living aphids too but has been lost in some groups; it seems that wax glands may have been lost and appeared again several times in more recent aphid groups.

Evolution of soldiers may have occurred just once in an early aphid ancestral to the related subfamilies Eriosomatinae and Hormaphidinae, and then been lost in the many genera that do not possess soldiers (Table 11.2). Or they may have evolved later more than once, when the tribes with soldiers were differentiating into genera. The second possibility is the more likely because the morphology and behaviour of soldiers is different in the different tribes. The weapons of Colophina and Hemipodaphis (Eriosomatini) include enlarged and toughened fore-and midlegs, while in Pemphigus (Pemphigini) the hindlegs are enlarged. In the Cerataphidini, there are at least two distinct types of soldiers: one with horns on the head and with greatly thickened forelegs with claws, and the other with stout hairs on the head and without enlarged forelegs. In both types, some soldiers are also sterile. The evolution of soldiers several times, independently, would emphasise their biological significance, that soldiers benefit the whole colony despite their cost. The weapons and butting/kicking behaviour of the soldiers could have evolved gradually, at first to remove honeydew or to dislodge siblings and rivals of the same species from preferred feeding sites. Later, they would stab enemies or push them or their eggs out of the gall. Later still, soldiers of some species became sterile, cleaning and defending their galls throughout their lives and, in a few Cerataphidini species, soldiers have evolved in the non-galling colonies on the secondary host as well as in the galls on the primary host.

Galls containing a clone of identical individuals, as in aphid galls, are an ideal place for the development of this apparently altruistic behaviour, especially in enclosed galls in which cleaning and defence are so necessary. But this social behaviour is rare in aphids as a whole–why is it not universal or at least more common? The answer may be that cheats can undermine it: if unrelated aphids enter the clone and remain undetected, they will benefit from the ‘altruism’ but make no contribution in return. In an enclosed gall, a soldier can be confident that its neighbours are clone-mates and so are genetically closely related. But outside the opening and on the surface of the gall, any aphids are probably not siblings and so are worth attacking. Soldiers develop in relatively long-lived galls that remain open for a considerable time: the Pemphigus spyrothecae gall lasts five months and has soldiers while the shorter-lived P. bursarius gall has none. Eriosoma and Pachypappa galls are looser and more open and individuals probably regularly migrate between galls, and some of these migrants might include cheats. Even so, some species have soldiers while others do not.

A small and unusual aphid group, unrelated to any other gall-causing aphids, has developed inquilines, though no soldiers–the Tamalia aphids (Aphididae: Tamaliinae; Table 4.4, Chapter 4). These cause galls in the leaves or flower heads of Arctostaphylos and its relatives (Ericaceae) in North America. There are four known gall-causing Tamalia species and one inquiline in the same genus: this is T. inquilinus (which is probably a species–complex of several closely related sibling species). The best known galling species is T. coweni, an unusual aphid in that both the fundatrix and her granddaughters cause galls. These females are wingless while alternate generations are winged and disperse to other sites. The one sexual generation has winged males and females that disperse before mating and laying eggs, and these eggs hatch into the fundatrices. Galls of T. coweni often contain females of both gall-causing generations–they share the gall apparently amicably. Cohabiting perhaps has evolved in Tamalia because both galling females are clone-mates, and this might represent the start of a type of social behaviour. It contrasts strongly with the highly territorial behaviour of Pemphigus fundatrices, which are not clone-mates. Inquiline Tamalia are well synchronised with development of the host’s gall. Mated T. inquilinus females lay eggs in the spring and the first instars invade the young galls of the host aphid before the galls become sealed; the inquilines mature and produce daughters, some of which are winged. These disperse and produce more daughters, which invade more galls when they open to release their winged migrants, and eventually winged sexual females and males are produced that mate to start the lifecycle again. The inquiline and host aphids tolerate each other without fighting, even though the inquilines are tougher-skinned and hairier than their hosts.

Evolution of inquilines that are closely related to the gall causers is almost unknown in aphids, although in gall wasps it is common (see below). The Tamalia inquilines, it seems, have evolved very recently. The gall-causing Tamalia are also not ancient. Although their host plants are widespread, known from across the northern hemisphere, the aphids are restricted to the Nearctic region; their absence from the Palaearctic suggests an origin around 10 million years ago, when North America split from Europe and the northern Atlantic widened. Tamalia first galled Arbutus, then shifted to the related Comarostaphylis and finally to Arctostaphylos, and it is on this host that the first inquiline Tamalia appeared. Inquilinism opened up a new opportunity for these aphids. Analyses of mitochondrial DNA indicate that inquilinism evolved only once and arose from a gall-causing species. The inquilines have diversified rapidly and recently, over only the last 2.4 million years, and no doubt they continue to diversify today. The inquilines are developing host races or sibling species along with their host plants rather than in parallel with their host gallers: host plants and inquilines seem to be coevolving, each driving the adaptation of the other. On the other hand, the Tamalia gall causers have not coevolved with their host plants. The gallers have apparently diversified more slowly than the inquilines and have evolved new species by periodically shifting to new hosts. Why should the Tamalia inquilines have evolved differently from the Tamalia gall causers? It may be due to the opportunity for invasion of a gall by the inquiline; the opportunity ‘window’ is brief and restricted by development of the plant, requiring the inquiline to be completely synchronised with the plant. In contrast, the window available for initiation of the gall may be wider, so that timing between the galler and the plant is less critical.

Diptera: three-way galls and the evolution of flower head galls

The Diptera, one of the largest insect orders, includes the gall midges (Cecidomyiidae), a group that contains more gall-inducing species than any other insect family. The first Diptera appeared in the early Triassic, about 250 million years ago (Table 11.1), after the first beetles and sawflies had evolved, but probably none of these Diptera were gall causers. The earliest known fossil cecidomyiid (a species of Catotricha, probably a fungus feeder) dates from late Jurassic or early Cretaceous deposits, about 150 million years old (see Box 4.8, Chapter 4). The family probably evolved during the late Permian and early Triassic along with related families, the fungus gnats (Sciaridae and Mycetophilidae) and, more distantly, the crane-flies (Tipulidae). Adult cecidomyiids are typically small delicate flies with long, many-jointed antennae, reduced wing venation and relatively long legs.

Many adult gall midges have been found trapped and fossilised in amber. Amber is fossilised resin produced by conifers to seal and protect wounds–as it oozed down the tree trunk it engulfed small insects that probably were attracted to the flow of sap from the wound, and preserved them perfectly. These fossils, which include representatives of existing gall-inducing genera such as Cecidomyia, Contarinia, Dasineura and Lasioptera, are mostly from Tertiary deposits of the Eocene and Oligocene, 55 to 25 million years old, and so are comparatively recent. The oldest known cecidomyiid galls in the fossil record are more recent than that and include Mikiola fagi on beech (Fagus sylvatica; described in Chapter 5, Gall midge pouches), recorded from the Pliocene (2–5 million years ago) in France, and Sequoiomyia kraeuseli recorded on Sequoia langsdorfii (Taxodiaceae), also Pliocene, found in Germany. So, although Cecidomyiidae is one of the most frequently encountered families of Diptera in fossil deposits, there is virtually no fossil evidence of the transition from fungus-to plant-feeding and, later, to gall-inducing; this must have occurred in the second half of the Cretaceous period, from about 100 million years ago, at the same time as the major evolutionary expansion of the angiosperms.

Only the most advanced subfamily, the Cecidomyiinae, includes gall causers. Phylogenetic studies of the Cecidomyiinae based mainly on morphological characters of adults and larvae are not yet fully developed (and DNA analyses are not common) but they do provide some indications of evolutionary trends. The subfamily is currently divided into four supertribes (see Table 4.8, Box 4.8, Chapter 4), but only two, the Cecidomyiidi and Lasiopteridi, include gall causers. Each supertribe includes several tribes with gallers (Table 4.8) and it is clear that gall-causing has arisen independently many times. But the transitional stages between a non-galling and a galling gall midge species are not fully understood.

The Cecidomyiidae were, probably, originally closely linked with woody plants in forests where the larvae fed on fungi on decaying organic matter. Then, following the rapid evolution of flowering plants during the Cretaceous, they became able to feed directly on living tissues of plants. Two major routes for this adaptation have been suggested. First, via wounds: the larvae could have invaded wounds in trees along with fungi and fed on the fungi and on sap exuding from the dying tissues. Such mixed feeding has been termed ‘phytomycetophagy’ and is typical, for example, of living species of the non-galling genera Resseliella, Ledomyia and Clinodiplosis, and of Lasioptera, which is particularly associated with the development of stem galls. A second possible route involved larvae invading the increasingly diverse and complex flowers of angiosperms. Here they perhaps fed initially on fungi and liquid products of decay (i.e. also phytomycetophagy) before adapting to feed on the sugary sap of nectaries and on sap from other flower parts and, later, on the tougher tissues of ovaries, fruits, buds and other plant organs. Many such associations with angiosperm flowers still exist, especially in major gall-inducing genera, such as Contarinia and Dasineura, and this evolutionary path may have resulted in the development of the tribes Cecidomyiini, Asphondyliini and Oligotrophini. Evolution of the whole supertribe Cecidomyiidi has been mainly associated with the angiosperms; relatively few species use ferns or conifers as hosts and those that do are probably of recent origin. On angiosperms some genera such as Dasineura and Contarinia have hosts in many unrelated plant families whereas other genera, such as Rhopalomyia on Anthemideae (a tribe of the Asteraceae), Rabdophaga on Salix, Caryomyia on Juglandaceae, Planetella on Cyperaceae, Psectrosema on Tamarix, Mayetiola and Orseolia on Poacaeae, are restricted in their host ranges. Evolution by host-shifting appears to be the main route by which new species evolved but knowledge about these radiations is still inadequate; it may improve as more DNA analyses are carried out and the molecular taxonomy of gall midges becomes better known.

Amongst the gall causers, some larvae cause ambrosia galls, unique to the gall midges, and these may still be dependent on fungi (several examples are described in Chapters 8 and 9). The young larva and the fungus are present at gall initiation and in some species at least, both are needed to initiate the gall. The larva may feed to some extent on hyphae that grow to line the gall chamber, or on sap from both plant and fungal cells, or only on the plant cells. This resembles the primitive condition of phytomycetophagy but it is probably a secondary adaptation, especially as ambrosia galls are known in three tribes, the Asphondyliini, the Alycaulini and the Lasiopterini (Table 4.8), none of which are closely related. In some ambrosia galls, the interaction between larva and fungus may be less intimate; the fungus may behave as an inquiline in the gall and have little effect on the larva.

Ambrosia galls involve a symbiosis between the gall midge and a fungus. The fungus, with the gall midge larva, penetrates the plant so initiating the gall, and grows to line the gall chamber, absorbing nutrients from the plant. The larva in the chamber may feed on the fungus (although in no case is this proven) and may be protected by it from parasitoids. The fungus also depends on the gall midge to transport it to new plants each year. The fungal symbionts in most ambrosia galls seem to belong to only one or a few closely related species; their identity is difficult to sort out as they seldom produce spores. In contrast, the pouches (mycangia) in which the female midge transports spores and hyphae to a new host plant are different in each of the ambrosia gall midge tribes, indicating independent origins of the trait. It may be that a fungus first caused plant cells to differentiate into a gall that was exploited by the fungus-feeding gall midge larva, and that gall midges later developed the ability to cause galls on their own. Now, it seems, the ambrosia fungus cannot cause a gall without the help of the gall midge.

FIG 225. A cladogram for Japanese and some European Asphondylia species (based on DNA analysis; modified after Uechi & Yukawa, in Osaki et al., 2006: details for the related genus Pseudoasphondylia are not included). Note that several species have not yet been named. The life history patterns 1–5 are described in the text.

Asphondylia is a large genus (about 270 species are known worldwide) in the tribe Asphondyliini, which is a monophyletic group. Many species with ambrosia fungi cause galls in flower buds, where they prevent normal fruiting. Most species, like most gall midges, are specific to one or a few related host plant species, but a few are able to gall hosts in several plant families. Although they all evolved from one ancestor, Asphondylia species illustrate a variety of lifecycles and habits, with some alternating between two organs on the same host or between two host species (reminiscent of some aphids and gall wasps but unusual in Diptera). Fig. 225 is a cladogram for a group of Japanese and European Asphondylia species; it indicates that these ambrosia midges are not constrained by the type of lifecycle or by the part of the plant galled, or by family of host plant: these characters vary in both the more primitive and the more advanced species. There are five types of life history in this group:

(1) Univoltine (one generation a year) and mono-or oligophagous (specific to one or a few closely related host plant species). The first instar larvae spend summer, autumn and winter in the gall, they develop rapidly in the following spring and adults emerge in early summer when the new plant organs, flower or leaf buds or young fruits, are available for galling.

(2) Bivoltine (two generations a year) and mono-or oligophagous. The gall midge alternates between two organs on the same host species. The British and Japanese species A. sarothamni is an example, alternating between leaf buds and young pods on broom Cytisus scoparius.

(3) Multivoltine (many generations a year) and monophagous. The gall midge has several generations galling young fruits that are available throughout the year. An unusual lifecycle, known for only one species in Japan.

(4) Multivoltine and polyphagous (with hosts in several plant families). The midge has several quick generations in spring and summer on hosts that die back in autumn and it winters on an alternate host. A. gennadii occurs in Europe as well as Japan and galls hosts in four families.

(5) Most individuals of the gall midge species are univoltine and oligophagous but some are bi-or multivoltine and utilise alternate hosts. An unusual lifecycle with only one species of Asphondylia known in Japan.

Most gall midges are restricted to a particular host and are tightly synchronised with its growth, so that the initiating hatchling is penetrating a bud or other part at exactly the correct stage of development. Asphondylia may be less constrained than most gall midges because of the fungal symbiont, which probably can grow on slightly older as well as on very young plant cells. This flexibility may have allowed a variety of strategies to evolve on a variety of hosts, and has resulted in the appearance of many closely related species. Asphondylia adults are relatively strong fliers too. Many gall midges are very fragile and are blown about passively by the wind, but Asphondylia females can direct their flight and search out suitable host plants. Polyphagy and multivoltinism increase the reproductive potential of a species, they increase the proportion escaping from enemies and may enable more nutritious plants to be galled. Speciation and shifting to new hosts in Asphondylia is probably a fairly recent phenomenon and has allowed (is allowing) large numbers of species to evolve.

Apart from the gall midges, gall causers are found in six other families of Diptera, all robust and bristly, and belonging to the Brachycera: Cyclorrhapha (the ‘higher’ flies), though the number of galling species is relatively small. Cyclorrhaphan flies seem to have arisen in the Jurassic and to have diversified in the Cretaceous (Table 11.1). Amongst those causing galls, only the Tephritidae are well known; it is thought that tephritid galls evolved relatively recently, about 25 million years ago.

Galls caused by the Tephritidae range from simple accumulations of callus to complex woody galls with an elaborate structure (see Chapters 7 and 8). Most of their galls are associated with host plants in the Asteraceae, particularly thistle-type plants (e.g. burdocks, thistles and knapweeds) in the subtribes Carduinae and Centaureinae of the tribe Cardueae. The flower heads are particularly rich sources of food for insects and a variety of larvae lives and feeds inside them, and some of these cause galls. These insects group into characteristic ‘guilds’. The most complex guild is composed of three sets of herbivores:

(1) Gall causers that exploit the young flower head and manipulate its vascular system, drawing nutrients to the head from elsewhere in the plant and creating a strong nutrient sink. Some tephritid flies (and gall wasps, Cynipidae) fill this role.

(2) Larvae that attack the flower head slightly later and mine through the receptacle and immature achenes (seeds). Non-galling tephritids and weevils are the commonest insects in this category.

(3) Larvae that attack the flower head after flowering and feed on mature achenes as well as the rest of the head; these include opportunistic moth caterpillars that become predators, killing and eating any of the earlier species that they come across.

In addition parasitoids and predators attack the herbivorous larvae and often specialise on one or other of these three categories. The guild structure is complex in thistle flower heads all over the Palaearctic region, from Britain to Japan. But it is less complex elsewhere and in related host plants. In the Nearctic, there are no complex galls in flower heads: the first group is missing. This may be because Cirsium and related plants (in the Carduinae) in North America have evolved and diversified only recently, in the Pliocene and Pleistocene over the last 5 million years, while in the Palaearctic, the thistle and knapweed subtribes of the Cardueae radiated in the late Oligocene or early Miocene, 20–30 million years ago (Table 11.1). Other subtribes of the Cardueae and related tribes with similar flower heads are less rich too–there are no gall causers in carline thistles (Cardueae: Carlineae) or in globe thistles (Cardueae: Echinopeae), nor in the small flower heads of goldenrods (tribe Astereae) and other species. Berkheya species (tribe Arctotideae) are the native thistles of southern Africa and many species have large flower heads. They have not evolved galls either, although some species have a rich flower head guild of non-galling tephritids and weevils.

FIG 226. Preliminary cladogram for the tribes of the subfamily Tephritinae (Tephritidae); those that include galling species are indicated with red lines (based on morphological characters; modified after Korneyev et al., in Raman et al., 2005). The tribes above the dashed horizontal lines are lower Tephritinae and those below are higher Tephritinae.

Only about 5 per cent of the 4300 described Tephritidae cause galls and nearly all belong to the subfamily Tephritinae (see Box 7.1, Chapter 7). The tribes of the Tephritinae are divided into lower and higher groups and galling has evolved in several of them (Fig. 226). The higher Tephritinae are monophyletic: the five tribes evolved from one ancestor (see Box 11.1). The lower Tephritinae are polyphyletic, probably with two ancestral species; the Terelliini is basal to the rest and is less closely related than the other tribes are to each other. The Terelliini are non-gallers except for Terellia tussilaginis, a borderline galler that causes slight swelling of the achenes in burdock heads (see Chapter 7, Tephritid Galls). The Cecidocharini is intermediate between the lower and higher groups and includes some gallers. All of the tribes in the subfamily include non-galling species and, in those tribes that also have gall causers, the latter have arisen independently. So, galling is not an ancestral trait in the Tephritidae. In the non-gallers, the females lay eggs into or between the florets of open flower heads; the more primitive species have blunt ovipositors, not adapted to penetrating deep into plant tissue. In Urophora (tribe Myopitini) and other gall causers, and in the more advanced non-galling species including the leaf-mining species in the subfamily Trypetinae, the ovipositor is needle-like or a fine saw and penetrates deep into flower heads or into leaf buds to insert the eggs. Feeding behaviour differs too between galling and non-galling larvae: in the more primitive non-galling species (e.g. most Terellia, Xyphosia), larvae excavate cup-like depressions in the receptacle without stimulating extra tissues and drink the sap that collects in the cups. The gall-causing larvae are more specialised and feed on nutritive tissues (see Chapter 7).

FIG 227. Cladogram for genera in the tribe Myopitini (Tephritinae) (based on morphological characters; simplified after Korneyev et al., in Raman et al., 2005). Red lines indicate that most species in the genus cause galls. *One species belongs to this Asimoneura genus and causes a simple gall; it is not closely related to the rest of the genus and no doubt it will be renamed in the future.

Most gall-causing tephritids belong to the tribe Myopitini (Fig. 227) and cause a range of galls from slight swelling of achenes to complex woody galls. Achene-feeding is thought to be the primitive habit in the tribe. The Eurasimona–Goedenia clade probably is basal to the rest of the tribe; Eurasimona larvae are found in flower heads of Anthemideae in Europe and Asia, and Goedenia larvae live in Solidago in North America. Spinicosta larvae feed in the large achenes of Berkheya and Vernonia in southern Africa, but they do not cause galls. Most Urophora, Stamnophora and Myopites cause conspicuous woody galls, though they differ in structure. Stamnophora larvae in Vernonia flower heads live in separate chambers and the adults emerge through a common exit that forms a ‘chimney’ extending above the achenes. Some Stamnophora species cause stem galls. In Myopites and Urophora, each larval chamber has its own exit, also with chimneys in Myopites. Myopites attacks flower heads of Inula and Pulicaria (tribe Inuleae) and Urophora attacks thistles and knapweeds in the Cardueae, and both tephritid genera are native to Europe and Asia.

There are 60 species of Urophora worldwide and their galls vary, from simple achene swellings to complex woody galls, usually in the flower heads but with two species in stems. Fig. 228 shows the relationships between the species–complexity of the galls increases down the diagram. The ancestral species probably mined into mature achenes without enlarging them; this is a common habit in larvae of related genera, e.g. Spinicosta (Fig. 227). It requires precision in placing the eggs and an elongated ovipositor, both useful pre-adaptations for a future gall causer. From this stage, a simple non-woody achene gall evolved, as is found in the U. quadrifasciata–group of species on knapweeds Centaurea (see Chapter 7 and Fig. 110). Although only a simple gall, the achene is enlarged and patches of nutrient tissue develop at the base of the larval cavity. The larva acquires more food than does a simple miner and grows larger than related non-galling larvae. The next step in gall complexity is the woody cup-shaped gall, formed of tissues of the receptacle as well as the achene and caused by the U. affinis–group (Fig. 228 and Fig. 110). The gall of U. affinis itself (not a British species) is a single cup while in U. jaceana, the woody tissues of several cups usually fuse together and incorporate more of the receptacle. Nutritive tissue is more extensive than in the U. quadrifasciata gall, developing first in patches along the wall of the chamber and then more extensively at its base. Most species in the U. affinis–group gall knapweeds. The most highly evolved and complex Urophora gall in flower heads is the block-shaped gall typified by U. stylata in spear thistle Cirsium vulgare in the U. cardui–group (Fig. 228 and Fig. 110). Young larvae chew down florets and feed on the ovary in a similar way to U. jaceana and its relatives. Then the larvae mine more deeply into the receptacle so that more tissues are affected, resulting in a larger gall. Probably these block-shaped galls have arisen independently several times in Urophora–they are found at several points in the cladogram. They are particularly distinctive in the U. cardui–group specialised on Cirsium: U. stylata, U. congrua and U. cardui. U. solstitialis on Carduus, U. terebrans on Cirsium and U. mauritanica on Carthamus cause similar galls.

FIG 228. Cladogram for Urophora; the tip of each branch represents a single species and selected species are identified (based on morphological characters; modified after Korneyev et al., in Raman et al., 2005).

The complex galls of the more specialised species show advantages over the less complex but there are costs too. U. quadrifasciata is more successful in some environments than the more highly evolved U. jaceana, even though both can attack the same host plant. Table 11.3 summarises the costs and benefits of the two species’ lifestyles. The more specialised U. jaceana has a higher fecundity (it produces 2½ times as many eggs) and can adjust the size of its egg clutch to the quality of the host’s flower head. The larvae extract more energy and nutrients from the plant (the gall creates a stronger nutrient sink), and this results in larger larvae and larger adults that lay more eggs. The disadvantages for U. jaceana are a longer developmental time caused by the more complex gall and its dependence on only two host species. U. quadrifasciata has a broader host range and develops faster so that two generations a year are possible. It also is more active and can disperse over long distances to find new host plants–it is therefore a more successful coloniser of new habitats. In stable predictable environments U. jaceana has the advantage but in variable and changing conditions, U. quadrifasciata does better. The woody gall of U. jaceana provides some protection from enemies, at least from generalist predatory caterpillars (see Chapter 10), but not from specialist parasitoids.

Amongst the Urophora gall causers, two closely related sister species, U. cardui on Cirsium arvense and U. misakiana on C. setidens in Korea, cause block-shaped galls in the plant stems. This appears to be a late development in Urophora, and there is evidence that they evolved from species inducing flower head galls, perhaps because gall size was limited in the small flower heads of their host plants. Another tephritid genus Eurosta (in the tribe Dithrycini, Fig. 226) causes stem galls in goldenrods Solidago species, in North America. This tephritid–plant interaction has been studied in great detail (the fly and its galls are described in Chapter 8). The studies provide examples of host-shifting and of the effects of enemies upon the tephritid.

The host plants of Eurosta solidaginis are Solidago altissima and S. gigantea, closely related sibling species and members of the Solidago canadensis species complex. The plants have overlapping distributions across eastern and central North America from southern Canada to Florida and Texas. E. solidaginis infests S. altissima throughout its range but is found on S. gigantea only in the north, in southern Canada and northeast USA. In areas where the tephritid is found on both hosts, the flies form distinct host races–each race attacks only its own species. Host races form because the flies are reproductively isolated: they emerge from their galls at slightly different times, they mate only with males or females that have emerged from the same host species, and they prefer to lay their eggs in the same host species that they emerged from. Hybrids between the fly races can be produced in the laboratory and are fertile, but they do not perform well; they are less able to cause galls and mortality is higher than in pure flies and, probably, hybrids do not occur in the wild or are very rare. Here is an example of natural selection in progress: reproductive isolation separates the races even if they are living together in the same meadow. The genetics of the two races is slightly different and, eventually, they could become distinct enough to be classified as two species of Eurosta. S. altissima is the ancestral host species; it seems that flies first shifted to S. gigantea in the northeast corner of the United States and then spread north and west. The initial shift may have been due to a ‘mistake’ in oviposition: the female laid her eggs on the wrong plant growing in the same meadow and, because S. gigantea is closely related to S. altissima, the shift was likely to succeed.

Being able to shift to a new host species is advantageous to Eurosta for reasons that probably apply to other gall insects too–but there are few detailed investigations. Where both host plants grow together, E. solidaginis usually survives better on S. gigantea than on S. altissima due to reduced attack by enemies. Attack by the endoparasitoid Eurytoma obtusiventris and by downy woodpeckers (see Chapter 10 and Fig. 170) is reduced because, where there is a choice, they both prefer Eurosta in S. altissima. So, the gallfly is partially protected on the new host, although the escape is not absolute; both enemies can attack the tephritid in both host plants.

This leads to a question: if host shifts occur in the gall causer, does this drive a complementary host shift in the galler’s enemies? In the Eurosta–Solidago system the answer seems to be both yes and no, depending on the enemy considered. The ectoparasitoid Eurytoma gigantea (Fig. 170) has not followed the host plant shift of Eurosta solidaginis. Parasitoids from hosts on Solidago altissima or from S. gigantea seem to be identical in their behaviour: the adult females emerge at the same time, mate with any male, and oviposit in Eurosta galls in either host plant regardless of the species they emerged from. The beetle inquiline Mordellistena convicta (Fig. 170) is different, though it can live in both plants. The larva either feeds in a gall, usually eventually killing Eurosta, or it bores into non-galled goldenrod stems. The beetle has formed at least two host races: the ancestral race from ungalled stems and a derived race from galls, and it may be in the process of forming additional races depending on the particular Solidago species. Adult stem borers emerge from stems about ten days before the gall borers emerge on either host plant–although this may not isolate their mating because the adult beetles live for a long time, feeding from the flowers of other plants as well as goldenrod. The female beetles can mate with males from stems or galls of either plant species but they prefer mates from the same source as themselves. They also prefer to lay eggs into the host species in which they lived as larvae and mortality of their offspring is lower in these hosts too. Parasitism tends to be higher in stems than in galls, and higher in galls on S. gigantea than on S. altissima. Perhaps this differential mortality, and also differences in size of the adults (their size and therefore their fecundity varies considerably), will help to maintain the beetle’s shift from stems to galls and to favour beetles in galls on S. altissima rather than on S. gigantea–and result in several host races. So, in this example, the shift of the herbivorous gall-causing insect has stimulated a shift in a predator, in an insect in a higher trophic level and in one that is totally unrelated taxonomically. This phenomenon may be common–but proving it requires detailed long-term work that apparently has not been done for any other gall insect.

Hymenoptera: (1) Sawflies and the evolution of gall structure and position

Amongst Hymenoptera, the sawflies (Symphyta) are an ancient group of insects. Fossil adults are known from the Permian, 290 million years ago, and their origins are earlier. The earliest known fossil gall (see Fig. 221, Chapter 11) may have been caused by a sawfly. Most gall-causing sawflies belong in the Tenthredinidae (Table 4.6, Box 4.7, Chapter 4). This family is at least 200 million years old, and galling seems to have evolved independently within it eight to ten times. It was in existence long before the main radiation of flowering plants in the Cretaceous and, even today, a quarter of all sawflies feed on ferns and gymnosperms, while most of the rest are found on woody host plants in the more ancient families of angiosperms.

FIG 229. Cladogram for sawflies in the tribe Nematini causing galls on willows Salix spp., with type of gall and host plant indicated (based on DNA analysis; modified after Nyman et al., 2000; Roininen et al., in Raman et al., 2005).

Most gall-causing sawflies gall willows (Salicaceae) and are closely related within the tribe Nematini in the Tenthredinidae. There are four gall-causing genera on willows and the species have been subject to a recent cladistic analysis using DNA. There is a lot of uncertainty about the base of the cladogram (Fig. 229, bottom left), which indicates that Phyllocolpa is not monophyletic: some Phyllocolpa species appear to be more closely related to species in other genera than they are to other Phyllocolpa species. It is likely that galling arose only once within this tribe and that Phyllocolpa species show more of the ancestral characters than the other genera. An evolutionary series can be traced from Phyllocolpa through Pontania and Eupontania to Euura, with the galls becoming gradually more complex up the series (although none of these galls are as complex as those of gall wasps and many other groups). Gall-causing sawflies have diversified on the many species of willows and are unusual because their species richness increases northwards, in contrast to the poles-to-tropics trend of most insects. Most gall-causing sawfly species have a cold temperate, boreal or arctic distribution rather than warm temperate or tropical, probably due to their restriction to willows, which show the same distribution. This group of sawflies has not managed to diversify on to plants in other families.

Phyllocolpa species cause part of a leaf to thicken and the margin to roll or fold downwards causing a simple open gall, not very different from the ungalled leaf–and the larva sometimes feeds outside as well as inside the fold. The ovipositing female causes the leaf to fold. She moves down the leaf pricking it several times with her ovipositor and with each prick she injects a fluid before finally laying an egg (described in more detail in Chapter 4). Chemicals in the fluid cause the upper leaf cells to enlarge, thickening the leaf and causing it to fold downwards, and the gall will develop completely whether an egg is laid or not. Phyllocolpa galls also occur on Populus and on other Salicaceae in North America and Asia but, in contrast to the willows, galls on the other hosts have not evolved further and leaf-folding on these hosts seems to have been an evolutionary dead end.

Most willow galls are closed swellings, with adults and larvae sufficiently different from Phyllocolpa to belong to different genera. The first step in development of the Pontania galls is a small swelling around the egg, sometimes called a procecidium. The more primitive Pontania species are still leaf folders but they also cause a procecidium that encloses the egg inside the fold. When the egg hatches, the larva feeds on the procecidium and also on the rest of the thickened leaf. The sausage-shaped galls of Pontania dolichura (Fig. 229) are thought to represent the earliest completely closed galls, with P. proxima-type bean galls (described in Chapter 9) evolving a little later. These galls develop within the leaf blade and bulge more on the upper surface; their structure is simple, with an inner green layer reminiscent of a similar layer in the procecidium of the Pontania leaf folders. P. dolichura also retains the Phyllocolpa habit of injecting the leaf several times with fluid before laying the egg. The most advanced leaf galls form a single clade (i.e. a monophyletic group; Fig. 229) named Eupontania (now a full genus recently elevated from a subgenus of Pontania). These galls are all closely associated with the midrib, some species causing ovoid galls in the blade near the base of the leaf and the more advanced forming pea galls that hang down beneath the leaf; in these, only a scar is apparent on the upper surface, as in Eupontania pedunculi. The female Eupontania inserts her ovipositor through the midrib before piercing the blade alongside, and injects just one dose of fluid as she lays the egg (see Chapter 9). The gall has a slightly more complex structure than the Pontania galls and the hatchling must feed inside for a while before the gall develops to its full size. Gall size varies considerably in Eupontania and, in small undersized galls, the egg or young larva has usually died.

Euura causes the most advanced willow sawfly galls, all in a distinct monophyletic clade (Fig. 229). The galls are relatively simple like Eupontania, to which they are most closely related, but develop in stems, petioles and buds rather than in leaves. Euura galls retain the characteristic of Pontania and Phyllocolpa in growing to full size before the larva hatches. The basal Euura gall seems to be a swelling in the young stem (E. atra–type in Fig. 229). Later, galling moved up into the bud and petiole, also sometimes into the base of the midrib. Stem galls have reappeared in Euura amerinae, which represents the peak of sawfly gall evolution. This gall is often larger than other willow galls and usually contains several larvae in separate chambers.

In willow sawflies, therefore, galling has probably evolved only once from a Phyllocolpa-like ancestor and has proceeded gradually in a series of small steps to Pontania, Eupontania and Euura. The ovipositing female seems to be responsible for the changes and for the gradualism of the changes: sawfly galls are similar wherever they develop; their evolution is ‘constrained’. In no other group of insects does oviposition result in a complete and fully formed gall. The precise spot where the female injects the egg determines the type of gall that develops and ‘mistakes’ in location could be the main reason that the site of the galls has shifted–from leaf edge to leaf centre, base and midrib, and thence to stem, bud and petiole. The trend is from the edge to more central parts of the plant where the vascular tissues are more concentrated, and this results in a nutrient sink that becomes progressively stronger. There are other advantages of a more central position too. Stems and buds are less likely than leaves to be shed from the plant before the gall causer is fully developed–some plants selectively shed galled leaves early. And more central galls are more likely to receive the lion’s share of nutrients, reducing the quantity that reaches the leaf edges; competition for the basal position on leaves occurs in pemphigine aphids and in cynipid wasps (see Chapters 5 and 9) and probably occurs between sawflies too. Why should the female make mistakes when she is laying eggs? Eupontania and Euura oviposit earlier than Pontania and Phyllocolpa, into very young shoots. At this stage, the leaves, petioles and buds are tiny and only 1–2 mm apart at the tip of a shoot. Slight shifts in the exact position of the egg will make a large difference to the position of the gall when the shoot is fully elongated. After each shift in position, from leaf edge to midrib to stem and bud, a new radiation of species has occurred (Fig. 229).

Each new radiation of sawfly species is enhanced by shifts to new host willow species. Each gall type can occur on several willows (Fig. 229), primitive or advanced species. Although the phylogeny of Salix species is not well known, Fig. 230 suggests that the subgenus Salix sensu stricto (with S. fragilis, S. alba, S. triandra and S. pentandra) is more primitive than subgenus Chamaetia (includes S. glauca, S. reticulata and S. polaris) and that Vetrix (all the remaining species) includes the most advanced species. Comparing Figs 229 and 230 indicates that host-shifting between closely related willow species is no more likely than shifts between distantly related hosts. Choice of host by the female is based on only a few chemical compounds (phenolic glucosides), which are similar in most Salix species. So, from the female’s point of view, most willows are chemically suitable to receive her egg and shifts to new host species may be governed by opportunity rather than by chemistry of the host plant. Close neighbours of a galled plant, particularly if common and with abundant shoots growing vigorously when the females are flying, are more likely to become galled than are distant plants. Common and widespread willow species have more species of sawfly galls than rare and restricted species, shrubs with their abundant young shoots have more species than trees, and both trees and shrubs have more species than prostrate forms. So, host-shifting for sawflies has been relatively easy. Although host-shifting between willow species has occurred often, shifts to other plant genera are rare: most nematine sawflies (i.e. in the tribe Nematini) are restricted to Salix. They have rarely jumped genus barriers unlike other insects that gall willows, gall midges for example, perhaps because of restrictions caused by the method of oviposition and the chemistry of the fluid injected with the egg. Perhaps the ovipositor, which makes galling unique in sawflies, also constrains gall formation, restricting the galls mostly to willows.

FIG 230. A hypothetical phylogeny for willows Salix spp. (based on morphological characters; from Roininen et al., in Raman et al., 2005). The species are divided into three subgenera (Salix s.s., Chamaetia and Vetrix) as indicated on the left; the species with asterisks belong to Chamaetia.

Hymenoptera: (2) Gall wasps and fig wasps–the most complex of all galling conditions

The Euhymenoptera or Apocrita includes all Hymenoptera apart from the Symphyta and, in contrast to them, is characterised by parasitism: most species are parasitoids. A few groups are plant-or fungus-feeders and these have evolved from parasitic ancestors. Gall-causing occurs in three superfamilies: in the Cynipoidea (the gall wasps, all in the family Cynipidae), in the Chalcidoidea (six families of chalcid wasps, including the fig wasps Agaonidae) and in the Ichneumonoidea (only the braconid wasps, family Braconidae). In the Cynipidae and Agaonidae, gall-causing is common. In the other families it is sporadic and rare and has evolved several times in species that are not closely related.

The Cynipoidea is a relatively small parasitic superfamily, with about 3,000 living species. It includes five families existing today and two more known only as fossils. The two largest families, the Cynipidae and the Figitidae (each with about 1400 known species worldwide) are related sister-groups and together make up the ‘microcynipoids’ (there are three ‘macrocynipoid’ families, with only about 200 species between them and no gall causers). Unlike most cynipoid families, the gall wasps (Cynipidae) are plant feeders, gallers or inquilines, and species of one subfamily of Figitidae are inquilines too, though none are gall causers. The first Cynipoidea apparently evolved in the Jurassic (maybe 200 million years ago; Table 11.1), although no fossils as old as this have been found. Adults have been fossilised in Cretaceous and Tertiary ambers, the earliest about 90 million years ago, and these probably had parasitic larvae, parasitoids of wood-and cone-boring moth, wood wasp and beetle larvae, similar to macrocynipoids today. Most of the Figitidae are parasitic, many of them parasitoids of aphids and aphid predators and of larvae of ‘higher’ flies. The Cynipidae are largely gall-causing and are monophyletic, evolved from a parasitic ancestor. Galling by gall wasps seems to have evolved in the Tertiary; the oldest fossil gall wasp is about 45 million years old (Eocene), found in Baltic amber. The oldest fossil galls found are less old, from the late Tertiary to the late Pleistocene, and are considerably younger than their host plants–oaks, for example, probably evolved in the mid-Cretaceous and diversified through the Palaeocene and Eocene (65–35 million years ago). The immediate ancestor of the first gall-causing wasp may have been a parasitoid of other gall-causing insects; perhaps it then became inquilinous and added gall tissue to the parent gall and later became able to induce its own galls.

Gall wasps

The family Cynipidae contains two subfamilies: all gall wasps belong in the Cynipinae apart from one extinct species known as a fossil (subfamily Hodiernocynipinae). At present, the Cynipinae is divided into six tribes (Table 9.1, Box 9.1, Chapter 9). Fig. 231 is a cladogram for the tribes within the subfamily, with the Figitidae included as a related sister–group. The Cynipinae is a monophyletic group derived from a parasitoid ancestor that also gave rise to the Figitidae. This phylogeny suggests that the ancestral gall wasp belonged to the tribe Aylacini and gave rise to more aylacine-type species and the Synergini on the one hand and the ‘woody rosid gallers’ on the other (‘woody rosids’ because all of their hosts belong to trees and shrubs in the plant subclass Rosidae; the oak family Fagaceae belongs to this subclass). The Aylacini is clearly not a monophyletic group. The woody rosid group includes the well-known galls on oaks and roses, and all the inquilines are related in the Synergini. The Cynipini and Pediaspidini are assumed to be sister-groups because, in both, their lifecycles involve heterogony or cyclical parthenogenesis, alternation between a sexual and an agamic generation (Chapter 9 and Box 9.1).

FIG 231. A cladogram for the tribes of gall wasps (Cynipidae: Cynipinae) with host plants (in brackets), showing the position of the Figitidae, sister-family to the Cynipidae (based on morphological characters) (from Csóka et al., in Raman et al., 2005, slightly modified, from information in Ronquist & Liljeblad, 2001).

FIG 232. A cladogram showing the host plants of gall wasp genera in the tribes Aylacini and Synergini (based on morphological characters); genera of the woody rosid tribes are not included (modified after Stone, 1999; Csóka et al., in Raman et al., 2005; from information in Ronquist & Liljeblad, 2001). Line symbols indicate host plant family.

Fig. 232 is a more detailed analysis of aylacine genera. It suggests that the ancestral gall wasp galled a herbaceous plant in the poppy family (Papaveraceae; or perhaps a mint, Lamiaceae). It probably caused simple swellings with one chamber in developing seeds and adjacent tissues inside the poppy capsules, as Barbotinia and Aylax do today. This ancestor then gave rise to two major lineages, one that led to the inquilines and aylacine genera on Rosaceae (but not on roses), on Lamiaceae and on some Asteraceae, and the other that produced the aylacines on Papaveraceae and other Asteraceae and the woody rosid gallers. Fig. 232 suggests there have been at least three independent shifts from herbaceous to woody host plants: (1) by Diastrophus on to brambles (Rubus, in Rosaceae); (2) by the ancestor of the woody rosid group; and (3) by the ancestor of the inquilines. Once on a woody host, gall wasps have not shifted back to herbaceous plants; this character seems not to be reversible. Although in the gall wasps as a whole evolution to new host plants has been by host-shifting, host shifts have been rare compared to those in other herbivorous insects; gall wasps are remarkably conservative, faithful to particular host plants. Rather paradoxically though, when hosts shifts have occurred, they have involved unrelated or distantly related plants often already exploited by other gall wasps. In the Synergini, this had to happen because the inquilines are dependent on galls already on oaks, roses, sycamores and acacias. In aylacines, several herb gallers have converged on to related hosts in the same plant family: the Aulacidea–Isocolus branch and the Phanacis–Timaspis complex (Fig. 232) are not closely related and both have converged on to the Asteraceae independently, often using the same host genera. Some plants seem to be more ‘gallable’ than others, perhaps because gall wasps are limited by the chemistry of their hosts. Interestingly, when galling the same host plant, the galls of the two groups are similar. Although host-shifting, rather than coevolution (see above), is the rule in gall wasps, it is possible that species within some genera (i.e. at the tips of the ‘twigs’ in the family tree, Fig. 232) have diversified in parallel (perhaps coevolved) with species in a particular plant genus. An example might be the Isocolus gall wasps that have diversified on knapweeds, species of Centaurea–but coevolution for these is not proven; this may be an example of host-shifting that has occurred at an unusually rapid rate.

The phylogenies in Figs 231 and 232 are based on detailed morphological analysis. More recent molecular DNA evidence is altering the interpretation of cynipid evolution, however, although it is not yet clear how much previous conclusions need to be modified. But two important reinterpretations are probable:

The woody rosid gallers are probably not a monophyletic group. Apparently, the Diplolepidini–Pediaspidini–Eschatocerini (Fig. 231) form a monophyletic clade that is basal to all the other Cynipinae, suggesting that the ancestral cynipid galled a woody host rather than a poppy-type herb. The Cynipini, also monophyletic, form a separate clade within the Cynipinae. An important inference follows, that alternation of generations (heterogony) evolved twice, independently: once in the Pediaspidini and once in the much larger tribe, the Cynipini.

Saphronecrus–Rhoophilus and Periclistus–Synophromorpha (Fig. 232) are groups of related genera distinct from Ceroptres, and all of these inquilines are in one clade that also includes the gallers Diastrophus and Xestophanes. Synergus, Saphronecrus and Ceroptres are inquilines in oak galls, Rhoophilus is an oddball from a South African moth gall, and Periclistus and Synophromorpha live in rose and bramble galls, respectively. Diastrophus and Xestophanes gall hosts in the Rosaceae too. Inquilinism, it seems, has evolved more than once.

The position of the synergids has changed over the years. In the 1980s, it was suggested that the group was polyphyletic, each inquiline perhaps derived from its host gall wasp and therefore related more closely to it than to other inquilines. It was suggested too that inquilines were primitive forms that had not yet evolved the ability to cause galls on their own. The detailed morphological evidence that became available in the 1990s and the first years of the twenty-first century modified these ideas, and new molecular evidence is changing interpretations once again. Recent DNA evidence does not contradict two conclusions based on morphology, however. The inquilines are derived from gall causers that have lost the ability to initiate galls, although they can add gall tissue to an already-existing gall. And the original ancestral gall is still likely to have been a simple single-chambered swelling in reproductive tissues, though probably in a tree seed or fruit rather than in a herbaceous plant.

The Cynipidae is thought to have evolved in Asia and spread westwards into the Western Palaearctic and eastwards, reaching the Nearctic (Central and North America) from both directions. Rose and oak galls have a Holarctic distribution with known species richness greatest in the Nearctic, and their tribes (the Diplolepidini and Cynipini) may first have appeared there. The oak gall wasps have tracked their host plants, mainly Quercus species, which are also more diverse in the Nearctic than in the Western Palaearctic. But very little is known of the oaks and their gall wasps in the Eastern Palaearctic, especially in China. The diversity of oaks is high in the Eastern Palaearctic and new evidence (Abe et al., in Raman & Gupta 2007; Stone et al., 2009) suggests that both oaks and oak gall wasps may have originated in the Eastern Palaearctic before moving east into the Nearctic and west into the Western Palaearctic. The scenario probably is more complex, with several lineages moving eastwards and westwards between the Nearctic and the Eastern Palaearctic (across the Bering land bridge, which has been dry land for long periods) and, independently, between the Eastern and Western Palaearctic, and the Nearctic and Western Palaearctic. In the Nearctic and the Western Palaearctic, centres of speciation of oak gall wasps are Mexico and southeast Europe, Turkey and Iran, with three or four times as many oak gall wasps and oak species in the Nearctic as in the Palaearctic. The rose gall wasps (Diplolepidini) and their host roses are also more species-rich in the Nearctic (they may be highly diverse in the Eastern Palaearctic too). The sycamore gall wasp group (the Pediaspidini) is very small and is restricted to the Western Palaearctic. The acacia gall wasps (the Eschatocerini), too, is a small group, known only from Central and South America with species on Acacia (Mimosaceae) and Prosopis (Fabaceae). Acacias are common and widespread particularly in Africa and Australia, but the gall wasps are unknown there; they attack acacias in only a small part of their range.

Radiation of species within the large genera in the oak and rose gall wasp tribes, particularly in Andricus, has been stimulated by two types of event: by ancestral species shifting from one host to another, which is rare in gall wasps, and by much more common shifts between host organs, between buds, flowers, fruits, leaves, stems and roots. In the Western Palaearctic, white oaks (e.g. Q. robur and Q. petraea) and black oaks (e.g. Q. cerris) probably derive from ancestors in eastern Asia that spread westwards along the Himalayan foothills. Later, since the Pliocene (which started about 5 million years ago), oaks have diversified in Italy, the Balkans and Turkey. An ancestral Andricus species in the same area, with both sexual and agamic generations on white oaks, shifted successfully just once on to a black oak and then diversified into a large group of heteroecious species. These retained their agamic generation on the ancestral white oak with the sexual gall on the black oak. They include the well-known A. kollari, A. lignicolus, A. quercuscalicis and their relatives. Although host-switching is rare, when it occurred in Andricus it led to a rich diversification of species (Fig. 240, Chapter 11). Not so in Callirhytis, the only other genus of oak gall wasps with heteroecious species and not closely related to Andricus; Callirhytis has sexual galls on white oaks and agamic galls on black oaks and has remained a small genus–at least in Europe; it may be more speciose in the Nearctic. The occasional host switch has occurred in North America too, from an ancestral species on white oaks to a rich variety of derived species on red oaks (section Lobatae) but in none of these has heteroecy evolved (this conclusion may be premature as the lifecycles of many Nearctic oak cynipids are not known in detail).

Oak gall wasps that require Turkey oak Q. cerris for part or all of their lifecycle have been restricted, until comparatively recently, to southeast Europe and Asia Minor where the tree is native. Since the seventeenth century, and since 1735 in Britain, Turkey oak has been introduced into gardens and parklands in northern and western Europe and has enabled several gall wasps to increase their range. The first to arrive in Britain was Andricus kollari, the gall imported for its commercial use (dyeing, ink-making, etc.; see Chapter 13), and A. lignicolus, A. corruptrix, A. quercuscalicis, A. aries, A. lucidus, A. grossulariae and A. gemmeus have followed since. All of these have sexual galls on Turkey oak (except for A. aries, known only by its agamic gall at present). Although host-shifting involving heteroecy has provided opportunities for speciation and dispersal, it also may have increased rates of extinction in some areas because of the necessity for oaks of two sections of Quercus to be present. Species of two other genera have recently been found in Britain too: Aphelonyx cerricola with its asexual gall on Turkey oak (its sexual gall is unknown) and Neuroterus saliens with both generations on Turkey oak. Other species whose range now includes Britain following introduction of their host plant to Cornwall are three species of Plagiotrochus on holm oak Q. ilex. They, and P. amenti (= P. suberi) on cork oak Q. suber in California and Argentina, were probably introduced accidentally with their hosts (both black oaks). The related gall wasp Dryocosmus kuriphilus, a native of China and Korea, is another introduction and is now a pest of chestnut Castanea in Japan and the USA and elsewhere.

TABLE 11.4. Characteristics of six Diplolepis galls on two species of Rosa in Ontario, Canada (from information in Shorthouse et al., 2005)

Compared to shifts between host species, shifts between different organs or locations on the same host plant have been relatively common in gall wasps and have resulted in episodes of rapid speciation. Relocating within a host plant involves changes in the site of oviposition by the parent female and evidently is ‘easier’ for gall wasps than adapting to the new genetics and chemistry that a new host plant involves. This contrasts with gall-causing sawflies on willows, which have jumped to new willow hosts relatively frequently–but perhaps the chemistry of Salix species is more similar than in different species of oak. Diplolepis on roses and Andricus on oaks provide examples of species shifts between host organs.

FIG 233. Diagrammatic section through a rose leaf bud indicating the position of the eggs of five Diplolepis gall wasps: A. D. polita, B. D. bicolor, C. D. nodulosa, D. D. triforma, E. D. spinosa. The section shows the shoot tip with young leaves arising around the apical meristem (densely stippled) and becoming progressively older and larger behind the meristem; vascular strands (dashed lines), pith (lightly stippled). (Drawn from a photomicrograph in Shorthouse et al., 2005). (See text for further explanation.)

Diplolepis has diversified into many species on wild roses, particularly in North America where Rosa is species-rich. Host shifts have occurred occasionally but shifts between host organs have been much more common: between buds, leaflets and stems. In Ontario, Canada, two Diplolepis species gall Rosa acicularis and four more gall R. blanda (Table 11.4). D. polita and D. triforma gall R. acicularis, while D. bicolor, D. nodulosa, D. spinosa and D. fusiformis gall R. blanda. D. polita and D. bicolor are leaf gallers and the other four gall the stems, although in most the gall is initiated in a bud. As in all gall wasps, the eggs are placed very precisely during oviposition (Fig. 233) and it seems that slight differences in timing and location–the eggs of different species are placed only a few mm apart–has led to the different host organs being galled. Five species oviposit into buds and the sixth, D. fusiformis, lays eggs in rows along a young stem, each egg inserted into one epidermal cell. Perhaps these differences arose through slight errors in positioning of the eggs or to delays in the timing of egg-laying. The rare event of oviposition on to another host species, which could lead to more speciation, may also be due to an egg-laying error.

FIG 234. Sections of galls induced by a selection of European Andricus species in their sexual or agamic generations (from Stone & Cook, 1998; also reproduced in Stone, 1999; with permission from Graham Stone). In each figure, the central unshaded space is the larval chamber, or several spaces in multichambered galls, and cut gall tissue is shaded black; all are drawn life size, except for f, l, o and r, which are twice life size. (a) A. coriarius (agamic), (b) A. mayri (agamic; now known to be the agamic generation of A. grossulariae), (c) A. seckendorffi (agamic), (d) A. lucidus (agamic), (e) A. inflator (sexual), (f) A. grossulariae (sexual), (g) A. coronatus (agamic), (h) A. quercuscalicis (agamic), (i) A. viscosus (agamic), (j) A. hungaricus (agamic), (k) A. quercustozae (agamic), (l) A. gallaeurniformis (agamic), (m) A. hartigi (agamic), (n) A. conficus (agamic), (o) A. gemmeus (agamic), (p) A. polycerus (agamic), (q) A. kollari (agamic), (r) A. curvator (agamic), and (s) A. foecundatrix (agamic).

Andricus species gall the roots, trunk, shoots, leaves, buds, catkins or acorns of oak trees. Most species (perhaps all) are heterogonous, with alternating sexual and agamic generations, and galls of each generation are generally restricted to just one organ. The causes of the shifts in organs galled are unknown; errors in oviposition may be involved but, because some organs are sited far apart and develop at different seasons, there must be other causes too. A few species gall more than one organ in one generation, though this is unusual; one example is the agamic hedgehog gall of A. lucidus, commoner on buds but also found on acorns. Outside Andricus, the sexual currant gall of Neuroterus quercusbaccarum is common in catkins or young leaves and, in Pediaspis aceris on sycamore and maples, the sexual generation may gall both leaves and flowers.

Gall wasps and especially the oak gall wasp tribe induce a greater variety of gall structures than any other group, greater than the variety produced by the whole gall midge family (Cecidomyiidae), for example. Why is there such variety and what are galls of different shapes and sizes for? The large genus Andricus illustrates the many types of gall and is used to suggest some answers.

Fig. 234 shows sections across a variety of European Andricus galls. Photographs of selected galls are shown in Figs 235–8 and more are found in Chapter 9: Fig. 152b–A. foecundatrix, Fig. 156–A. kollari, Fig. 160–A. quercuscalicis and Fig. 163–A. lucidus. Some galls are multichambered with a spiny surface (Fig. 234a–d). In others, the single inner chamber is surrounded by an air space (e–k). Some are hard, thick-walled and woody with a single larva in a central chamber (p, q), while in others, the larval chamber is surrounded by modified bracts (r, s). A. gallaeurniformis (l) causes simple galls along the underside of the leaf midrib. Three species have galls unlike any other: the inner chamber of the A. hartigi gall (m) is surrounded by spines with broadened ends that join to enclose an air space; the A. conficus gall (n) is fleshy and soft; and A. gemmeus (o) is tiny with its surface covered with fleshy red tubercles. Andricus galls have other traits too: hairy surfaces, sometimes like cottonwool; surfaces covered with sticky resin or with sugary nectar; or the gall is spongy, incorporating many small air spaces; and some galls have two chambers, one housing the larva and the other empty.

FIG 235. The agamic gall of Andricus coriarius. (Graham Stone)

FIG 236. The agamic gall of Andricus gallaeurniformis. (Graham Stone)

FIG 237. The agamic gall of Andricus quercustozae. (Michael Chinery)

Gall shape and size, different for each species and generation of oak wasp, is determined by the gall wasp even though the gall is made of plant tissue–the gall is the extended phenotype of the insect (see above, Chapter 11). One reason for the variety seems likely, but is difficult to prove: that the galls protect the larva inside. An individual inside a gall with spines, sticky resin, thick walls or with many chambers, etc., is more likely to survive and so leave more descendants than one in a gall without these characteristics. The main enemies of oak gall wasps when inside the gall are chalcid parasitoids, birds and rodents (see Chapter 10) and some characteristics may be more effective against one set of enemies than another. Fine spines and hairiness, for example, may be effective against parasitoids but stout spines and coarse projections are more likely to deter small mammals and birds, especially as these are rarely restricted to one food source and can find easier meals elsewhere. The only parasitoids that can reach larvae in large, thick-walled, woody or spongy galls enveloped in hairs or cottonwool are those with very long ovipositors. In some species of Torymus and Megastigmus, selection has favoured ever-longer ovipositors (see Torymus affinis Fig. 218, Chapter 10). Some parasitoids have solved the problem by attacking the gall when it is small and immature so that a short ovipositor is adequate. But at this size, the gall wasp larva is tiny, not large enough to support most chalcids; so these species are koinobionts, their larvae delaying their development until the host is large enough to provide an adequate meal (see Box 10.1, Chapter 10). Sticky surfaces, e.g. on young knopper galls (agamic galls of A. quercuscalicis, Fig. 160, Chapter 9), can gum up the delicate valves of the ovipositor as the female drills into the gall. After laying an egg, she carefully cleans the ovipositor with her legs and antennae, a procedure that takes time that cannot be used for egg-laying. While cleaning, too, the female is vulnerable to her own enemies, like predatory ants, and sometimes she gets stuck fast by the glue. So, the result is fewer parasitoids and a degree of protection for the gall wasp. An inner air space is a common characteristic of oak wasp galls and, again, probably protects the larva inside the inner gall. A drilling parasitoid may miss the inner gall or not get enough purchase to penetrate it if it is loose and rolls about or swings suspended from a thread. Sometimes parasitoid eggs are laid in the air space, or in a decoy chamber (e.g. in A. galeatus galls; Cynips disticha galls have the same feature)–the larva may hatch but be doomed to starve, unable to reach the gall wasp larva. The most sophisticated protective devices involve ants and aphids, e.g. in A. infectorius (previously A. gallaetinctoriae; Fig. 239) and A. polycerus (Fig. 238), both southern European species. Aphids, attracted to the sap in vascular tissues that supply the larval chamber, congregate on the gall surface sometimes covering it completely. The aphids produce honeydew, which attracts ants, and the ants protect the aphids by driving away their enemies and, incidentally, any female parasitoids attempting to drill into the gall to reach the gall wasp larva. Some gall wasp galls cut out the middleman (the aphids) and actively secrete nectar on to the gall surface (Dryocosmus cerriphilus in S. Europe is an example)–ants collect the nectar and protect the gall wasp larva inside from drilling parasitoids. Oak trees do not usually produce nectar, so the gall wasp and its extended phenotype must have tapped into a developmental pathway that is not normally expressed by the oak.

FIG 238. The agamic gall of Andricus polycerus. (Gyuri Csóka)

FIG 239. A cluster of three (green) agamic galls of Andricus infectorius (= A. gallaetinctoriae; one is young, smaller with distinct tubercles), with two (brown) cola nuts of Andricus lignicolus. (Gyuri Csóka)

Despite the variety of gall shape and structure, mortality due to enemies is often high and some gall traits do not seem to increase survival significantly. Perhaps the trait was effective in times past when the gall first evolved but no longer protects against modern parasitoids: this has been called ‘the ghost of parasitism past’. Even if no longer useful, the trait would not be lost unless it became a liability, if the larvae inside were less likely to survive than those in galls without the trait. Because it is the host plant that produces the galls, they are not costly to the gall causer and so there would be no pressure for a trait to be lost during evolution unless it became detrimental to the galler’s survival. Some gall characteristics may be more effective against a harsh environment, like desiccation or fire, than against particular enemies. But, although galling is probably a general advantage for herbivores in some environments, compared to leaf mining or free-living grazing for example, the effectiveness of spines, hairs, tough walls and other structures against particular environmental stresses has not been demonstrated.

Most of the characters found in gall wasp galls on oaks are shared by several species. Are species with the same trait closely related, indicating that new species have evolved after the trait appeared? Or does a character appear in galls that are not so closely related, which would suggest that the character is adaptive and has evolved more than once to cope with similar stresses imposed by enemies or by the physical environment? The family tree (Fig. 240) suggests that both of these alternatives are true. The gall ancestral to all of these species was probably small and simple, with a single chamber, and without surface adornments of spines, resin, etc. and is likely to have been the sexual gall, which usually is the simpler gall. Closely related species often cause similar galls. Bud scales enclosing the inner gall of the agamic generation are found in A. foecundatrix, A. curvator and others in the A. foecundatrix clade. The galls of the three species in the A. lucidus clade are spiny, covered with sticky resin and have many chambers. Typical of the A. quercuscalicis clade is a sticky surface and an internal air space; and the galls of the A. kollari and A. coriarius clades are hard and woody with thick walls. These traits evolved before the clade diversified into several species (conserved evolution). Some characters have appeared independently several times in more than one clade, which suggests that less closely related gall wasp larvae have solved some problems of survival in similar ways (convergent evolution). In the species in Fig. 240, spiny or hairy galls have evolved from smooth galls three times (2), multiple chambers have appeared twice, derived from single-chambered galls (3), and sticky surfaces have evolved from non-sticky galls twice also (4). The A. hartigi clade is unusual in that all of its galls are different despite their close relationship (Fig. 234, above); this demonstrates that new gall structures can evolve relatively rapidly. North American Andricus galls show similar mixtures of conserved and convergent evolution, too.

FIG 240. A cladogram for European Andricus species illustrating the evolution of several traits (based on DNA analysis; modified from Stone & Schönrogge, 2003; Stone et al., 2008): 1, heteroecy (species above the dashed horizontal line gall both white and black oaks; those below it gall white oaks only); 2, spininess; 3, many chambers; and 4, sticky surface resin. The arrow indicates that all of these species evolved from a common ancestor. Figures (photographs) of selected galls are indicated. (The genus Andricus is the subject of active on-going research and species are regularly added to the cladogram; as a result, details are regularly modified and updated and may alter in the future.)

FIG 241. Agamic gall of Andricus caputmedusae. (Graham Stone)

Andricus galls illustrate three general evolutionary patterns, and all may have a defensive function:

Conservation, in that most galls in a clade have the same character(s).

Convergence, with galls that are not closely related developing similar structures so that they come to look similar.

A general increase in gall complexity over time.

Parasitoids, and lethal inquilines, cause more deaths than most other mortality factors and these three patterns all could be responses to parasitism. Gall causers can escape from parasitoids by shifting to new host plants–at least for a time, until the parasitoids catch up with them. But this has not happened often in oak gall wasps. The more common solution, the evolution of traits that that are protective, encourages the parasitoids to ‘fight back’, and this could result in a coevolutionary arms race (or ‘Red Queen’ scenario, see Chapter 10) between parasitoids and gall causers. Coevolution may sometimes involve one parasitoid and one gall trait, such as the trend for ever-longer ovipositors associated with larger and thicker-walled galls (mentioned above). But, in oak gall wasps (and rose gall wasps too), a whole community of insect enemies attacks and competes for a variety of galls on the tree at the same time; most parasitoids and lethal inquilines are not specific to particular species of gall. So, the success of a trait in one gall cannot be judged against one particular parasitoid; interactions between the galls and their enemies are far more complex and have to take into account the whole gall and enemy community in a place over a particular time. Overall, though, protection against enemies is the most likely explanation for the rich variety of shapes and structures of oak cynipid galls.

The fig wasps

The final detailed example concerns the fig wasps (Agaonidae). This is the only family within the Chalcidoidea to be largely plant-eating. The Chalcidoidea is a huge superfamily with 20,000 species known and many more to be discovered; the overwhelming majority are parasitoids. Although the agaonids have many rather primitive or ancestral characters, the original chalcidoid is thought to have been a parasitoid. By inference from DNA phylogenies, the association between fig wasps and figs is old, going back 87 million years, but the earliest fossils come from less than 60 million years ago. Figs and fig wasps are tropical and subtropical in distribution and are found all over the world, while most other gall-causing chalcids are found only in Central and South America and in Australia. The symbiotic association of the figs and their pollinating fig wasps is unique–in no other gall insect is there any evident benefit to the plant host. (There are a few other cases of mutual benefit to the gall causer and to the host: the bacteria Rhizobium and Frankia in root galls of legumes and alder trees (see Chapter 3, Solid Swellings caused by Bacteria); also the association of mycorrhizal fungi and plant roots, which causes gall-like swellings on the roots.)

The fig–fig wasp relationship represents the best evidence we have of coevolution between insect gall causers and their plant hosts. The pollinating fig wasps (subfamily Agaoninae) is the only major insect galling group in which coevolution has occurred–at least, all available evidence supports this conclusion. But coevolution is difficult to prove. Not only must the phylogenies of the insect and host plant groups parallel each other, but the branching of each cladogram must be synchronous, occur at the same time. Otherwise, parallel cladograms could be due to rapid host-shifting by the insects colonising new host species almost as soon as they appear. Coevolution involves figs and their pollinators and is driven by the dependence of each on the other: the only insects capable of pollinating fig flowers are fig wasps and the fig wasps can only live in the florets of specific figs, galling them in the process. Not all fig wasps are pollinators. Some gall the florets without pollinating them and some have become parasitoids of other fig wasps (see Table 7.2, Box 7.2, Chapter 7), and these have not coevolved. Fig. 242 shows the relationship between the pollinating fig wasps in the Agaoninae, the other agaonid subfamilies, and other chalcid families that are closely related. The Agaonidae is clearly not monophyletic; some of its subfamilies are more closely related to the Eurytomidae, Torymidae or Pteromalidae than they are to the Agaoninae. Classification at this level needs modification and some subfamilies will eventually be removed from the Agaonidae. The Agaoninae, which includes all of the fig pollinators, is monophyletic, however, and is basal to the other subfamilies and to other chalcid families. The fig–fig wasp mutualism arose only once, too–perhaps about 87 million years ago.

FIG 242. Cladogram for the subfamilies of the Agaonidae showing their relationships with other chalcid families (in bold type; based on DNA analysis; after Weiblen, 2002).

FIG 243. Evolution of two characters in the mutualism between Ficus species and their pollinating fig wasps (15 genera are included with one, Ceratosolen, split into several species). (a) Breeding system of the host fig: monoecious, dioecious or polymorphic (species of figs in the section Sycocarpus are either monoecious or dioecious). (b) Pollination behaviour of the fig wasp: active, passive or polymorphic (four genera contain active or passive species); Ceratosolen galili has lost its pollination behaviour (see text). (Based on DNA analysis; from Machado et al., 2001, slightly modified.)

The mutualism of fig wasps with their figs has resulted in several characteristics that illustrate the coevolution of the two groups. Two of these characteristics involve the breeding system of the host and the pollination behaviour of the fig wasps: the figs are monoecious or dioecious and the wasps are passive or active pollinators (described in Chapter 7). About half of all Ficus species are monoecious: each fig inflorescence produces and disperses both fertile seeds (the female function) and pollen (the male function). The remaining figs are dioecious: seed-producing inflorescences occur on different trees to those that produce pollen. Fig wasps that are active pollinators collect pollen into special pouches and transfer it purposefully on to receptive stigmas, while pollen dusts the bodies of passive pollinators and gets distributed passively as they crawl over the female flowers. The parallel evolution of these characters (Fig. 243a & b) coincides remarkably closely. Monoecious figs and passive pollinators are the original primitive conditions–found in species of Tetrapus, the basal genus that pollinates figs of the subgenus Pharmacosycea, the most primitive of existing figs. Tetrapus wasps are Neotropical and fossils of adults have been found in South America dated to the Oligocene, 34.5 million years ago. Figs of the subgenus Urostigma (Fig. 243a), found in the tropics around the world, retain the monoecious condition. Subsequently, dioecy arose twice, first in the ancestor of the Ficus–Sycidium–Sycomorus clade and later when the lineage branched into Sycidium and Sycomorus–Sycocarpus. Meanwhile, the ancestor of the Sycomorus–Sycomorus figs had reverted from dioecy back to monoecy. In Fig. 243b, active pollination arose just once in the ancestor of all non-Tetrapus pollinators, and these developed pollen pockets of various types in order to carry the pollen. Active pollination reverted to passive pollination independently at least five times: in Waterstoniella and in some species of Platyscapa, Pleistodontes, Blastophaga and Wiebesia; all of the species in these genera have reduced or vestigial pollen pockets. Ceratosolen galili is unique in this group having lost the ability to pollinate. It has become ‘parasitic’ on the fig florets, causing galls without pollinating the stigmas. The remaining genera and species in Fig. 243b are active pollinators. There are one or two instances where coevolution probably has not occurred. The fig subgenera Ficus and Sycidium are closely related sister-groups but their pollinators, Blastophaga and Wiebesia on Ficus and Kradibia and Liporrhopalum on Sycidium, are not, although they occur in the same clade (Fig. 243a). It seems that host-shifting has occurred occasionally even in a group that has otherwise coevolved. As in all phylogenies, the story will change as more molecular data become available (not least in sorting out the Ceratosolen species: this is a large genus and is clearly not monophyletic).

The stability of the mutualism between figs and fig wasps over its long evolutionary history is extraordinary. The interaction is in no way altruistic in either partner, though: each is in it for what it can get out of it. So, why should active pollination by the wasps evolve? It requires pollen pockets to develop and the appropriate complex behaviour to evolve, both of which take resources and time that could be used for other things (to produce and lay more eggs, for example). Pollination of the fig floret allows the seed to grow and provides more and better-quality food for the fig wasp larva, and development of the gall in the ovary enhances this. Active pollination also results in more developing seeds in a fig inflorescence and more successful galls, and so higher numbers of gall wasps develop despite the extra resources required from the wasp. And each fig produces more fertile seeds too, despite having to pump extra resources into the galled florets. So, pollination, and particularly active pollination, enhances survival of the fig wasps as well as the figs, despite the costs involved on both sides, and helps to explain the stability of this long-term mutualism. But the balance between costs and benefits can change, as is illustrated by reversions to passive pollination in some fig wasps and the loss of pollination altogether in others (in Ceratosolen galili, at least). Thus, this intimate fig–fig wasp interaction is not an evolutionary dead end; it can lead to other things.

Some fig species attempt to reduce the costs of providing galls. In these, the inflorescence partly fills with fluid after the florets have been pollinated, a fluid that creates breathing problems for the wasp larvae–it seems as if the fig attempts to get rid of the fig wasp larvae after the adults have done their job of pollination. But the fig wasps ‘fight back’. The spiracles (breathing holes) of larvae in fluid-filled figs are enlarged, with hairy rims that repel water, an adaptation to the semi-aquatic environment inside the fig. Several species of the large genus Ceratosolen are examples, adapted to the fluid-filled monoecious figs of the subgenus Sycomorus. So, the fig wasp survives, benefiting the fig species in the long run as wasps are available to pollinate future generations of figs. The coevolution of the fig–fig wasp partnership proceeds by adaptations to reduce costs on both sides but with knock-on effects that benefit the other side. And so the mutualism is reinforced and stable over the long history (87 million years) of the interaction.

Evidence for the synchronous timing of the fig–fig wasp coevolution comes from plate tectonics and movement of continents (see Chapter 11). In the early Cretaceous, 140 million years ago, the southern landmasses of South America, Africa, India, Antarctica and Australia were fused together as one supercontinent known as Gondwana. About 100 million years ago, Gondwana began to break up, and Africa separated from Antarctica and South America and started to move northwards. The ancestral species of both the figs and the pollinators appeared about 87 million years ago and spread throughout Gondwana, without Africa, from South America across Antarctica (which was not frozen then) to Australia. The ancestral species became the Pharmacosycea figs and the Tetrapus fig wasps (Fig. 243). India separated from Gondwana about 80 million years ago, an event that coincided with the appearance of most of the fig subgenera and fig wasp genera in Fig. 243; the ancestors of these groups spread through much of the Old World and colonised Africa via Madagascar, which remained attached to India for a while. South America, Antarctica and Australia were linked until the Palaeocene or Eocene (Table 11.1, Chapter 11). Australia separated and the fig wasp genus Pleistodontes diversified there about 64 million years ago, and South America finally split from Antarctica in the Eocene, coinciding with diversification of the neotropical Pegoscapus wasps. The remaining pollinators of the Urostigma figs radiated in the Eocene and Oligocene, and dispersed throughout the palaeotropics.

An afterthought in evolution: large insect groups with a few simple galls

Gall-causing in two large unrelated orders of insects, the moths and butterflies (Lepidoptera) and the beetles (Coleoptera), and in two large groups of Hymenoptera, the Chalcidoidea (apart from the Agaonidae) and Ichneumonoidea, is unusual and sporadic. Most of the galls are simple accumulations of callus tissue, not organised into distinct tissue layers. Moth and beetle galls are similar. The caterpillar or beetle larva feeds on the gall callus, in plant stems or roots or, in some beetle galls, in leaves and seed pods. Most larvae of related species are free-living and feed externally on leaves. Some are stem borers and may cause gall-like swellings of wound tissue if several larvae happen to be living together. These are not classed as true galls unless the larvae feed on the extra tissue–but often it is difficult to draw the line between wound-swellings and simple true galls, particularly if the habits of the larvae are not known in detail. Wound-swellings may represent an early stage in gall evolution or, perhaps, indicate that galling in a species has been lost–but there is no evidence for this, either way.

Moth and beetle galls have evolved independently several times in unrelated families. In the Lepidoptera, these are neither the most primitive nor the most advanced, while in Coleoptera, the more advanced weevils include most of the gallers (in the large families Curculionidae, Apionidae and Nanophyidae). Moth families are grouped into 46 superfamilies. Neither the nine most primitive superfamilies nor the eleven most recent (the Macrolepidoptera or butterflies and some larger moths) include any gallers. Galling in butterflies may not be possible because of their size. Their caterpillars are larger than gall causers in any other group, and too large to feed comfortably inside stems and leaves of many plants. Galling in all moth groups appeared late, in the most highly derived species in a family; galling in moths, and in beetles too, is not a primitive trait. An unusual moth family is the Cecidosidae (in the superfamily Incurvarioidea). It only has seven species but all cause galls and these are slightly more elaborate than most moth galls–they are hard rounded or tubular galls with an aperture plugged with a stopper or operculum. Cecidosidae occur in Africa and South America and gall species of Schinus and Rhus (Anarcardiaceae). The family probably evolved on the supercontinent of Gondwana in the early Cretaceous about 100 million years ago, before Africa and South America separated from Antarctica.

The only gall causers known in the Ichneumonoidea are a few recently discovered braconids. The first plant feeders in this group of parasitic wasps were discovered in the 1980s and the first gall causer was found ten years later–a few more galls have been discovered since but very little is known about them. There are 39–48 families of braconids and three include a few gallers; these are not closely related and their galls have evolved independently. Allorhogas (subfamily Doryctinae) is a large genus of parasitoids found worldwide, and the few gall causers known from the genus are from Brazil and Costa Rica. The galls are hidden inside fruits (of Fabaceae, Melastomataceae and probably other plant families) and are easily missed, so probably more gallers will be found in the future. Some Allorhogas species are parasitoids or inquilines in gall midge and gall wasp galls but their relationships with the host galls are poorly known. The Mesostoinae is a small subfamily found in Australia where it is probably endemic, and its galling species have specialised on Banksia (Proteaceae), an endemic plant genus. Monitoriella belongs to a third subfamily whose affiliations are uncertain. It is a neotropical genus found from Mexico to Peru and in Trinidad, and several species cause leaf galls on Philodendron species (Araceae).

Chalcid wasps are also mainly parasitic and include very few gall causers, apart from the fig wasps. A few Torymidae, Eulophidae and Eurytomidae cause galls. Probably most of them have evolved from seed or fruit feeders, or from stem borers–the Tetramesa species (Eurytomidae) arose from stem borers in grasses. Paragaleopsomyia cecidobroter (Eulophidae; described in Chapter 10, Ambrosia gall midge galls) is an unusual inquiline and causes ‘endogalls’ associated with the ambrosia galls of Asphondylia atriplicis; it cannot cause galls from scratch. Perhaps it is partway along the evolutionary route to true gall formation, or perhaps it has lost the ability to cause galls having evolved from gall-causing ancestors (in the same way as synergid inquilines in the gall wasps).

EVOLUTION OF GALLS: A SUMMARY AND SOME CONCLUSIONS

Although galling as a lifestyle has a long history (more than 300 million years), most evolutionary studies of gall insects, and of gallers in other groups, are relatively recent and many questions remain to be answered. Galling is a minority lifestyle for insects and has evolved independently in many unrelated groups. Other herbivorous strategies, such as leaf mining, stem boring and free-grazing, are more common: galling seems not to be cost-effective for these. Galls are made of plant tissue but the insect controls their structure and position on the plant, and the gall cannot evolve independently of either the insect or its host plant. Despite the intimate association, coevolution of insect and plant is rare: it seems that the plant usually leads the way with the insect following (host-shifting or shifting to new parts of the plant), often some time after the appearance of the plant species. The best-known example of coevolution occurs in figs and fig wasps where, unusually, both partners benefit from and depend on the association. Highly specialised lifestyles with dependence on one or a few closely related species could lead to an evolutionary dead end–but this has not happened in most groups of gall causers. They have colonised new sites on old hosts and on new host species, and often a burst of speciation has followed each shift. Galling clearly ‘works’ for these species. And galling can be lost and new lifestyles appear, such as parasitism, inquilinism and free-living herbivory. Examples occur in most of the galling groups included in this chapter (and also in gall-causing scale insects; information on their evolution may be found in Cook & Gullan, 2004, and Gullan et al., in Raman et al., 2005).

Apart from the gall causer, other organisms often benefit from the gall, and they have evolved along a variety of routes. Some inquilines and parasitoids are closely related to and have evolved from the gallers; others are unrelated. There seems little doubt that the rich variety of shapes and sizes of galls, particularly in gall wasps, is related to heavy attack by these enemies. In galls containing many related individuals, as in aphids (and in thrips and some scale insects, not included in this chapter; information on their evolution can be found, for thrips in Crespi et al., 1997, and Mound & Morris, in Raman et al., 2005; and for scales, in Gullan et al., in Raman et al., 2005, and Takagi, in Raman & Gupta, 2007), some siblings have evolved into soldiers that defend the gall, attacking and driving away potential parasitoids and predators. Here, social organisation reminiscent of other insects that live in communal nests has developed: as in honeybees, bumblebees, vespid wasps, ants and termites. Gall mites are also small enough for many individuals to inhabit one gall–has social behaviour evolved in any of these? None has been reported.

Other groups of gall insects are unusual for one reason or another. The gall midge family, the Cecidomyiidae, is large and widespread and, unique for a family, galling is the commonest lifestyle. The ambrosia gall midges are unusual too with three interdependent species, plant, gall midge and fungus, and the gall midge and fungus partners cannot survive without each other or their host plant. Evolution of the gall midges is not well understood, though; many species are not yet described and relationships between tribes and subfamilies is not clear. Conversely, sawfly galls on willows are caused by a tribe of closely related species whose evolution is relatively well understood. These are unusual for two reasons. They have an unusual distribution, commoner in the far north and at higher altitudes. And the galls of many species develop without the stimulus of a feeding larva, a characteristic that is not found in other groups.

Tracing phylogenies is a problem because the fossil record of galls and gallers is so fragmentary; cladistics provides useful additional information. But cladograms are only as reliable as the information they include, which may be sparse and based on too few species. The patterns of evolution they suggest are based on current knowledge that is usually incomplete; as more information is added, interpretations may alter and cladograms will become more reliable. The more information used the better, both from DNA and morphological analyses. Knowledge of gall-causing species, their way of life, their structure and their DNA, is increasing all the time and, as a result, cladistics are clarifying how gall insects have evolved.

CHAPTER 12

Galls and Geography

THIS CHAPTER LOOKS AT TWO aspects of the distribution and biogeography of gall insects: global patterns and gradients in species richness and their possible causes, and cases of range expansion of galling insects and their parasitoids and inquilines. The second aspect gives us a set of ‘natural experiments’ that help us to understand the first aspect. We concentrate on insects because they are better known than other gall-causing groups (mites, nematodes, fungi, bacteria).

Gall insects are found in every continent, apart from Antarctica. In many areas the gall fauna is poorly known–the galls of Europe and of North America north of Mexico and their distribution are the best studied (see, for example, the review articles in Raman et al., 2005). More species will certainly be found in all regions but it is especially the tropics, the southern hemisphere and much of the Eastern Palaearctic that are seriously under-studied (Raman & Gupta, 2007, bring together current knowledge of the biogeography of gall insects in the Eastern Palaearctic and Oriental regions). So, as knowledge accumulates, distributions apparent at present will need to be modified. Particular families of gall-causing insects are characteristic of different geographical regions probably due mainly to the distributions of their host plants. Most hosts are flowering plants, especially dicotyledons, but some groups attack monocots and also conifers and ferns. In most respects, the distribution of gall insects and the patterns of variation in their species richness worldwide follow the pattern seen in many groups of animals and plants.

Climate and geological history divide the terrestrial world into biogeographical zones (Fig. 244). The basic divisions are between temperate and tropical parts of the world. In the northern hemisphere, the Nearctic (North America and Greenland) is distinct from the Palaearctic (Europe and central and northern Asia), although these two are often combined as the Holarctic. In the southern hemisphere, the temperate land mass is much smaller because the continents extend less far south. The Antarctic region includes the tip of South America, the islands of the southern oceans and New Zealand, and Antarctica. There are four main tropical/subtropical regions: the Neotropical (most of Central and South America), the African, the Oriental (India and southeast Asia) and Australian (Australia and associated islands to the north and east). The southern tip of Africa, the Cape, is much smaller than the rest but is distinguished botanically because of its unique flora.

FIG 244. Biogeographical regions of the world (compiled by Alexander Cameron).

GLOBAL GRADIENTS AND PATTERNS OF GALLING

Four gradients or patterns have been proposed to account for the variation of plants and animals worldwide–though the reasons for the patterns are not completely understood. They apply also to gall causers, although some may be more relevant to some galling groups than to others.

Latitude–Species richness increases from polar regions to the tropics, reflecting the general amelioration of climate. An altitude gradient has a similar effect, with species richness increasing from high to low altitudes down a mountainside.

Area–Species richness increases as the area of land increases. This reinforces the latitudinal gradient: the tropical zone is the largest because the tropical latitudes either side of the equator are adjacent.

Harshness of environment–Species richness increases from harsh extreme conditions to ‘easy’ equable conditions. An environment may be harsh and extreme in terms of climate (hot and dry, for example), or soils (e.g. thin, acidic and nutrient-poor), or be subject to regular catastrophes (e.g. fires, floods, droughts). At the easy equable end of the gradient, the climate is mild, warm or cool (not too hot and not too cold), there is plenty of water but not too much, and there are deep nutrient-rich soils.

Host plant species richness–Species richness of herbivores increases as plant species richness increases.

These gradients help to describe the distribution of gall causers, with one gradient perhaps more important than another in particular galler groups. The latitude gradient occurs in a wide range of organisms, from protozoa, ants and woodpeckers to apes and monkeys, as well as in plants and gall insects. Why species richness should increase from the poles to the tropics probably has several explanations (over 25 mechanisms have been suggested, ranging from effects of glaciations, variety of habitats and environmental stability to competition between species, and chance). The gradient is not understood and must be due to many interacting mechanisms. Despite the overall patterns (1, 2, 3 or 4 above), in some gallers the gradient is opposite to that expected, with species richness increasing towards the poles or increasing in harsher environments. Examples are included that illustrate a variety of patterns. All explanations are tentative–we need much more information on the distribution of gall species, particularly from the tropics, before distributions can be adequately described and explained.

TABLE 12.1. Gall-causing Cecidomyiidae in tropical Costa Rica and in four temperate countries (from Hanson & Gomez-Laurito, in Raman et al., 2005; for calculation of number per 1,000 km², see the original text).

It seems logical to expect that the variety of gall causers should increase as the species richness of host plants increases and also as land area increases (i.e. in the tropics). These patterns do seem to be fundamental, even though local climatic or soil conditions may distort them. Tropical rainforests probably have more plant species per unit area than most other habitats and, probably, gall species richness is high too, perhaps higher than anywhere else. Because plant species richness tends to increase from the poles to the tropics the general trend of gall species richness is in that direction too. The main problem is lack of knowledge of tropical gall faunas: many temperate regions in the northern hemisphere are much better known than any tropical area. Costa Rica perhaps is becoming an exception; the variety of its galls is better known than in most other tropical countries, although many of its galls are unnamed and described only to family rather than to species. Costa Rica extends from 8° to 11° North in Central America and is relatively small, only the size of West Virginia or Denmark. Its gall fauna is rich: 967 species of galling arthropods (insects and mites) have been found on 711 species of plants in 131 families, all vascular plants, i.e. ferns, conifers and flowering plants but not mosses and liverworts (Hanson & Gomez-Laurito, in Raman et al. 2005), and no doubt this will turn out to be a considerable underestimate. Most (80 per cent) of Costa Rica’s gall causers are gall midges (Cecidomyiidae), with only 6–7 per cent or less each for the gall mites, gall wasps, psylloids and other groups. Compare figures for Japan, seven times as big (Table 12.1) and more thoroughly surveyed: about 1400 species of galling insects and mites are known of which only about 45 per cent are due to gall midges; 20 per cent are caused by Hemiptera (mostly psylloids) and about 16 per cent by gall wasps. In Costa Rica, the pattern of galling is generated by gall midges and seems to be due to their wide distribution across host plants–although they are mainly host plant-specific, gall midges attack most families of plants, unlike gall wasps, psylloids and scales that specialise on particular genera of hosts. Table 12.1 compares numbers of galling cecidomyiids in Costa Rica with four countries with temperate climates. There are more than twice as many gall midge galls in Costa Rica (290 per 1,000 km²) as in the temperate countries, probably because there are more than twice as many plant species in the tropics: one estimate in Costa Rica is of 3368 species per 1,000 km² compared with 1527 per 1,000 km² in North America. Gall midge galls follow the plant species richness, with a greater variety of both in tropical than in temperate regions. This temperate–tropical pattern seems to hold true even when complicated by unusual aggregation of particular gall causers on particular plants and in particular soil conditions (see below). Gall midges dominate the gall fauna worldwide, though they seem to be relatively more common in the tropics than in higher latitudes. But gall midges and their galls are not as well known as some other gallers, in temperate as well as in the tropical areas, and this is unfortunate in view of their commonness. We need more studies in all parts of the world before any patterns of distribution can be confirmed.

Cynipid wasps and gall-causing sawflies are uncommon in tropical gall faunas mainly because their host plants are typical of temperate and polar regions. Oaks (Quercus) are uncommon in Costa Rica except in the very wet cloud forests at high altitudes, which have a more temperate climate. Here, oak gall wasps are common at least on one abundant species, Quercus copeyensis, which supports 23 gall wasp species. Sawfly galls are unknown in Costa Rica although their non-galling relatives are not rare: there are over 150 species in Costa Rica. No doubt the lack of galling sawflies is due to the virtual absence of willows (Salix); only one genus of nematine sawflies, the group that specialises on willows, is known in the whole of the Neotropics.

Other areas of the world besides the wet tropics are rich in plant species. The Cape floral region at the southern tip of Africa stands out as one of the richest. This region is renowned for its high levels of endemism: many of its plant species have evolved there and are found as natives nowhere else. The Cape has a warm temperate, Mediterranean-type climate, with hot summers and mild winters and with seasonal rainfall. The soils are thin, dry and nutrient-poor and frequent fires are characteristic. The typical vegetation is a shrubby heathland known as ‘fynbos’, with a rich variety of woody plants less than 1 m tall. The heath merges with open woodland at higher altitudes where rainfall is higher and with semi-desert where rainfall is less. Species richness of low woody shrubs is higher in the fynbos than in other types of vegetation nearby and gall insects are also more diverse, with more than twice as many species in the fynbos as elsewhere (Wright & Samways, 1998), many of them endemic. On average, only one galling insect attacks each fynbos shrub species and most galls are caused by gall midges, as in the tropical fauna of Costa Rica. Psylloid, cynipid, tephritid and apionid (weevil) galls are all rare. As in the wet tropics, gall insects of the fynbos, especially the gall midges, track plant species richness.

Fig. 245 illustrates a poles-to-tropics trend of gall causers linked to increasing plant richness but with gall species richness peaking in the warm temperate zones (latitudes of about 23°–45° North and South) and then reducing towards the equator. The pattern reflects local diversity of gall causers: there are more gall species and each is more abundant in an area of a few square metres in these warm temperate climates than in the tropical zone. There are almost certainly more gall species in the tropics as a whole (i.e. regional diversity) but they are sparser, their density (abundance) being lower at the local level. The zones either side of the tropics have Mediterranean-type climates with hot summers and mild winters, and with a long dry season, with rainfall concentrated either in winter or in summer. The zones extend further north and south than the true Mediterranean climatic zones: at relatively high altitudes in the tropics where rainfall is seasonal rather than year-round, and in warm deserts at higher altitudes where rainfall is more abundant than is characteristic of deserts. This pattern is not unique for gall causers; it occurs in parasitic wasps and other insect groups in North America and in Europe south through Africa. All sites in Fig. 245 with a high number of gall species (more than 12 per local sample) have ‘scleromorphic’ vegetation. This is a dry heathland type of vegetation, rich in species of low woody shrubs with small, tough, leathery, long-lived leaves that often are resinous and aromatic. These plants tend to be indigestible and toxic to herbivores, because of their high fibre content and high concentrations of tannins and other phenolics. Scleromorph vegetations have been given different names in the different parts of the world that have a Mediterranean-type climate: maquis in Israel, Greece and elsewhere around the Mediterranean, fynbos in the southern tip of Africa, kwongan in southwest Australia, and chaparral in South America and in California and Arizona in North America. Outside the true Mediterranean zones, similar scleromorphic vegetations are the cerrado and campiña of Brazil and the Sonoran Desert vegetation of Arizona. The soils of all of these areas are thin, dry and nutrient-poor, and sometimes contain high levels of heavy metals toxic to many plants. This vegetation is fire-prone too, adapted to regular fires. So, these environments are harsh, with a long dry season, frequent fires and impoverished soils. The gradient of gall species richness increases with harshness, running in the opposite direction to that expected.

FIG 245. Species richness of gall insects on a latitudinal gradient north or south of the equator, with tropical, warm temperate, cool temperate and arctic zones indicated. All data are from local samples, i.e. all galls found in a standard small area were collected. Key to symbols: mesic vegetation (closed circles); scleromorphic vegetation (open circles); scleromorphic vegetation in relatively mesic sites (indicated by x). Squares indicate samples from fynbos vegetation: scleromorphic vegetation (open squares); riparian woodland in mesic sites (closed squares). All samples with a species richness of more than 12 are from scleromorphic vegetation (indicated by the vertical dashed line). (Modified from Price et al., 1998; with permission from Peter Price and the Journal of Biogeography, Blackwell Publishing.)

It is insect and mite galls that are species-rich in Mediterranean-type vegetations. Fungal galls are not well known but are probably rare in such dry environments. Why are arthropod galls so common? Scleromorphic plants are not good sources of food for externally feeding herbivores, like caterpillars and leaf beetles, or for leaf miners: the toughness and toxicity of the leaves provide good protection against these groups. But gall insects can bypass these deterrents and manipulate the plant in order to acquire adequate supplies of food. Inside galls, the larvae are enclosed in or sit upon a layer of tissue that, as a nutrient sink, concentrates nutrients and water drawn from elsewhere in the plant; so the gall larvae do not go short. The long-lived leaves of these host plants also benefit the larvae. The tough tissues and toxins tend to be concentrated in the outer layers of the galls, where they deter grazing caterpillars and other free-feeding herbivores and pathogenic fungi, and so benefit the gall larvae. Parasitoids, inquilines and predators might do well too, able to avoid the plant’s toughness and toxicity and protected by the plant’s defences in the same way as the gall causers. Overall, it seems that mortality of gallers in scleromorphic vegetation is generally lower than in nearby more mesic sites (which are less extreme: less hot and dry, with more fertile soils), but specific causes of death are not well known in these vegetations.

Gall insects seem to be more species-rich on sclerophyllous vegetation in wet tropical rainforests too (sclerophyllous leaves are relatively tough and leathery, but not to the same extent as in true scleromorphic vegetations). The tops of tall trees that form the high canopy of these lofty forests suffer greater water and nutrient stress than do the lower parts of the trees and trees in the understorey. Water and nutrients come from the soil via the roots and have a long way to travel to get to leaves at the top of a tall tree, and are likely to be in short supply at high levels. Leaves in the high canopy are adapted to withstand drought even in a rainforest: they are sclerophyllous, tougher and with more toxins than those on the same tree lower down. In a survey in a rainforest in Panama, Central America (Ribeiro & Basset, 2007), the diversity of galling insects increased with leaf sclerophylly and with climatic harshness, reflecting the same pattern as in the fynbos vegetations of Mediterranean climates. Species richness of gallers increased with height, with high numbers in the high canopy. Survival of the galler larvae increased with height too, and in the more sclerophyllous leaves. Inside their galls, insects suffered less parasitism and predation, and the galls were less likely to be attacked by fungal pathogens, than those in softer leaves at lower levels. In the same study, free-feeding herbivorous insects were compared with galling insects at increasing height in the canopy. The species richness of the free-feeders reduced with height and decreased as the toughness of the leaves increased while the gallers did the opposite. The high canopy of tropical rainforests has been seriously under-sampled for insects in general, and for gall insects in particular. If the pattern shown by the Panama forest is general, it will considerably increase estimates of the species richness of galling insects in the tropics.

In the Mediterranean heathlands, gall midges are the commonest gall insects (this is probably true in the topical rainforests too but families of gallers were not recorded in the Panama study). Even though the adults are delicate and short-lived, they are good dispersers and readily find isolated patches of vegetation, so they can colonise rapidly after fire. Their high species richness is reinforced by individual plant species that have become centres of speciation for gall midges, particular host plants that are particularly ‘gallable’, supporting unusually high numbers of closely related species. The creosote bush Larrea tridentata in Arizona supports galls of 16 Asphondylia species, Atriplex canescens in southern California has eight Asphondylia species, and Acacia tortilis in a similar vegetation in Kenya supports 18 gall-causing midges. Once a gall midge becomes adapted to a particular host plant in these dry habitats, particularly a host with a patchy, scattered, distribution, it may diversify rapidly into new species, each specialising on different parts of the same plant. Scleromorphic vegetations are already species-rich in plants and in cecidomyiid galls and, if one genus of gall midges specialises and speciates on a particular plant species, this will add to the species richness. So, the high species richness in the fynbos, for example, could be due partly to high speciation rates on a few plant species rather than to the gallers being attracted to the rich variety of plant species alone. Plants that are highly attractive to gall midges are known outside Mediterranean-type vegetations too; for example, two Fagus (beech) species in Japan support 21 species of Hartigiola gall midges. Some cecidomyiid genera have specialised on particular host genera: Rabdophaga, for example, has 80 species known worldwide, all causing galls on the species-rich genus Salix. But in this case, each willow species supports only one or a few species of Rabdophaga so that numbers of gall species follow the increase of plant species richness. This gradient is likely to be clearest where gallers are spread evenly across plant species, typified by the one galler per plant species that is characteristic of most cecidomyiid gallers. In the wet tropics, where plant species richness is also high with no one species dominating the flora, the same pattern should be apparent–but we need more data before this can be concluded.

Although many galling groups increase in species richness from the poles to Mediterranean latitudes and the tropics, sawflies on willows are an exception. There are more species of both in colder northern latitudes than further south, although they also decline in the far north, north of the Arctic Circle. The variety of both sets of species also increases with altitude, e.g. in the Alps, Himalayas, Japan, southern China and the Rockies. No sawfly galls are known in Australia even though one family, the Pergidae, is indigenous there and has richly diversified on eucalypts. The north–south pattern has been illustrated by a study in Japan, a chain of islands running roughly north–south between latitudes 45º and 30º North (Roininen et al., in Osaki et al., 2006). A 1250 km distance was surveyed from Hokkaido in the north to Honshu in the south and, at several points, transects were followed down mountainsides, covering a range of 1900 m in altitude. Sawflies of the four nematine genera, Phyllocolpa, Pontania, Eupontania and Euura, were found and both willow and sawfly species richness declined towards the south: 13 willows and 21 sawflies occurred in Hokkaido and only five and two, respectively, in the most southerly Honshu site. The abundance of galls on individual plants also declined southwards. The same pattern was apparent over the range of altitude–species richness and density were both higher at higher altitudes than in the delta regions of rivers. The sawflies declined more rapidly than their host willows, probably because patches of willows were smaller and more fragmented further south and at lower altitudes. There are probably several reasons for the sawfly decline. Dispersal by adults from one isolated wood to the next is less likely than in continuous Salix woodland. Also, as willow stands become less abundant and less uniform, the sawflies become less well synchronised with tree phenology. For successful gall formation, the female must lay her eggs when leaf buds and shoots are at the appropriate stage of growth, usually as the shoots are elongating rapidly in spring. In northern climates, leaf flush is well synchronised between trees and occurs rapidly, over a relatively short period. As long as the females are flying at this time, there will be no shortage of egg-laying sites; females will be able to lay all of their eggs and the resulting gall density will be high. Further south and at lower altitudes, the timing of leaf flush is less predictable; it occurs over a longer period but individual trees are more variable, some becoming green earlier than others. So, if most females are flying at the same time, there may be a shortage of leaf buds and shoots at the correct stage of growth, which will result in higher mortality and fewer galls. Also, in warmer climates, full-grown sawfly larvae spend longer dormant in their cocoons (Peter Price, pers. com., 2007) and this can lead to higher mortality than in colder climates, deaths due to higher metabolic costs as well as to the greater chance of predation. In Japan, continual invasion from the mainland to the north via Sakhalin and the Kuril Islands probably reinforces the abundance of sawflies in Hokkaido. As sawflies disperse southwards, their movement may be prevented or slowed down by stretches of sea and the need to ‘island-hop’. Some sawflies are less able to cross water than others: Salix caprea and S. triandra are without sawfly galls in Japan even though they are galled in Sakhalin and on the mainland nearby.

Apart from these broad geographical patterns, there are also variations in the numbers and kinds of galling insect species depending on the structure of the plant. Generally, the number of herbivores increases as complexity of structure increases, with fewest on herbaceous plants, more on shrub-sized plants and most on trees. Gall insects only partially follow this trend, however. Gall species richness is higher on woody than non-woody host species, but shrubs tend to be richer in species than trees, and younger and smaller individuals of tree species often bear more galls than older and taller specimens. Scrub woodland and woodland edge are often richer in species than nearby mature and ancient woodland. Most cynipid wasps on oaks and sawfly gallers on willows illustrate this pattern. The main reason is probably the greater abundance of fast-growing buds and shoots on younger plants and on shrubs rather than on trees; gall causers require the continual production of young, active plant tissues. Willows with a shrubby growth form produce many shoots sprouting from low down and from ground level, while trees ‘concentrate’ on gaining height as rapidly as possible, their terminal buds and leading shoots taking the lion’s share of the plant’s resources. This results in trees having fewer buds and young leaves to gall than shrubs. Natural pruning of a woody plant to remove its main leading shoot has a rejuvenating effect, resulting in an abundance of young, vigorous side shoots and a bushy growth form, and this provides an abundance of sites for gallers. Regular fires may have a similar effect, stimulating a rich flush of new growth over the next few years and helping to explain why fire-prone vegetations have a high species richness of galls. But some gall causers do not follow this trend. The American aphid Pemphigus betae (described in Chapter 5) attacks large cottonwood trees Populus angustifolia rather than shrub-sized individuals although, on a particular tree, the fundatrix selects young, rapidly growing leaves on which to lay her eggs. P. betae is probably restricted to mature trees by the conditions necessary for its overwintering egg to survive: the egg needs to be protected in a deep fissure in the bark and, in young trees, the bark is too smooth. A few Eupontania sawfly species, too, are more common on tree-sized willows. These species lay their eggs in the commonest leaf buds regardless of whether they are on more or less actively growing shoots and, because short, less vigorous shoots are usually more abundant than long vigorous shoots, the galls are more common on the short ones. These sawflies emerge very early in spring when willow buds are just starting to grow and the differences between more and less vigorous shoots are not yet apparent (even to a female sawfly). So, the females select the commonest shoots rather than the most vigorous, unlike sawfly species that emerge later in the spring.

Gall-causing eriophyoid mites, despite being very common, are less well known than gall insects. Like insects, most gall mites are specific to a single host plant and their species richness tends to increase as plant species richness increases, particularly as the number of species in a genus increases. In the boreal forests of Finland (Niemalä et al., in Osaki et al., 2006), two species-rich plant families, the Salicaceae and the Betulaceae, are hosts to 42 per cent of the country’s gall mite fauna. In addition, the abundance of the host plant, its geographical range and the average size of its leaves affect gall abundance: widely distributed and abundant plant species with relatively large leaves have more gall mite species, whether or not the plant is shrub-or tree-sized. Birches and willows are abundant and dominate these northern forests and would be highly apparent in the landscape. Commoner trees are more likely to be colonised regularly by gall mites than rarer species, especially as gall mites tend to be passively distributed by the wind.

CHANGING RANGES OF GALLS AND GALL INSECTS

Over long time scales, i.e. over geological and evolutionary time, the distribution of gall causers has changed (as is true for all organisms). Some of this change is connected to movement of continents and ancient changes of climate, and therefore to change in the distribution of host plants (see also Chapter 11). Distribution is also related to more recent climatic changes triggered by the Pleistocene glaciations: there have been at least nine glacial periods between 1 million and 18,000 years ago. In particular, the general warming in climate that has occurred since the last glacial period ended, 12,000–10,000 years ago (in Britain), has affected distribution. This warming has allowed plants to spread naturally out of the refuges in southern Europe to which they were restricted during the last glaciation and to extend their ranges northwards, and the gall insects to follow them. Glaciations have affected North America too and also the southern hemisphere, though less severely there because existing landmasses do not extend so far towards the pole as in the north. At least part of the latitudinal trend (described above) may be due to delays in the host plants, and even more their gallers, ‘catching up’ with these climatic changes. In addition, gall insects have extended their ranges because of human activity, over time scales of only a few hundreds of years. Host plants have been planted outside their native ranges and some gall causers have followed them, under their own steam or aided by humans, and their enemies (parasitoids and inquilines) have followed the gallers. And some gall insects have been accidentally or deliberately introduced into a new country over a time scale of only decades–perhaps becoming pests or being introduced in order to control a pest (this topic is expanded in Chapter 13). In this section, the expanding ranges of a tephritid and several cynipids and their parasitoids and inquilines are described. Their patterns of spread may apply to other gallers too, though few are yet known in sufficient detail.

Tephritid flies–Tephritidae

The tephritid fly Urophora cardui causes a stem gall in creeping thistle Cirsium arvense (described in Chapter 8), and its distribution in Europe is known in more detail than that for other tephritids. Although its host plant is widespread and common across Europe, U. cardui is patchy, with large gaps in its distribution particularly in central Europe. U. cardui has expanded its range naturally since the end of the last glaciation following expansion of creeping thistle, and its range continues to expand at the present time. This may be because its postglacial expansion is still under way or perhaps it is a response to human-induced changes to habitats or to global warming. Over a small scale of distance up to 10 km, the distribution of U. cardui varies dramatically, both from place to place and over time. Local populations frequently disappear: a patch of creeping thistle with good numbers of galls in one year may have none the next. But it seems that local extinctions are usually more than balanced by colonisations of empty patches. Both are variable, though, and in some years extinctions are commoner than new colonisations. It seems that regular colonisation–extinction events are characteristic of U. cardui populations.

U. cardui adults, like many tephritids, seem not to be very active: they sit around on thistle plants and are easy to capture in a pill box or specimen tube, and have been thought to fly only short distances. But studies by Eber & Brandl (1997) have shown that they are highly mobile and are able to move several kilometres a year, so that there has been more than enough time since the last glaciation for the species to have spread throughout the range of creeping thistle. The tephritid fly has probably repeatedly colonised northern areas and repeatedly disappeared–perhaps due to suboptimal environments or to local interaction with parasitoids. The centre of distribution for C. arvense and U. cardui during the Pleistocene glaciations was in the large river valleys of the western Mediterranean and Greece, glacial refuges for both species. During the present interglacial, creeping thistle has spread north along two main routes west and east of the Alps following the rivers Rhone–Rhine and the Danube–Oder–Elbe. U. cardui has followed behind but remains patchy. It does not occur throughout the whole of the creeping thistle’s range: it is absent from parts of southwest France and Italy (perhaps due to the very dry summers there), and also from Brittany, Switzerland, Norway and Sweden. It extends considerably further north in mainland Europe than in Britain: it reached southern Finland in 1981 but has not yet been recorded in Scotland or in England north of south Yorkshire (where it was recorded for the first time in 2006).

The patchiness of U. cardui seems to be due to its colonisation–extinction dynamics. The adults are mobile enough to find host plants in favourable environments and to migrate outwards from a parent population, establishing new daughter populations as a series of ‘stepping stones’ (see Box 12.1). In a site, numbers of galls characteristically will fluctuate from year to year, perhaps dying out for a while and restarting as new adults fly in. Local extinction may be due to the climate becoming warmer or drier or to loss of habitat (e.g. due to control of the thistles, which are regarded as weeds), or it could be caused by parasitism. The chalcid Eurytoma robusta (see Chapter 10 and Fig. 171) can decimate a population: it is an ectoparasitoid that also feeds on the gall tissue and can break through from one chamber to the next destroying most or all of the host larvae in a gall. E. robusta follows U. cardui; as it catches up with some of the U. cardui populations, it survives for a while or declines and disappears, perhaps having caused its host to die out, before moving on to new patches of U. cardui. Genetic variation of U. cardui populations declines from south to north: over 400 km from northern Germany through Denmark, for example (Johannesen & Seitz, 2003). This is expected if relatively small proportions of southern populations have moved north (see Box 12.1). The most northerly and most recent populations in southern Finland are less variable than those in France, and perhaps a similar decline occurs from south to north in Britain too. But genetic variation of E. robusta does not follow the same pattern and there is no south–north decline over the same geographical area. E. robusta is not specific to U. cardui, however; it also parasitises tephritids in flower head galls of thistles and knapweeds (see Chapter 10). It seems that E. robusta is opportunistic, specialising on U. cardui from time to time as it catches up with it but attacking other hosts too.

Oak gall wasps–Cynipidae

Cynipid wasps and their host plants, like Urophora cardui and Cirsium arvense, were confined to glacial refuges during the Pleistocene glaciations and have spread northwards in the interglacial periods. In the present interglacial, most gall wasps have followed the expanding range of their host plants and many now occur naturally wherever the host occurs. In Europe, where the oak cynipid fauna is best known, the glacial refuges in the south have the richest variety of species, and species richness decreases northwards. There are three distinct refuges for oak gall wasps: Iberia (Spain, Portugal and the Mediterranean coast of northwest Africa), Italy, and the Balkans and Turkey to Iran. These refuges were isolated during glacial periods, by ice in the mountains and by treeless plains too cold for the survival of continuous forest. During each glacial period, each refuge was isolated from the others for long periods (probably for more than 100,000 years), long enough for genetic variation specific to a refuge to evolve. Subsequent migration northwards, therefore, can be traced back to a particular refuge: northern populations may have come from only one refuge or be a mixture derived from more than one. Some species expanded their range rapidly after the ice of the last glaciation disappeared (12,000–10,000 years ago in Britain). Other species moved north more slowly, restricted by physical barriers (e.g. mountain ranges or seas) or slowed down by interactions with other species (competitors for too few galling sites, perhaps, or predators and parasitoids causing high mortality). And some species are still more or less restricted to their glacial refuge. It seems that all oak cynipids in the Western Palaearctic share a common phylogeographic history: the group speciated dramatically in Anatolia (eastern Turkey) at least 5 million years ago and subsequently species have migrated westwards into Europe several times, some reaching Iberia perhaps 1 million years ago. Their ranges have waxed and waned as ice sheets in the north retreated and advanced during the last 2.4 million years (see Table 11.1, Chapter 11) and more species evolved in each refuge when isolated during the glaciations.

The cynipids whose distribution is best understood are Andricus species in the Western Palaearctic with sexual galls on Turkey oak Quercus cerris, a black oak (section Cerris, see Chapter 9 and Box 9.2). Q. cerris is native to south and east Europe from Italy to Turkey, an area restricted by mountains to the north: the Alps, Tatras and Carpathians. Unlike Q. robur, Q. petraea and other white oaks (section Quercus s.s.), Q. cerris did not escape this refuge naturally after the last glaciation. But over the last 400–500 years, people have planted Turkey oak in parks and gardens in western and northern Europe, as far as eastern Ireland, northern Scotland, Denmark and Latvia. It now forms scattered ‘islands’ of single trees or small groups within woodlands dominated by white oaks, with Q. robur and Q. petraea the commonest species across most of Europe. The heteroecious (host-alternating) cynipids dependent on Q. cerris and on Q. robur and other white oaks (Chapter 9) are native to the same southeast European refuge as Q. cerris; some species have expanded their ranges north-and westwards following the plantings of Q. cerris and are now found wherever the two host oaks occur together. Some species have spread naturally while others have been aided by human introduction, intentionally or accidentally. Because the range expansion has occurred over only the last 400 years, the genetic history of populations remains distinct enough for invasion routes into northern Europe to be traced (see Box 12.1). After longer periods of time, newer and older populations that were genetically distinct become similar as adult gall wasps migrate between them. These Andricus species have not become pests in the invaded areas and so they have spread as they will, unhindered by attempts to control them–unlike a related cynipid Dryocosmus kuriphilus, a pest of sweet chestnut (Castanea species; discussed below and in Chapter 13). Q. cerris does not occur in Iberia; here its place is taken by the closely related cork oak Q. suber, also a black oak. A few heteroecious Andricus species have spread into Iberia from the east and use Q. suber for their sexual generation but most have not; the host switch from Q. cerris to Q. suber is difficult, it seems.

BOX 12.1. Tracing invasion routes: founders and bottlenecks

The pattern of spread of an invading species depends on its abilities to disperse from a parent population (a source population) and, for a herbivorous insect, the distribution and abundance of its host plant. Adult insects dispersing in different directions from the parent population are likely to give rise to, or ‘found’, a number of daughter populations that may be genetically slightly different from the parent population. So, over a geographical area that contains the native source population of a species and an invaded range, the population is likely to be divided into subpopulations that vary genetically. This variation can be used to trace patterns of invasion of a species whose range is expanding–perhaps along a single route progressing away from the source population or fanning outwards in several directions at once. Genetic variation of populations is assessed by comparing the presence and frequency of different forms (alleles) of particular genes. Original populations that have evolved in a place and existed there for a long time are likely to be more variable than invading daughter populations that are younger, particularly in the variability of their ‘allozymes’ and ‘microsatellites’. Allozymes are enzymes, derived from alleles, which exist in more than one form although all of the forms function in exactly the same way (i.e. they all do the same job of catalysing the same chemical reaction). Microsatellites are regions of a chromosome where the DNA is highly repetitive, and the number of repeat units in a microsatellite varies between individuals. The variation in allozymes and microsatellites does not affect the functioning or survival of the individuals, but it does allow the probable age of the population to be estimated and the ancestry of the invading species to be traced.

As a species spreads out of its native area and invades new areas, three factors will affect the genetic variation of the daughter populations:

1. The size of the source population and the numbers of founders. If there are few founders, their genes will be a small subset drawn from the larger parental gene pool, so that the genetic variation of the new populations will be less than in the old. The new colony will have passed through a genetic ‘bottleneck’. Rare alleles in the source population are likely to be missing from the invaders, and the number of allozymes and size of microsatellites will be smaller. Or if one of the founders happened to carry a rare allele, this could become more frequent in the daughter colony than it is in the parent population. On the other hand, if the daughter colony remains small, its rare alleles could be lost over time just by chance, resulting in a further decrease in variation. If the number of founders is large, the genetic difference between the parent and daughter populations will be less distinct.

2. The origin of the founders. Gall insects can only invade new areas if they can find new patches of their host plant. If the host forms small widely scattered patches, the founders are likely to come from the nearest source population, and the invasion will progress via a series of ‘stepping stones’, each giving rise to the next daughter colony. Each of the populations in the series will be founded by colonists carrying a subset of the alleles of the previous population, and so genetic variation is likely to decline along the line of the invasion. But if the new host plant patch is near other inhabited patches, founders can come from several source populations from several directions (i.e. from several neighbouring ‘islands’) and the new population will be more genetically variable.

3. The subsequent exchange of adults between the new colony and other populations. Exchange of migrants between new and older populations will reduce genetic differences between them even if the rate of immigration is low, and so new colonies will become more like the old. If the populations remain very different genetically, this suggests that gene flow between them is very low, perhaps because the dispersal ability of the adults is limited.

Ten oak gall wasps dependent on Q. cerris and native to its southeast European glacial refuge have invaded north and west into Britain, following the plantings of Turkey oak. Eight are Andricus species that require Q. cerris for their sexual generations. A. kollari was the first, aided by deliberate introduction. Then followed A. quercuscalicis, A. lignicolus, A. corruptrix, A. lucidus, A. aries, A. grossulariae and the most recent, in 2008, A. gemmeus. In addition are two species dependent solely on Q. cerris, Aphelonyx cerricola and Neuroterus saliens. A. infectorius has reached Germany but is not yet recorded in Britain. No doubt more species will arrive in the future. The rate of invasion across Britain varies between the species: some have moved rapidly (A. kollari, A. quercuscalicis, A. lignicolus and, since 1997, A. aries) and are now common, while others are expanding much more slowly and still have a relatively restricted distribution (A. corruptrix and the more recently arrived A. lucidus, A. grossulariae and Aphelonyx cerricola). The invasion history of A. queruscalicis and A. kollari is particularly well known and the details are described here.

Andricus quercuscalicis causes the familiar knopper galls on acorns of Q. robur and on hybrids between Q. robur and Q. petraea, but it is extremely rare on pure Q. petraea or other white oaks. Its host range is more restricted than most Andricus species. Knopper galls enclose the agamic generation. The sexual galls are tiny and develop on the catkins of Q. cerris (the lifecycle is described in Chapter 9). Both galls can be very common, sometimes the commonest Andricus species in a particular woodland–although like most insects, their numbers often vary dramatically from year to year. A. quercuscalicis started to extend its range naturally not long after Turkey oak began to be planted outside southeast Europe and it is now found from the Balkans and Turkey to Britain and Ireland (Sunnucks et al., in Williams, 1994). In 1631 it was recorded in southeast Germany, it was recorded from Berlin in 1787, the Netherlands in 1882, and reached England in 1958 and Ireland in 1986. Q. cerris was first planted in England in 1735 but A. quercuscalicis did not arrive for more than 200 years; it took time to move out of its native area, and perhaps its progress was slowed by the necessary sea crossing. It is not yet found north of the Scottish borders even though Q. cerris has been planted as far as the north coast of Scotland. It is probably limited by the scarcity of Q. robur; in north and west Britain Q. petraea is the commonest oak. It does not occur in Iberia because Q. cerris does not occur there–it has not made the switch to Q. suber unlike some Andricus species. At eastward limits of its range, in eastern Turkey, Ukraine and beyond, it is limited by the rarity of Q. robur, while northwards in mainland Europe absence or rarity of Q. cerris imposes limits. The agamic females are the main dispersal stage; they emerge from knopper galls on Q. robur and fly considerable distances to find a Turkey oak with catkins, relatively rare amid the Q. robur. The pattern of spread of A. quercuscalicis from its endemic range in southeast Europe seems to have followed a series of stepping stones, with new colonies appearing in patches of oak forest containing Q. cerris and each colony started by a few founder individuals from the previous population in the series (see Box 12.1). Its genetic variation follows such a cline and is highest in the endemic area, reducing northwards and westwards to the lowest in Ireland. Populations in eastern Hungary, closely followed by Slovenia and eastern Austria, are the most genetically diverse and contain a number of rare alleles that are missing further west. Variability has been progressively lost further north and west, with populations in southwest England and in Ireland the least variable. Britain may have been colonised more than once, in eastern England and again in the southwest. Alternatively, the first invaders may have arrived in the southeast and progressed westwards, losing variation on the way. It is likely, too, that there were very few original immigrants into Britain, probably fewer than 15 individuals.

By 1996, four alien heteroecious Andricus species, A. kollari, A. quercuscalicis, A. lignicolus and A. corruptrix, had invaded southern Britain and had spread varying distances northwards. The northward spread in Britain has been well documented and has provided a large-scale natural experiment on the impact of the new gall wasps on each other and on the native oak gall wasp community (Schönrogge et al., 2000; Schönrogge, 2001). In 1996, all four species occurred in southern England and had existed there for longer than further north. A. corruptrix was found south and east of a line from the Wash to the Dorset coast, A. quercuscalicis reached the Scottish borders, A. lignicolus was found as far north as the Great Glen in central Scotland, and A. kollari was the only species in the far north. A. quercuscalicis and, probably, A. lignicolus and A. corruptrix, spread gradually overland to Britain from southeast Europe, but A. kollari has a different history. The marble gall was deliberately introduced into Devon in the 1830s (for dyeing, tanning and ink-making, see Chapter 13). Huge numbers of marble galls were brought by boat to Britain from the Middle East and Turkey; Haleb (Aleppo) in Syria was the main centre for export. Most British and Irish individuals, and those in northern coastal Europe (Belgium, the Netherlands and northern Germany), are genetically distinct from their neighbours on the European mainland, which was gradually colonised overland from different sources in southeast Europe (Stone et al., 2007). Most A. kollari populations in Britain come from one or more very large introductions from the eastern end of A. kollari’s native range, from southeast Turkey and the highlands of Lebanon, Israel, Syria or Iraq. By contrast, one or two British populations have spread northwards overland, in the same way as described for A. quercuscalicis, from the Balkan glacial refuge and also, it seems, from Iberia. So, British A. kollari populations have a complicated history.

Marble galls were first seen in Britain in 1834 on oaks in the Exe valley in Devon (Walker et al., 2002) and were regularly imported after that. Since then A. kollari spread rapidly eastwards, perhaps boosted by more introductions elsewhere in southern England. In 1854 they were common in southeast England and spread northwards, reaching southern Scotland in 1871 and northern Scotland in 1882. Since 1980, A. kollari has been recorded on some of the islands of the Inner Hebrides. It is still absent from the Outer Hebrides and parts of northern Scotland, probably because of the absence or rarity of oak trees. The English Channel seems not to have been much of a barrier, allowing emigrants to spread back into Belgium, the Netherlands and northern Germany; these populations are more closely related to British than to southern A. kollari from the Balkans. For all of these exotic species, the agamic females that emerge from the marble galls are probably the main dispersers and so the sexual galls became established first. Since 1996, three of the species have further extended their ranges northwards, while A. kollari had probably reached the limit of its host trees before 1996, having been in Britain more than 100 years longer than the other species.

Invasion by new species is likely to result in changes in the local community. It might mean increased competition for sites to gall, i.e. for suitable buds on either or both of the host trees. Or there may be no shortage of buds but parasitoids, inquilines and predators of the local native galls might be affected: there may be changes in the community of insects associated with the galls of the local species as the enemies discover and include the new galls in their food range. The changes may be complex and indirect and, no doubt, will alter over time. Besides the number of invading species decreasing northwards, the diversity (abundance as well as variety of species) of the native oak cynipids and their enemies declines also: more than 20 oak wasp species were found in the south and only 15 in the north, and 17–18 parasitoids and inquilines occurred in the south, with fewer than 10 species in the north. This trend is superimposed on the usual year-to-year fluctuations of a species, which can be large. Where they are present, the four invaders are often the commonest species at a site, especially the sexual generation galls on Q. cerris. Despite this, and despite the general decline in cynipid species richness and abundance from south to north, the invader species in Britain have not apparently affected the native gall wasp fauna (or had not, by 1996). This suggests that there is no shortage of space and no competition between ovipositing females; invading gallers can always find suitable buds to gall.

In northern Scotland, Andricus kollari has reached the northern limits of Quercus robur, Q. petraea and Q. cerris. At this extreme limit of its range, populations of A. kollari behave differently from those further south: the lifecycle of most individuals takes two years to complete rather than one year, no doubt due to the colder climate. This results in regular alternation in its density, from high in one year to low in the next (see also Chapter 9). This fluctuation has knock-on effects on A. kollari’s parasitoids, particularly when A. kollari is the commonest oak gall at a site. The parasitoids continue to have annual lifecycles, as they do further south. In high-density years in northern Scotland there are plenty of hosts available but not enough parasitoids to exploit them because relatively few emerged from the low host density in the previous year. And those emerging from a year when the host was common will face a shortage in the following low-density year. So the proportion of A. kollari parasitised is low in high-density years and high in low-density years, not what is expected from ‘well-adjusted’ parasitoids.

Most parasitoids and inquilines that attack oak cynipids are not specific to particular galls: communities inside galls of similar size and position on the tree, and present at the same time, tend to include the same species (see Chapter 10 and compare the food web diagrams), although their abundance often varies. So, a new gall invading a community of native galls in an oak woodland, particularly if the gall is common, will provide alternative hosts for native parasitoids and inquilines as long as they recognise it as a source of hosts. After A. quercuscalicis arrived in Britain, the knopper galls were almost free of parasitoids for about 20 years. It seems that the local chalcids took time to recognise the new gall and to find the host larvae inside. After all, few gall wasp species in Britain gall the acorns and none form a large knobbly gall like a knopper. In the late 1970s, the first parasitoids to be recorded were Torymus cyaneus and Mesopolobus amaenus, and two ichneumonids (Mastrus diminuens and Gelis formicarius) and a diapriid (Spilomicrus stigmaticalis) were also recorded, all attacking the host larva. M. amaenus continued to be quite plentiful but the other species remained rare or have never again been recorded; they seem to have been opportunistic species, spilling over into knopper galls perhaps because they happened to be unusually common in the years when recorded. Inquilines were very slow to attack knoppers in Britain: the first, Synergus pallicornis, was recorded in the 1980s but it remained uncommon. The total mortality of A. quercuscalicis larvae caused by all parasitoids and inquilines together was usually less than 10 per cent. By the late 1980s, the variety of parasitoids had increased slightly although mortality was still uncommon. Species attacking other large woody galls in Britain were beginning to discover knopper galls: the endoparasitoid Sycophila biguttata, and the polyphagous ectoparasitoids Mesopolobus sericeus and Eupelmus urozonus, which also attack the galls in the native range of A. quercuscalicis in Europe (see Fig. 213, Chapter 10). Then, between 1990 and 1995, there came a sudden and sharp increase in abundance of parasitoid species, including new ones that were recorded for the first time, more than 30 years after A. quercuscalicis had reached Britain. All attacked the inquilines, especially Synergus gallaepomiformis. The attack rate on the gall causer has remained low, with usually less than 15 per cent of available hosts being parasitised. The larva is apparently well protected inside its inner gall and surrounded by the knopper’s thick outer wall. It is the inquilines, common in the outer wall and more easily reached by the parasitoids, which have increased the size of the community.

Recruitment of parasitoids to the small sexual galls of A. quercuscalicis on Q. cerris in Britain (there are no inquilines) has followed a different pattern, without the 20-year time lag. The parasitoid species are all native, and some of them were already attacking the small bud galls of A. kollari, common in Britain 100 years before A. quercuscalicis arrived. Leaf and catkin buds on the tree are not far apart when young and the local parasitoids clearly had no difficulty in finding the catkins. Mortality of the causer in the catkin galls is always much higher than in the knoppers, across the whole of the native and invaded ranges of A. quercuscalicis. Unsurprisingly, the small thin-walled catkin galls are more vulnerable than the knoppers.

In both sexual and agamic galls of A. quercuscalicis, the parasitoid assemblages in the invaded range at first were subsets of those in the native range in southeast Europe. Over time, the galls have recruited more species of parasitoids and the variety of species in the invaded range has converged on that in the native range. This has occurred despite differences in the cynipid faunas of the two areas and despite the different environments of northern Europe compared to the southeast. It suggests that the host tree, the part galled and the type of gall, and the phenology of the tree (i.e. the timing of its development: when it produces its leaves, flowers, fruits, etc.), are more critical for the parasitoids than climate and other local environmental factors. Parasitoid species recruited to a new gall could come from two sources. Either they are from the local community and have recognised a new food source and have switched to it; or they have invaded from the native range of the gall wasp, having followed it via the same stepping stones, perhaps losing genetic variability en route (see Box 12.1) in the same way as the host. In the invaded area, both sources could provide parasitoids at the same time, so that perhaps different ‘strains’ of parasitoid individuals attack the same host population.

It seems that the invading cynipids have had more effect on the local parasitoid and inquiline community than on the native gall causers because many enemies prefer to attack the invaders. Although the A. quercuscalicis knopper galls were slow to recruit enemies, the agamic galls of other invading Andricus species, which attack buds, may have acquired their enemies more rapidly. Native bud galls are common and both the new and old bud galls developing as near neighbours would readily be searched by native parasitoids and inquilines. Although many parasitoid and inquiline species were already in Britain before the invading gall wasps arrived, some species have arrived with the invaders or followed them here from mainland Europe. An example is the parasitoid Megastigmus stigmatizans (Torymidae), a large species that attacks the galler larva in knopper galls and in the invader agamic bud galls (see Chapter 10), and is not native in Britain. In its native range, M. stigmatizans attacks a wide range of large woody galls. In Britain, it has less choice, restricted here to the galls of the invaders. Two Synergus species, S. umbraculus and S. reinhardi, despite being native in Britain, are now specialists on the invader agamic galls here and more generalist in southeast Europe.

A. quercuscalicis in Britain has the potential to affect the parasitoid community by altering the sex ratio of the parasitoids that attack its tiny catkin galls. The same species (all Mesopolobus species) also parasitise the host larva in the slightly larger sexual bud galls of A. kollari, A. lignicolus and A. corruptrix on Q. cerris (see Chapter 10 and Figs 212b and 213b) and the host larvae in some native species on Q. robur and Q. petraea also available in spring (e.g. Neuroterus quercusbaccarum, N. numismalis, Andricus quercusramuli and Biorhiza pallida). Nearly all the Mesopolobus individuals emerging from the A. quercuscalicis catkin galls are males, while the invader bud galls and the native galls provide a larger food supply and produce a sex ratio that is more nearly 50:50 males and females. In years when A. quercuscalicis is particularly common at a site, this imbalance might affect the ability of the parasitoids to find mates and so could affect their numbers in future generations.

Most heteroecious Andricus species that require Q. cerris are not found in Spain and Portugal–Q. cerris is not native in the Iberian refuge and has not been planted there. A. kollari is an exception. The genetic structure of Iberian A. kollari is a subset of the genes of southeast-European A. kollari, indicating that Iberia was colonised from the east. It seems that this migration occurred at least one glacial cycle ago, perhaps well before the end of the Pleistocene. So, A. kollari is native to Iberia in the present interglacial even though it evolved in southeast Europe. Until the 1990s, it was suspected of having an asexual-only lifecycle in Iberia, having lost its sexual generation. But females emerging from Iberian marble galls are now known to lay their eggs in buds of cork oak Quercus suber, a black oak species in the section Cerris and closely related to Q. cerris. Q. suber is native to southwest Europe and the coast of northwest Africa, and eastwards to Corsica, Sardinia, Sicily and the west coast of Italy. A host shift from Q. cerris to Q. suber enabled A. kollari to colonise Iberia. Other species in Andricus, and in Callirhytis, Neuroterus and Synophrus, also exploit Q. cerris in eastern Europe and Q. suber in the west, so this host shift is not unique, but it is rare. The host shift has not occurred in reverse, though. A. kollari from Q. suber has not shifted back on to Q. cerris, and Iberian-derived individuals do not extend north of southwest France, the northern limit of the native range of Q. suber. Unlike Q. cerris, Q. suber has rarely been planted outside its native range (although it has been introduced into California). In northwest Europe, including Britain, Q. robur and Q. petraea derive from the Iberian glacial refuge rather than the southeast European refuge due to the more rapid warming of the climate in the west (influenced by the Atlantic Ocean and its ameliorating affect on the climate). In contrast, Q. cerris and A. kollari come only from the southeast. The Q. cerris–A. kollari and the Q. suber–A. kollari meet along a narrow front near the estuary of the River Gironde in southwest France, the natural northern limit of Q. suber, and they do not interbreed. This suggests that they should be classed as two distinct species, and they have recently been separated: A. kollari is the species in most of Europe and is endemic to southeast Europe, and a new sibling species, A. hispanicus, is endemic in Iberia. In western Italy, both Q. cerris and Q. suber are native; probably Q. suber originally spread eastwards from Iberia via North Africa. Perhaps A. hispanicus spread with it: there may be another front between the two sibling species in western Italy. In northern Italy, the marble gall wasp originates from the Balkans and uses Q. cerris and is A. kollari s.s., but there is no information on southern Italian A. ‘kollari’ populations.

So, the host shift of Andricus kollari from Q. cerris to Q. suber for its sexual generation has led to a new species in southwest Europe. Has this shift had any effect on the next trophic level, on the parasitoids of the old and new A. ‘kollari’? The host gall wasp on Q. suber has diverged genetically from its parent on Q. cerris, so its parasitoids might also have diverged. DNA analysis has been used to trace the origins of one parasitoid, Megastigmus stigmatizans (see also above). This parasitoid has a broad distribution, parasitising host larvae in marble galls derived from both Q. cerris and Q. suber sexual generations throughout its native range from Iberia to Iran and in its invaded area further north too (Hayward & Stone, 2006). In southeast Europe, M. stigmatizans attacks ten heteroecious Andricus species all causing large woody galls in their agamic generations, but in northwest Europe it is less polyphagous, and in Iberia it is thought to specialise on A. hispanicus only. M. stigmatizans has apparently tracked A. kollari over two time scales: over millennia between the glacial refuges of southeast and southwest Europe and over a few centuries following the human plantings of Q. cerris across northern Europe. Before these plantings, M. stigmatizans was not found in northern Europe. Analysing its populations across the Q. cerris–A. kollari and Q. suber–A. hispanicus front in southwest France indicates that this parasitoid has mirrored its host. It first evolved in the southeast Europe glacial refuge and then spread westwards into Iberia, following A. kollari and parasitising hosts in marble galls on a variety of different white oaks (Quercus s.s) that derived from sexual galls on Q. suber. This host shift in Iberia apparently occurred just once, as in the host A. hispanicus. Iberian M. stigmatizans are particularly genetically uniform, much less diverse than those in southern, central and southeast Europe.

Escape of heteroecious species from their glacial refuges, and of their parasitoids too, seems not to have been common. A. kollari probably reached Iberia and evolved into A. hispanicus long before M. stigmatizans caught up with it. Both species must have migrated during interglacial periods, following the oak forests as they spread across Europe. A. hispanicus seems to have appeared in Iberia at least one interglacial before M. stigmatizans, perhaps 200,000 or 300,000 years before. (But dating such movements is not an exact science.) M. stigmatizans certainly reached Iberia well before the present interglacial, but this started only 15,000 to 12,000 years ago. Glacial periods have always been longer than interglacials, so during most of the intervening period A. hispanicus would have been isolated in the Iberian refuge, safe from attack by M. stigmatizans (and perhaps other parasitoids) from the east.

M. stigmatizans and A. kollari have also expanded their ranges into northern Europe over only the last few centuries, following the human plantings of Q. cerris. All of the A. kollari are derived from Q. cerris ancestors from southeast Europe. And A. kollari from the east has not moved into Iberia in recent times, a move that would have required a second host shift on to Q. suber. A. kollari and A. hispanicus cannot cross the divide in southwest France in either direction. But M. stigmatizans has been less restricted: individuals in France north of the Gironde estuary, and further north in areas where Q. cerris has been planted, have ancestors either from Iberia or from southeast Europe. Not only have the parasitoids from Iberia managed to utilise marble galls on a variety of white oak species, they have also shifted from Q. suber–A. hispanicus to Q. cerris–A. kollari. Thus M. stigmatizans from the Iberian glacial refuge is no longer confined there, unlike A. hispanicus. Conversely, M. stigmatizans from southeast Europe has not invaded Iberia in the last few hundred years; in southwest France, its range expansion has been one-way, out of Iberia. In northern Europe, therefore, M. stigmatizans on A. kollari derive from two sources, from Iberia as well as southeast Europe. The parasitoid has been more flexible than its host, able to recognise marble galls on a variety of white oaks whether derived from sexual galls on Q. cerris or Q. suber. Although heteroecious gall wasps spreading into northern Europe, including Britain, all derive from southeast Europe, any parasitoids behaving like M. stigmatizans will follow them from Iberia as well as southeast Europe. This could affect native and invading oak gall communities in the north as the foreign parasitoids invade. In Britain, M. stigmatizans is restricted to heteroecious Andricus species, but other parasitoids may have a broader host range and may attack native species too, and have unpredictable effects on the parasitoid communities.

A few other black oak species apart from Quercus cerris have been planted outside their native range and are acquiring invading cynipids, although more slowly than Q. cerris has done. Cork oak, rarely planted outside its native Iberia, has been introduced into California and there it is attacked by the European gall wasp Plagiotrochus amenti (= P. suberi), probably introduced accidentally with the host plant. Holm oak Quercus ilex, introduced to Britain from southern Europe around 400 years ago and common in the south of England, has recently (since 2004) become host to three further invaders, all species of Plagiotrochus (P. quercusilicis, P. australis and P. coriaceus).

Heteroecious species of gall wasps are known only in the genera Andricus and Callirhytis and only in the Western Palaearctic, and they alternate only between white and black oaks (in the sections Quercus s.s and Cerris). There are 29 species of oaks in these sections in the Western Palaearctic. In the Eastern Palaearctic, white and black oaks are more species rich: 17 species are recorded in the Himalayas, 32 species in China and six in Japan (Abe et al., in Raman & Gupta, 2007) and there may be more to be found. Perhaps host-alternating gall wasp species exist also–but there are none known and the total number of oak gall wasps recorded is less than in northern Europe and in Britain. It seems certain that many oak gall wasps remain to be discovered in the Eastern Palaearctic. The Quercus subgenus Cyclobalanopsis, the cycle cup or ring cup oaks, is found exclusively in Asia and gall wasps, perhaps some heteroecious species, may be found on these oaks too.

Gall wasps have rarely become major pests. An exception is Dryocosmus kuriphilus, which is the one gall wasp in the tribe Cynipini to gall sweet chestnut Castanea (Aebi et al., in Osaki et al., 2006). It induces a many-chambered gall on chestnut trees and has one generation a year. It is native to China where it is parasitised by 11 chalcid species including some that also attack oak gall wasps, and is not a pest there; probably the parasitoids control its numbers in its native region. But in the 1940s it was accidentally introduced to Japan where it is a major pest, severely reducing the crop of chestnuts produced by attacked trees. In 1999, it was found in Nepal, also accidentally introduced from China or, perhaps, Japan. A chalcid parasitoid, Torymus sinensis, specific to D. kuriphilus, has been introduced to Japan from China and has successfully controlled it, and will probably also do so in Korea, which has a similar problem. More recently, D. kuriphilus has arrived in North America and Europe where it is also reaching pest status (more information on the biological control of D. kuriphilus is included in Chapter 13).

In Japan, Korea and Italy, several parasitoids attack D. kuriphilus, although they have not prevented it from becoming a pest. The parasitoids are species with a broad host range that also attack local oak cynipids right across the Palaearctic. Examples are Eurytoma brunniventris, Eupelmus urozonus, Ormyrus pomaceus and Sycophila species (see Chapter 10). The pattern may be different in the USA; there the species has not been adequately studied. Some of these cosmopolitan parasitoids also attack cynipid hosts on sycamores or on roses, and Eupelmus urozonus even attacks an inquiline cynipid in galls of cecidosid moths on Rhus in South Africa. The time lag between arrival of the invading chestnut gall wasp and development of its parasitoid community has been very short. There apparently has been no problem of detection of this new host by native parasitoids despite different environments, and host-shifting has been easy, even for endoparasitic species (e.g. Sycophila variegata and S. biguttata). Even though D. kuriphilus is unique in attacking Castanea, it is closely related to the oak cynipids and the gall clearly is sufficiently similar to them for the oak parasitoids to exploit. The main difference seems to be the lack of inquilines in the chestnut galls: these are rare in the invaded range and are unknown in the native area in China. Perhaps, because the inquilines feed on plant tissue, they adapt less easily than parasitoids to a new host plant. There are two Castanea species in Europe. C. sativa is native to Turkey, Greece and the southern Balkans, and perhaps also Italy, and like Quercus cerris, it has been planted north-and westwards as far as Britain. C. crenata is native to Japan and has been planted in Italy, Spain and Portugal. C. sativa is present in most of the Western Palaearctic and it is possible that D. kuriphilus will spread as far north as Britain, in the same way as the oak cynipids on Q. cerris have spread from southeast Europe. Compared to Andricus quercuscalicis knopper galls in Britain, D. kuriphilus is acquiring parasitoids more quickly. Knopper galls were largely free of parasitoids until they recruited inquilines, more than 20 years after arriving in Britain. In Italy, only ten years after its introduction, there are already more parasitoids (more than 60 species) on D. kuriphilus than on all the oak cynipids in Britain. Probably polyphagous parasitoids that attack woody oak galls will readily include D. kuriphilus in their host range should it ever reach Britain. In the long run, host-specific ecotypes of parasitoids may evolve, genetically distinct on D. kuriphilus on chestnuts and on Andricus on oaks. And cynipid galls on sycamores in southern Europe could become alternative hosts too. It is possible that all invading species, D. kuriphilus as well as the oak cynipids, may cause changes in local parasitoid communities and this, to be fully understood, will require long-term monitoring.

IN CONCLUSION

This chapter has considered only two aspects of the biogeography of gall insects: their pattern of distribution across the world and changes in the ranges of selected species over a number of time scales. More information on the distribution of each group of gallers can be found in Chapters 4 to 9, particularly in their associated boxes. Explanations suggested for the distributions are necessarily tentative at present; we need more information on galls in the tropics and eastern Asia as well as in other parts of the world before patterns can be adequately explained. But with publications on galls appearing all the time (see, for example, the review articles in Raman et al., 2005; Osaki et al., 2006; Raman & Gupta, 2007; Stone et al., in press; and their lists of references), information on distribution should accumulate in the future.

The ranges of galling insects have changed in the long-distant past and more recently, and continue to do so today. The movements of the cynipid species described are known in impressive detail and, coupled with analysis of their DNA, fascinating accounts of their evolution and spread are emerging. Studies of this type require detailed and long-term monitoring and a lot of manpower; despite this, we hope that they will continue in the future and will be expanded to include species in other galling groups–particularly in the Cecidomyiidae, the family that includes more gall causers than any other. The invasions of oak cynipids into Britain have not, it seems, had detrimental effects on the native cynipid community and the variety of native galls has not declined. This implies that the invaders have colonised without competing with native species, that there were empty niches available. So, invading oak gall wasps are increasing the biodiversity of galls in Britain, a welcome note to counter the continuing loss of species resulting from destruction of habitats, global warming and other detrimental effects of humans on the native fauna and flora.

CHAPTER 13

Galls and People

IN TIMES PAST, AND MUCH more so than at the present day, people were good at exploring the properties of things around them and putting them to good use. Galls were no exception and people have been using galls for a variety of purposes since the fifth century BC and probably earlier. They have also attracted some superstitions and folk customs. In this chapter, the uses to which galls have been put are considered; these are mostly historic. We then look at the ‘downside’, the cases, both ancient and modern, where galls have harmed people or their activities, even when the galls were not recognised as such. Finally, we deal with the main modern use of galls and their causers: as agents for the biological control of weeds and, by contrast, examples where it has become necessary to introduce parasitoids to control gall-causers that themselves have become pests.

The next chapter, Chapter 14, forms a pair with this one. It describes some of the early studies and reports on galls and the way in which knowledge and understanding have increased over the centuries.

GALLS AS MEDICINES AND DRUGS

The earliest use of galls was as herbal medicines. The first records are from the Classical Greeks: Hippocrates in the fifth to fourth centuries BC and Theophrastus in the third century BC. By ‘gall’, they and most subsequent references to galls until recent times meant the Aleppo gall or gall-nut, caused by Andricus infectorius. The identity of the Aleppo gall has been confused throughout history, probably because of its use and importation all over Europe and further east in Asia for making medicines, dyes and inks and for tanning leather (see below). Until recently, the scientific name generally used was A. gallaetinctoriae or A. tinctoriusnostris, but these probably refer to mixtures of similar species used in trade, including the marble gall A. kollari. So, these names should not be used (Stone et al., in press). A. infectorius is a version of the original scientific name, Cynips infectoria, given to the species by Hartig in 1843. Aleppo galls occur on Quercus infectoria and related oaks. Aleppo galls have a high tannin content, higher than most other galls, and it is this that gives them their astringent* properties and reputation for curing a multitude of ills. In the first century AD, Pliny the Elder listed 23 ailments that could be alleviated using gall-nuts, ranging from treating burns and staunching blood to soothing mouth ulcers and toothache, and mending broken nails. Culpeper’s Herbal of 1653 recorded that oak galls dried up rheums and were effective against diarrhoea, dysentery and bloody fluxes. The patient chewed the inner part of the young galls or drank the liquor, wine or a mixture of water and vinegar, used to quench them after burning them. Galls were also ground and made into an ointment, ‘unguentum gallae’, to apply to the skin. In the seventeenth, eighteenth and nineteenth centuries in Europe and the USA, galls continued to be used as a tonic, as an astringent for haemorrhages and as an ointment. They were used to treat animals too; in 1868, farmers near Harrogate in Yorkshire used an infusion of galls to cure diarrhoea in cows, apparently successfully (Fagan, 1918). Oak apples (caused by Biorhiza pallida and native in Britain) produced less satisfactory results but could be used to treat rheums, fluxes and similar problems. And the bedeguar on rose (caused by Diplolepis rosae) was recorded by Pliny as a successful treatment for baldness when ground up and mixed with honey!

In other parts of the world, other species of galls have been used as medicines. In China, the aphid gall ‘wu pei tzu’ has been used in traditional medicine for millennia. The gall is caused by Schlechtendalia chinensis on sumach Rhus semialata (in the US, sumach is spelt ‘sumac’) and, like the Aleppo gall, it has a high tannin content. It is used for many of the same ailments, too, as an astringent and an expectorant, and as a dressing for wounds and swellings. Tamarisk galls, on several species of Tamarix found from Morocco to India and also caused by aphids, are known locally as ‘tak-out’ or ‘teggaout’ (Howes, 1953) and have been used as astringents too, although their tannin content is not as high. And aphid galls on Pistacia known as ‘afs’ or ‘afs-el-batum’ have been sold by herbalists and druggists in Tripoli and in northwest India, and used as a tonic and an expectorant. Both tamarisk and pistacia galls have also been used to treat coughs, asthma, fever, lack of appetite and stomach pains.

Besides being used as herbal remedies in the past, some galls are used as drugs today. Ergots are the source of one set of drugs. Ergots are caused by the fungus Claviceps purpurea found on several species of grasses (described in Chapter 3), and cause the painful and often lethal disease ergotism (described below). They also have a long history as a useful medicine. The ergots contain alkaloids (e.g. ergotamine, ergometrine, ergotoxine, all derivatives of lysergic acid or LSD; van Dongen & de Groot, 1995) that cause smooth muscles to contract. Controlled small doses of the drug can induce labour by speeding up contractions during childbirth, and they help to control haemorrhage after the birth by constricting the smaller blood vessels and capillaries. The Chinese have used the drug since the eighth century and the Europeans since at least the sixteenth century, well before its properties were understood (Spooner & Roberts, 2005). Nowadays ergots are cultivated commercially on rye crops, particularly in Russia and in Spain, and increasingly are grown as laboratory cultures. The drug was officially recognised in 1820 in the USA and in 1836 in Britain. Its main use continues to be in obstetrics, though it can also be used to treat the pulsating headaches of migraines.

Yew trees Taxus species, and the leafy artichoke gall caused by Taxomyia taxi (described in Chapter 6), are the source of another set of drugs, discovered in the mid-1960s as a treatment for cancer (Hageneder, 2007, summarises the information). Taxanes are poisons produced by all parts of the yew and are lethal in large doses. The gall is involved incidentally if it happens to be harvested with the leaves and shoots used to produce the drug; the concentration of taxanes in the leaves of the gall is no higher than in the ungalled shoots. Like ergots, yew as a medicine has a long history. The Greeks and Romans used extracts of leaves to stimulate menstruation, to treat adder and insect bites and worm infections, and for skin problems and ulcers. From the Middle Ages until the twentieth century, yew was used to treat additional ailments: epilepsy, diphtheria, rheumatism, arthritis, tonsillitis, bronchitis and asthma and to promote abortion. American Indians treated similar conditions and skin cancers with extracts of yew, and used these as a general tonic and ‘to gain strength’. The anti-cancer properties of taxanes come from their ability to inhibit cell growth. Nowadays, taxanes are used to treat several cancers: of the breast, ovaries, prostate and lung, and some types of leukaemia.

OTHER USES OF GALLS

Besides being remedies for diseases, oak galls could be used for burning in lamps. Theophrastus describes two types (see also Chapter 14). A large dark-coloured resinous gall that burns well without adding oil is the Bassorah (i.e. Basra) gall caused by Cynips insana (Senn, 1941), now classified as a form of Andricus quercustozae (A. quercustozae f. insana). The second gall Theophrastus described as a soft woolly ball; this is the gall of Andricus theophrastea and was used as a wick in oil lamps. In general, though, the main uses of galls, other than as food (see below), made use of the high concentrations of tannins found in some galls, particularly those on oaks.

Tannins are substances with astringent properties produced by plants, and are best known for their ability to combine with the proteins in animal hides to convert them into leather. They are common in woody plants but are rare in herbaceous species. Rapidly growing tissues, such as meristems and young leaves, have little tannin but mature leaves, stems, roots and fruits of some trees, e.g. oaks, have high concentrations; oak bark has been used for tanning leather for centuries. Tannins protect the plant because they make the plant tissues less palatable to herbivores and protect against growth of fungal pathogens. This is evident in spring and early summer in British oak woods when the abundance of caterpillars and other grazing insects feeding on the young foliage is much higher than later in the summer when tannin levels in the leaves have increased. Besides oaks (family Fagaceae), other trees have high tannin concentrations; pistacias and sumachs (Anacardiaceae) are particularly rich and some galls on these have high tannin contents too. In all of these plants and their galls the tannic acids are chemically similar.

Not all cynipid galls on oak have a high tannin content. Aleppo galls (caused by Andricus infectorius) are the richest with about 65 per cent (Fagan, 1918, records 75 per cent) and knopper galls (A. quercuscalicis) contain 50 per cent. Marble galls (A. kollari) and artichoke galls (A. foecundatrix) have only 14 per cent and 10 per cent respectively, and oak apples (Biorhiza pallida) and spangle galls (Neuroterus species) have less than 5 per cent, a concentration similar to oak leaves without galls. The common spangle gall (N. quercusbaccarum) was bottom of the league of those tested with no measurable tannin content (all figures are per cent dry weight; from Leach, 1986). Chinese galls, caused by the aphid Schlechtendalia chinensis on sumach, are rich in tannins with about 60 per cent, comparable to Aleppo galls, and some pistacia galls have high levels too.

Galls in dyeing, tanning and ink-making

The colouring properties of tannins have been used for dyeing cloth and skins for millennia, from classical times until the twentieth century. Ink-making and tanning are rather less ancient and also continued into the twentieth century. Aleppo galls were a common article of trade for the Egyptians in the fifth and fourth centuries BC and the Greeks in the third and second centuries BC, and were used to dye wool and woollen cloth. Pliny (first century AD) stated that oak galls produced a black dye for colouring hair and the best preparation for dyeing leather and skins. For producing a permanent black dye it seems there was no substitute for the Aleppo gall. Most valuable were ‘blue’ galls, i.e. young galls before the insect had emerged. ‘White’ galls, lighter in colour and in weight and with insect emergence holes, were inferior but could be used for dyeing light-coloured linens. Aleppo galls were exported all over Europe for dyes and later also for tanning leather and ink-making. Some were shipped to Bombay (Mumbai) and then exported as Bombay or Indian galls, often to China. Large quantities from the Mediterranean arrived in London, for the American as well as the British market. Although there is a rich variety of cynipid galls on oaks in North America, there are no records of their use by American Indians or by the European settlers; sealskin furs were transported to London to be dressed and dyed before being exported back to America. By the early twentieth century, though, and probably earlier, the US had its own dyeing industry based on imported Aleppo galls; in 1914, gall-nuts worth $17,174 were imported from Baghdad (Fagan, 1918).

Other galls have been used to produce dyes although much less commonly. In Turkey and Iraq, an oak cynipid gall (probably the Bassorah gall Andricus quercustozae) was mixed with cochineal and tartar to make a fine scarlet dye known as ‘Turkey red’. Other common names for the Bassorah gall are the ‘mad apple of Sodom’, the ‘Dead Sea fruit’ and the Mecca gall. Knopper galls (Andricus quercuscalicis) were often substituted for Aleppo galls, and marble galls (A. kollari) were used occasionally where they are native, in southeast Europe and the Middle East. According to Fagan (1918), gall-nuts were used by Somali women as a tattooing dye, probably the Aleppo gall or a similar species. In China, Japan and northern India, the Chinese gall (Schlechtendalia chinensis) produced a black dye and, in southeast Europe and in India, pistacia aphid galls produced a scarlet dye when mixed with other ingredients. Like the oak cynipid galls, the aphid galls produce the best dyes when young, before the aphids have left the gall.

Galls, particularly Aleppo galls, have been used to produce high-quality, permanent writing-inks since medieval times. In the ninth, tenth and eleventh centuries, monks used gall ink for copying manuscripts (although the galls then were often called ‘oak apples’ rather than gall-nuts or Aleppo galls). Aleppo galls continued to be used in Europe and the USA for legal documents that required permanent and durable ink until the first half of the twentieth century. The first inks made, however, were carbon inks and did not include galls. Carbon inks used soot or lampblack made from burning vegetable oil, resin and tar and mixed with water, and with gum arabic added to keep the soot in suspension. The result was a thick fluid so that writing would have been similar to painting. Ancient peoples used carbon inks in 2500 BC (Eusman, 2001). Good quality carbon ink had a blue-black colour that did not discolour with age; but it could easily smudge and did not adhere to a greasy surface, and was easily removed from a document by sponging with water. Chinese carbon ink, also ancient, was similar and was mixed with glue so that it could be moulded into sticks. Carbon inks were used with papyrus on which the pigment was trapped between the fibres, making the writing more permanent than that on parchment or paper. In the Middle Ages, carbon inks had largely been replaced by iron-gall ink, although for a while both were used.

The reaction of tannin and an iron salt to create a blue-black colour has been known since antiquity, recorded by Pliny in the first century AD. A recipe for iron-gall ink appeared in a fifth century encyclopaedia (Eusman, n.d.) and gall inks were increasingly used from the ninth to the fifteenth centuries. It was not until about 1800, however, that tannin derived from oak galls became the most important and widely used ingredient for writing-ink. Iron-gall ink was used until the mid twentieth century–in 1974, the German government still specified gall ink for legal documents. Later, gall ink was increasingly replaced with tannins from other sources, e.g. logwood inks, and nowadays most inks use synthetic aniline dyes. Iron-gall ink is produced by the reaction of the tannic acid in the galls with an iron salt, e.g. ferrous sulphate. The result is a ferrous tannate complex that at first is colourless and water-soluble, allowing it to penetrate the surface of the paper. When exposed to air on the paper, ferric tannate is formed by oxidation; this is an indelible pigment and forms a permanent blue-black ink. Gum arabic is added to make the ink visible when writing, to prevent the ferric tannate from settling out of the solution, and to improve the viscosity of the ink so that it flows more freely from the pen. Further refinements were to add logwood extracts or indigo, which deepen the colour and prevent it fading or bleaching in daylight. Various recipes for gall inks are described in Box 13.1. Aleppo galls with their high tannin content were favoured in writing-ink recipes. Native British galls contain less tannin and marble galls, introduced to Britain in 1834, did not fulfil their potential for ink-making because of their relatively low tannin content. Other cynipid galls from southeast Europe and the Middle East have been imported from time to time, often mixed with consignments of Aleppo galls. The most important non-cynipid gall and with tannin levels similar to the Aleppo gall is the Chinese gall. It has been used for making writing-ink in China, Japan and northern India for as long a time as the Aleppo gall in Europe. The Chinese gall was first imported into Europe in the early eighteenth century and then seems to have been forgotten until 1844 when it became an important ingredient in gall ink. The cost of imported galls was related to their tannin content: blue Aleppo galls cost 70 shillings a hundredweight, green galls (slightly older galls) were 60 shillings, while white galls were only 50 shillings (MacEwan, 1921). Chinese galls were at least as valuable as the best Aleppo galls.

BOX 13.1. Recipes for writing-inks using galls (iron-gall ink) (From Lucas, 1907; MacEwan, 1921; Eusman, n.d.)

By the nineteenth and early twentieth centuries, the best and most widely used inks for writing and for commerce in the western world were produced from Aleppo galls, caused by the cynipid Andricus infectorius (confused for a long time with A. gallaetinctoriae; Graham Stone, pers. com.). In the east, Chinese galls, caused by the aphid Schlechtendalia chinensis, were used.

A basic recipe for blue-black ink using Aleppo galls is described in Lucas (1907):

4 oz. sound Aleppo galls, bruised in a stone mortar.

¹/4 oz. bruised cloves.

2 pints cold water.

Macerate (i.e. allow to ferment) for a month, shaking occasionally; strain through a filter of linen or cotton cloth and press the macerated galls using a wooden press. To the filtrate, add:

1½ oz. ferrous sulphate (known as copperas).

40 grains neutral indigo powder.

Keep for at least a month, stirring occasionally. The tannin in the galls reacts with the ferrous sulphate to produce ferrous tannate, which on exposure to air becomes a blue-black colour, ferric tannate, which is permanent but takes time to develop. The indigo colours the ink blue-black immediately so that it is easier to write with and cloves deter the growth of moulds. Other ingredients can be added as refinements:

Gum arabic (from acacias native to the eastern Mediterranean countries); it keeps the pigment in suspension, improves the viscosity of the solution so that it flows freely from the pen, and produces a deep brilliant colour on the paper.

Sulphuric acid (or other acids); it keeps the pigment in solution and speeds up the maceration process.

The procedure using Chinese galls is different:

7 oz. Chinese galls, coarsely ground.

Thoroughly moisten with distilled water or rain water and keep in a warm place (about 70º or 80º F.) for 8–10 days or until the surface is covered with mould, sprinkling with more water as necessary to keep the galls damp. The mould includes yeasts that provide enzymes to promote fermentation; unlike Aleppo galls, Chinese galls do not contain these necessary enzymes. Add 1 pint of water and heat over a water-bath for an hour, strain and press the galls in a wooden press. Add ½ pint more water, heat for another hour, strain and press again. To the filtrate, add:

3½ oz. ferrous sulphate in solution.

3 grains methyl blue.

As for the Aleppo gall ink, oil of cloves and sulphuric acid can be added to determoulds from contaminating it and to improve the colour of the ink, and to keep it in solution.

In the 1870s, the cost of materials and labour for producing a good-quality writing-ink for bank notes, ‘Bank Black Ink’, is given below (information provided by the Bank of England; Jonathan Briggs, pers. com., 2001). For 60 gallons of ink:

The Bank used, on average, 20 gallons of ink per month. Mr Smee, the Bank of England’s Medical Officer from 1857–77, stipulated that only the best-quality nut-galls and gum should be used, which made the ink used by the Bank much more costly than ordinary writing-ink.

Tannins have also been used for centuries for tanning animal skins, i.e. for hardening hides to turn them into leather. In Britain, a traditional tanning material was oak bark, collected from coppiced woods at least from medieval times. By 1800, there were shortages of bark and two or more materials were being mixed, including imported materials. Wood and bark from tropical trees had been used since the seventeenth century and were often superior to native oak; through the eighteenth, nineteenth and early twentieth centuries they were imported in greater quantities. Aleppo galls were imported for tanning from the Middle East well before 1800. In the eighteenth and nineteenth centuries, there were several unsuccessful attempts to introduce Aleppo galls into Britain–in contrast to the marble gall of A. kollari, which was introduced in the 1830s with dramatic success (see Chapter 12). Its increase and spread was so rapid that it was feared that it would lead to the loss of the acorn crop, important winter fodder for pigs. Marble galls never became a substitute for Aleppo galls as their tannin content is too low, but they did not decimate the acorn crop either.

Tamarisk galls from northwest India were used for tanning too, exported from Bombay to Europe. In Morocco, goat-and sheepskins were tanned with tamarisk galls to produce fine Morocco leather. The fresh galls produce a light almost white leather with a slight rose or violet tint. As it ages, the skin becomes darker and brownish. Several species of pistacia galls from the Middle East and India have been used to produce good-quality light-coloured leathers too.

Although not a gall, another source of tannin used increasingly through the nineteenth century was valonia–the acorn cups of Quercus macrolepis and related oaks native to southeast Europe and Turkey (acorn cups are known as vallonia in modern Italian). In 1885, 30,000 tons of valonia were imported into Britain. The trade declined in the twentieth century; only 5,000 tons were imported in 1949 (Howes, 1953). Valonia tannin produces tough, high-quality leathers used for the soles of shoes. In the 1950s, J. T. Palmer (1992) was a cargo surveyor whose job was to examine materials damaged when in transit for insurance claims. Sometimes these included jute hessian bags containing assorted galls and valonia shipped from Syria and Turkey and en route to a small tannery in Liverpool. By 1992, only two British firms were still using valonia, although only as a tiny proportion of the tanning materials used (most were wood, bark and other products from tropical trees). Galls were used along with valonia until 1976, when the British Dyewood Company of Glasgow still manufactured tannic and gallic acids. (The firm no longer exists; Jonathan Briggs, pers. com., 1996.) The species of galls used was not recorded but, as China, Syria and Turkey were the main sources, one assumes that Chinese galls and Aleppo galls formed the bulk of the imports.

GALLS AS FOOD

There are a few records of galls being used as foods for humans or for domestic animals. The maize smut fungus, caused by Ustilago maydis, is the best known (its lifecycle is described in Chapter 3). It affects the young maize shoots and particularly the cobs, with individual grains becoming swollen and silvery white before turning black as the spores mature. Young galls at the silvery white stage are highly nutritious but should not be eaten after they start to turn black: the mature spores induce allergic reactions and are toxic to animals as well as to humans. The host plants, maize and a close relative known as teosinte, are native to Central America. Maize has been grown in Europe since at least the eighteenth century, in Britain since the nineteenth century, and the smut arrived here with the crop; it has been known in southern England since about 1850. The Aztec civilisation of Mexico was based on maize and the galls as food have probably been eaten since well before the sixteenth century. Nowadays the galls are a delicacy known as ‘ambrosia of the Aztecs’ or ‘huitlacoche’ and are as good to eat as chanterelles, morels and truffles; they are exported in tins to London and elsewhere. A recipe for huitlacoche quesadillas is given in Box 13.2.

BOX 13.2. Quesadillas de huitlacoche (From Banuett, 1992)

1 lb. huitlacoche, coarsely chopped.

¹/4 onion, finely chopped.

1 garlic clove, finely chopped.

1 sprig epazote (Chenopodium ambrosioides), finely chopped.

1 sprig cilantro (Coriandrum sativum), finely chopped.

2 chilis poblanos, roasted and cut into strips.

salt to taste

Cook onion and garlic in a little oil until soft, not browned. Add chilis, epazote, cilantro, huitlacoche and salt, and cook until soft. Put spoonfuls of mixture in centre of uncooked tortillas, fold each in half and fry in hot oil until brown. Serve immediately, with guacamole, a sauce based on avacados. Can also be used to stuff crepes, served with cream.

A few cynipid galls have been used as food. Fagan (1918) noted galls of Disholcaspis weldi being offered for sale on a fruit stand in Mexico City. Children in Mexico regularly ate other oak galls, particularly those caused by species of Callirhytis and especially those covered with nectar; apparently gall nectar makes them sweeter than sugar. In 1558, cynipid galls on Salvia species were regarded as delicacies in Crete and the Middle East (Fagan, 1918); it seems they have an acid, aromatic flavour and are especially good with honey and sugar. They were collected in May and sold in village markets. Liposthenes glechomae on ground ivy Glechoma hederacea, too, is said to taste pleasantly sweet and has been eaten in France and Sweden. Galls of the honeydew gall wasp Disholcaspis eldoradensis on Quercus lobata apparently produce so much nectar that foraging honeybees in California stored 30–40 lb in their hives (Felt, 1940). Galls of Dryocosmus deciduus on oaks in Missouri and Arkansas are known as ‘black oak wheat’ or ‘wheat mast’ and are rich in starch; they were regularly used to fatten cattle, sheep, pigs and poultry.

Other insect galls are eaten occasionally too. In 1811, Linnaeus recorded Adelges galls on spruce in northern Scandinavia being collected like berries (Fagan, 1918; they are described in Chapter 6). The following is an anecdote from Australia:

Our bush tucker delicacy for that night was bloodwood apples–a gall created by an insect which burrows under the bark of the bloodwood tree. These are quite tasty and full of nutritional value, if you don’t think too much about what you are eating or look too carefully at the juicy grub in the middle.

From Land Rover International, May 1999; see Clayton, 1999.

Bloodwood apples, or bush coconuts, occur on bloodwood eucalypts Corymbia species, and can be very abundant in central and northern Australia. They are caused by a scale insect, Cystococcus pomiformis (Eriococcidae). Each mature gall contains a large female up to 4 cm long, with her mouthparts inserted in the inner wall of the chamber and her abdomen plugging the exit hole. The inner wall is a 1 cm thick layer of edible white flesh that provides food for her small male offspring. Aborigines prize this gall as a delicacy. They break it open and eat the female and the male nymphs–she is watery and the nymphs have a nutty flavour–and then scrape out and eat the white coconut-like flesh of the inner gall (Gullan & Cranston, 2005).

SUPERSTITIONS

Over much of historical time, the nature of galls has not been understood and superstitions and folklore have developed around them. Pliny assumed that galls were fruits of oak trees produced in alternate years to acorns. He stated that Aleppo galls grew rapidly like mushrooms, appearing in a single day and shrivelling as soon as exposed to the heat of midday. In later centuries, there were medical superstitions. The thistle gall of Urophora cardui was carried in the pocket, credited with curing haemorrhages, and the rose bedeguar or robin’s pincushion, caused by Diplolepis rosae, prevented toothache. (These galls are described in Chapters 8 and 9, respectively.) Galls were thought to foretell the future too. John Gerard (1542–1612; also Dodoen’s Herball of 1578) records prophesies of oak apples (Biorhiza pallida), beliefs already at least three centuries old: if the oak apple contained an ant, there would be grain in plenty; if a white worm, a murren (i.e. a plague) of beasts and cattle; if a spider, pestilence amongst men. In England, 29 May became Oak Apple Day, which commemorates the restoration of King Charles II to the English throne on 29 May 1660. Porches of houses, lych gates of churches, pub doors and sometimes horses and, later, locomotives of trains were decorated with oak branches. This date was also known as Shick-Shack Day, a much older tradition and nothing to do with Charles II (Robbins, 1992). ‘Shick-shack’ is a term of abuse. On this day, people wore a sprig of oak with an oak apple (though often without) in their buttonhole or hat or carried in the hand. Apples or potatoes (after they had been introduced to England) could be used instead. Sometimes the oak apples were gilded with gold paint. If an oak sprig was not worn, punishments followed, with minor variations in different parts of the country. You were whipped about the hands and face, kicked, punched or sworn at, or drenched with water or pelted with eggs. Or you were ‘nettled’. The same date was ‘Oak and Nettle Day’ in Nottinghamshire; if you had forgotten your oak sprig, your face, hands and neck were whipped with nettles. This tradition continued into the twentieth century at least until the 1930s, meted out by schoolboys nettling the bare knees of smaller boys in short trousers. The oak sprigs had to be removed at midday, when all punishments stopped.

GALL DISEASES AND PESTS

Not all galls are useful or harmless. Several are pests or cause diseases of crops or of forest trees and sometimes cause severe economic loss. A few cause human diseases that can be life threatening. Many groups of galls contain damaging or nuisance examples: bacterial and fungal galls can cause disease, some galls caused by plants can be a nuisance, and several galling nematodes, gall mites and galling insects are pests.

Crown gall, caused by Agrobacterium tumefaciens, causes the best-known bacterial disease; it affects many plants and is a nuisance on fruit trees, brassicas and other crop plants. Diseases now known to be caused by fungi have been known since antiquity and some are gallers. Two of the most serious, causing ergotism and black stem rust disease, are described in more detail below. There are other gall pathogens of crops; examples are wart disease of potatoes (caused by Synchytrium endobioticum), clubroot of brassicas (Plasmodiophora brassicae), powdery scab of potatoes (Spongospora subterranean), peach leaf curl (Taphrina deformans), and diseases of conifer plantations (e.g. canker of larch Lachnellula wilkommii; white pine blister rust Cronartium ribicola). For more information on these and other fungal and bacterial diseases, see Chapters 2 and 3, and Large (1946), Ingram & Robertson (1999) and Spooner & Roberts (2005). Pest galls caused by plants can be a nuisance too, although not usually in Britain; the most serious are probably the dwarf mistletoes (Striga species) that attack cereal and legume crops in semi-arid areas and savannahs of Asia, Africa, Australia and the USA.

Some animal gallers are crop pests. The nematode Anguina tritici attacks wheat seeds causing galls known as ear cockles. These look very like corncockle seeds or like small ergots (caused by Claviceps purpurea; see below). They have not been seen in Britain since the 1950s, probably eliminated by efficient seed-cleaning techniques. The best-known gall mite pest is the blackcurrant mite Cecidophyopsis ribis (see Chapter 6), and many gall midges are pests of vegetable, fruit and cereal crops and of other cultivated plants including trees, shrubs and ornamental herbaceous plants (see Barnes, 1948–56; Nijveldt, 1969; Harris, 2002). Some cereal gall midges are described in more detail below. The most notorious galling insect pest is the grape phylloxera Daktulosphaira vitifoliae, which almost destroyed vineyards in Europe and elsewhere. It is described further below too (see Chapter 5 also). Only one gall wasp, the chestnut gall wasp Dryocosmus kuriphilus, has become a pest, although it is now successfully controlled (see the Biological Control section below).

Ergotism: caused by Claviceps purpurea

The fungus Claviceps purpurea is a pathogen of many grasses, including cultivated cereals, particularly rye Secale cereale. The galls are ergots that replace the grain (the biology of the fungus is described in Chapter 3). The ergots have the distinction of being one of the first galls to be illustrated, by Caspar Bauhin in 1685 in his Theatrum Botanicum; the figure is reproduced by Ainsworth (1976). Ergots, and the alkaloids they contain, have been used in folk medicine for centuries and more recently in orthodox medicine too (see above). But the dose must be strictly controlled. If ergots are eaten in any quantity, they cause severe disease and often death, in domestic animals as well as in humans. An early name of the disease was ‘Ignis Sacer’ (holy fire). Later it became known as ‘St Anthony’s Fire’ after miraculous cures occurred at the church of La Motte au Bois in France where the bones of St Anthony are buried. St Anthony was an Egyptian hermit who lived between 251 and 356 AD (van Dongen & de Groot, 1995). The monastic order of St Anthony became linked with treatment of the disease. Fig. 246 is a woodcut of St Anthony with a victim who has lost a foot and has his left hand enveloped in flames. St Anthony himself also suffered from Ignis Sacer. His suffering is depicted in an engraving by Martin Schöngauer (Fig. 247) and in Bosch’s St Anthony Triptych (1500–05; described in Dixon, 1984). The disease has been recognised throughout the Middle Ages, since at least the tenth century, although its link with ergots was not understood until much later. Through the seventeenth and eighteenth centuries and as it became clear that bread made from ergotted rye was the main cause of the disease, seed-cleaning techniques improved and ergotism became less common. But it remained a hazard even into the twentieth century, especially in poor communities in rye-growing areas. In 1927, there was a mild outbreak amongst Jewish immigrants in Manchester, caused by bread made from ergotted rye grown in Yorkshire. Since then, more serious outbreaks have occurred, in France in the 1950s, India in 1975 and Ethiopia in 1978. Nowadays the fungus is controlled in cereal crops, at least in Britain, but it remains common in wild grasses. The disease is still a problem of cattle, sheep and horses even in Britain; in the 1980s, ergotism increased due to contamination of hay and silage crops.

FIG 246. St Anthony with a victim of gangrenous ergotism, from a woodcut by Hans von Gerssdorrf, dated 1517 or 1551 (reproduced in Barger, 1931; van Dongen & de Groot, 1995).

FIG 247. St Anthony tormented by seven hell-born creatures, usually interpreted as the effects of ergotism. From an engraving by Martin Schöngauer provided by the Wellcome Library, London. (I am indebted to Mark Nesbitt for finding this image.)

There are two forms of ergotism, gangrenous and convulsive or spasmodic ergotism, both with horrible symptoms. Gangrenous ergotism begins with lethargy and pains in the back and limbs, especially in the calf muscles of the legs, followed by swelling of the hands and feet or of the whole limb. Accompanying this is a sensation of violent burning alternating with icy cold–it is this form that is known as St Anthony’s Fire. The small blood vessels in the fingers and toes, hands and feet, become constricted so that the blood supply is cut off and this leads to shrivelling and gangrene. In mild cases, the patient loses fingernails or single fingers or toes; if severe, hands, feet or whole limbs can be lost. Although the severest cases result in death, often the patients survive unless they eat more ergot. Convulsive ergotism has a wide range of symptoms but no loss of limbs. At first there is a numbness of the fingers and toes with ‘formication’, a sensation like ants (sometimes described as mice) running about on or under the skin. Then the limb muscles go into spasm and contract violently so that fingers, toes, hands, feet, arms and legs are flexed and contorted; the effect is excruciatingly painful and limbs cannot be straightened by the sufferer or by anyone attempting to help. More symptoms follow: convulsions, dementia, maniacal excitement and loss of sight. Recovery is possible if the attack is mild, but often the patient dies or is left with mental and physical disabilities. In the eighteenth century, ergotism was particularly common in France and Germany, the two forms more or less separated by the River Rhine, the gangrenous form to the west and the convulsive form to the east. The convulsive form may be related to deficiency of vitamin A; in the worst affected parts of Germany, people were desperately poor with no milk or cheese to improve their diet, and no choice but to eat large quantities of rye bread that was often contaminated.

Ergotism in the fourteenth to eighteenth centuries in Europe may have been more common than is generally realised and may have been a major cause of the high mortality and low fertility rates of those times. The disease may also have helped to cause the outbreaks of religious fervour that were common. Convulsive ergotism, with its fits, hysteria and maniacal dancing, was often associated with witchcraft, and may have led to the witch hunts of East Anglia and Salem, New England (the Salem witch trials occurred in 1692). Both East Anglia and Salem, Massachusetts, have dry sandy infertile soils where rye was the main cereal crop grown.

Black stem rust disease of wheat: caused by Puccinia graminis

Black stem rust has caused devastating disease of cereals since Roman times, particularly of wheat crops, and is still a problem in some parts of the world. Like most rusts, it has a complicated lifecycle with five spore stages and requires two host plants; a grass is its primary host and barberry Berberis vulgaris (also Mahonia in the USA) is its secondary host (its detailed lifecycle is described in Chapter 3). Galls develop only on barberry and do little damage but, on wheat, the attack can be devastating–at best the yield of grain declines, but severe attack leads to shrivelling of the grain and wilting of the plants so that the whole crop is lost. The fungus diverts nutrients from the developing grain, water is lost from the whole plant and all infected plants die. The disease appears on the wheat crop in late spring or early summer, streaking the leaves and stems with orange (the uredinia and urediniospores) and colouring the whole field so that it appears to be on fire. The spores spread from field to field like wildfire and, in late summer and autumn as the black teliospores develop, the remains of the crop and the soil are dusted black, like soot.

In Italy in the seventh century BC, the Romans sought to avoid diseases of their cereal crops by propitiating Robigus, the Corn God. Each year on 25 April, the usual date that the first rust symptoms appeared on the wheat plants, they celebrated the festival of ‘Robigalia’. A procession followed a particular route, prayers were offered and a red-coloured dog and a sheep were sacrificed in order to safeguard the crops from disease. Epidemics of cereal crop disease have occurred throughout history, although the causes were often obscure. Greek and Roman accounts of the symptoms were often not precise enough to distinguish the effects of the rust disease from powdery mildew, smuts or bunts, or of loss of crops due to drought. In medieval Europe, barberry was thought to exert a malign influence over wheat (due to its reddish leaves and red berries) so causing the rust disease, so nearby barberry bushes were often destroyed. In the seventeenth century it was realised that fungi were associated with many plant diseases but they were assumed to be symptoms of diseases rather than their cause. It was not until the nineteenth century that fungi were shown to cause particular diseases and their lifecycles worked out. In 1854 the Tulasne brothers identified the spore stages of P. graminis on grasses, and in 1865 Anton de Bary finally worked out its whole life history, including the necessary alternation between wheat and barberry.

Barberry is a European species often assumed to be native in Britain. Like the cereal grasses, though, it probably evolved in the Middle East and was spread throughout Europe by the Romans. Both host plants, and the rust disease, were introduced to North America by European settlers and then spread across the wheat-growing prairies. Berberis was introduced as a useful plant, used for hedging to control stock and providing berries for making preserves. Losses of the wheat crop due to black stem rust were often severe and led to suffering and starvation in many pioneer communities. When the link with barberry was suspected, strenuous efforts were made to eradicate the bushes. The famous ‘Barberry Law of Massachusetts’, passed in 1755, was an order to destroy all plants before the deadline of 13 June 1760, with penalties imposed if the law was ignored (Ainsworth, 1976). In some states of the US, it is still an offence to plant barberry. Barberry is more widespread in North America than in Britain and the rust still causes crop losses there. P. graminis now attacks native American Berberis and Mahonia species, so there are reservoirs of the disease and epidemics have continued into the twentieth century. In warmer climates too, in the USA and elsewhere, P. graminis can survive all year without the spore stages on barberry, so shortcutting the lifecycle and making the disease more difficult to control. In Britain, the disease is now rare on cereals and is mostly found on rye-grasses Lolium species. Barberry is an uncommon plant too, not often found outside gardens. In the twentieth century, new varieties of wheat resistant to black stem rust were bred and have controlled the disease effectively. But often this control proved to be temporary because the rust mutated into variants capable of surviving in the new hosts. Nowadays, genetic engineering improves resistance in wheat and can provide protection much more rapidly than the old breeding programmes and so can keep ahead of the rust’s ability to adapt. Major epidemics are now rarer than they used to be.

The grape phylloxera: Daktulosphaira vitifoliae

In the mid-nineteenth century, a devastating ‘blight’ hit the vineyards of Europe, first appearing in France in the summer of 1865. The vines suddenly turned red or yellow and withered and the leaves dropped off before autumn, and the roots became black and shrivelled with very little development of fine rootlets and root hairs. A few grapes developed but failed to ripen and most plants did not survive the winter. The blight spread in a characteristic fashion, advancing through a vineyard from row to row. Several causes were suggested: inclement weather (drought, late frosts) or a fungal or insect-caused disease but, at first, no pathogen or pest was found. Eventually, yellow swellings were noticed on the roots and these proved to be the eggs and adult females of an aphid. Several names were given to it and eventually, in Europe, it became known as Phylloxera vastatrix. It is native to North America, however, and it was named there before it reached Europe; its current name is Daktulosphaira vitifoliae. But it is a variable insect and may turn out to be a complex of several closely related species. The phylloxera reached Europe on infested American Vitis species brought in as breeding material when attempts were made to control mildew that was common on European vines. This attempted control inadvertently resulted in the much more serious problem of the grape phylloxera. D. vitifoliae has a complicated lifecycle (described in Chapter 5), nearly as complex as a rust fungus. By 1868 the lifecycle on the vine roots was worked out; it involves simple galls that, when on the main roots, cut the supply of nutrients and water to the above-ground parts of the plants, causing them to wither and die. The root galls damage the plants directly too and allow fungal pathogens to invade the roots, an additional cause of root death. The females lay eggs continuously, without mating, and cause a rapid increase in the root gall population. So, the root stages increase and spread throughout the summer, affecting the vineyard in the manner of a spreading blight or rust fungus. Eventually, a generation of aphid nymphs climbs out of the soil and up the plants; they grow into adult females and are dispersed throughout the crop by the wind. This leads to the sexual stages and overwintering eggs and more generations of nymphs that induce the leaf galls. After introduction in 1865, the phylloxera spread to every vine-growing area in France and thence to neighbouring countries. In 1875, losses to the French wine industry reached £50 million a year and vineyards elsewhere were also badly infested. By 1885, the aphid had reached Algeria, Australia and the South African Cape, accidentally introduced by settlers taking European vines with them.

The grape phylloxera was threatening the wine industry everywhere, except in its native North America. In the US, D. vitifoliae feeds on many local species of grapevines and was originally thought to be a different species from Phylloxera vastatrix in Europe because it mainly caused galls on the leaves rather than on the roots as in France. It was realised later that the American species can feed on the roots but fewer individuals do so than in Europe. Leaf feeding causes less damage than root feeding, so that American vines were relatively little affected. In Europe, there is only one grapevine species Vitis vinifera, although this has been bred over the centuries into about 2,000 varieties. On V. vinifera, root feeding by D. vitifoliae became very common and its effect was disastrous, in contrast to its mild effect on the American vines. Vineyard owners in France attempted to solve the problem by planting resistant American species, but the resulting wine was unpleasant and inferior. The solution proved to be to graft European vines on to American rootstocks; this produced resistant vines that yielded palatable wine. But this solution does not work everywhere and may not be permanent. Research continues to improve the management of vineyards and, particularly, DNA studies are beginning to sort out whether D. vitifoliae is just one variable species or a complex of forms that perform differently on different species and varieties of Vitis (see Granett et al., 2001).

The reason that D. vitifoliae became so devastating in southern Europe was the milder winters. In much of North America, the insect goes through its full lifecycle, including the sexual stages and the tough overwintering eggs, necessary to withstand the cold winters there. The eggs hatch into the leaf-feeding stages and the galls on the leaves do much less damage than those on the roots. On some American Vitis species, too, the root forms do not develop. In the milder climates of Europe, the sexual forms and leaf-feeding stages are by-passed and the asexual females multiply through many generations on the roots; it is these generations that devastate the plants.

Gall midge pests of cereals: the Hessian fly Mayetiola destructor on wheat and Orseolia oryzae and O. oryzivora on rice

The Hessian fly is a serious pest of cereal crops in Europe, North Africa, North America and New Zealand, and in North America it is considered to be the most destructive insect pest of wheat known. Barnes (1956) published a detailed account of its history, biology and control and there is more recent information in the Crop Protection Compendium (CAB International, 2007). Although wheat is the main crop attacked, rye and barley are also affected, as well as some wild grasses. The hatchling larvae feed under the bases of developing leaves and cause swollen stem galls at the bases of young plants or at the internodes of older plants. Larvae pupate under the leaf-sheaths in seed-like puparia and there can be up to six successive generations during a growing season. Sustained attacks weaken plants and reduce yields, and during outbreak years grain losses are sometimes a quarter to half of the crop. Losses in the USA were especially high during the nineteenth and early twentieth centuries after the pest had been accidentally introduced from Europe. In about 1776 during the American War of Independence, infested straw bedding was allegedly imported to America by troops from Hesse in Germany–hence the common name. Hessian fly was one of the first insect pest species to be controlled by plant breeding; in 1945, the year before resistant wheat varieties became generally available, overall losses in the US were estimated at $37 million, compared with annual losses of only about $16 million by the 1980s. In North America, Europe and New Zealand (where it was accidentally introduced before 1888), there has been a general reduction in losses, mainly resulting from plant breeding. By 1992, more than 25 genes that conferred resistance in wheat had been identified. The use of resistant varieties combined with clean cultivation and management of sowing dates, to limit the periods during which crops are exposed to the egg-laying adult midges, have been especially effective in North America. This is a good example of integrated pest management. In Europe, there have been periodic outbreaks, but this pest has never been very damaging probably because, here, a complex of natural enemies usually controls it effectively.

The Asian rice gall midge Orseolia oryzae has been known as an important pest of rice since the late nineteenth century but the African pest O. oryzivora was not recognised as a distinct species until 1982, after serious outbreaks developed in West Africa. The two pests cause similar damage to cultivated rice. Barnes (1956) reviewed the information available on the Asian species and more recent accounts of both species are included in the Crop Protection Compendium (CAB International, 2007). Female midges lay eggs on the leaves of young rice plants and the young larvae, which hatch after a few days, crawl to the growing points of the young shoots where they feed for two to three weeks. A long tubular gall, similar to an onion leaf, develops around each larva and prevents the development of the rice inflorescences, resulting in complete loss of grain from infested shoots. The larvae pupate at the bases of the galls and, later, the pupae wriggle their way towards the tips, where they cut holes through which the adults emerge. This ensures that in crops flooded by rain or irrigation water, the adults emerge well above the surface of the water. The galls are usually 25–30 cm long but may be longer and they are probably the longest of any insect gall. Crop losses of 10 per cent are common and much greater losses may occur locally. In both tropical Asia and Africa, rice production has become increasingly intensified using high-yielding new varieties; but these are susceptible to the gall midges and crop losses have increased. The main control for both pests is by breeding resistant varieties of rice.

GALLS AS AGENTS FOR THE BIOLOGICAL CONTROL OF WEEDS

A serious threat to biodiversity, both of plants and the animals that depend on them, is the invasion of weeds that take over an area and replace the natural vegetation. This problem is, perhaps, second only to the destruction and loss of habitats. In addition, weeds reduce the potential of farmland: grassland weeds reduce the value of grazing land and weeds contaminate crops. So, weeds need to be controlled. Chemical control, with herbicides, is often effective in the short term but it is expensive, it needs to be regularly repeated and it damages the environment. Mechanical control, e.g. hoeing and weeding by hand, is often too labour intensive to be economically viable anywhere but in a small garden–although it is still the main method used in areas of subsistence farming throughout the world. Biological control may be costly in the early stages but has the potential to control a weed permanently and at no future cost. Integrated pest management, i.e. using a mixture of methods integrated with the weed’s habits and lifecycle, has potential too.

A plant becomes a weed if it increases in an uncontrolled fashion to the detriment of the habitat or crop, usually because it has been introduced to a new area where environmental conditions are ideal for it and where there is little competition from the native vegetation. Back in the native country, the plant species typically is not super-abundant but is controlled by a suite of natural enemies. In the new country, the plant has no serious enemies because the local insects are not adapted to feed on it. Biological control, or biocontrol, is usually based on the deliberate introduction of natural enemies of the weed, herbivores or pathogens from the native country that feed on the plant and reduce its abundance to a level at which it is no longer a nuisance nor economically damaging. There are three types of biological control. Classical biocontrol is the traditional type that introduces an enemy of the weed, usually an insect from the weed’s native area, which becomes established. Inundative biocontrol involves the repeated introduction of an enemy that does not become permanently established and includes spraying with the agent in water (e.g. nematodes or pathogenic fungi) as a bioherbicide. Manipulative biocontrol modifies the habitat to promote competition by native plants. If it is successful, biocontrol is the cheapest, most permanent, and most environmentally friendly option and may be the only sustainable method in the long term. Classical biocontrol is the main method discussed here because some biocontrol agents used are gall insects.

The main costs involved in a successful biocontrol programme are for experiments to ensure that a potential agent is safe to release and will not attack non-target plants. This involves testing the feeding and oviposition preferences of the insect while in quarantine conditions (and, in Canada, costs on average two scientist-years and $800,000; 2002 figures, Harris & Shamoun, 2002). Subsequent costs of successful programmes are low as the agent disperses itself to new patches of the weed and reduces it wherever it appears. The ideal outcome is for the agent and weed to be in balance: when the weed is abundant, the agent increases rapidly and reduces it, and when the weed is uncommon the agent becomes rare. It is important that the agent does not die out completely, so that it is on hand should there be a resurgence of the weed.

Gall causers that are successful agents tend to have certain characteristics:

They have narrow host ranges and do not attack related crop plants or native species.

The galls develop in rapidly growing meristematic tissues that are produced in abundance by the weed.

The galls are substantial in size and form strong nutrient sinks. The most efficient agents induce galls that divert nutrients from elsewhere in the plant via the vascular system, resulting in galls that are heavier than the ungalled plant parts.

Galls need to be common, to grow rapidly and to develop throughout the growing season of the weed, particularly when the galled parts are young.

And the galls must reduce the weed to a level at which it is no longer of economic importance. Often they do this by reducing seed production rather than by killing the plant.

The best-known examples of biocontrol of weeds using galls involve thistles and knapweeds. These plants have been introduced accidentally, without their native enemies, from Europe and Asia to new countries, particularly Canada and the USA, South Africa, Australia and New Zealand. In areas of the new countries with suitable climates and soils, particularly in prairies and pastures, many species have become weeds supplanting the native vegetation and reducing the area of grazing land for cattle and sheep. The success rate of biocontrol varies, though; some introduced gall insects are more successful than others. Some examples are included below, together with another success story involving a weedy acacia in South Africa, introduced from Australia. The final example concerns an unusual gall wasp that has become a pest of sweet chestnuts and has been successfully controlled biologically by introducing one of its parasitoid enemies.

Nodding or musk thistle: Carduus nutans

Carduus nutans is a weed of overgrazed pastures and disturbed sites, a problem in Canada, the USA, Australia, New Zealand and elsewhere. In western Canada, it spread over the dry prairies and formed dense stands of 150,000 plants per hectare, so abundant that large areas became useless for grazing cattle. The thistles disperse by seeds endowed with parachutes of hairs that allow the wind to carry them far and wide, and they germinate in patches of bare soil, which are plentiful in dry or overgrazed grasslands. The gall-causing weevil Rhinocyllus conicus, native in Europe and western Asia, was first released as a biocontrol agent in Canada in 1968 and in the US in 1969 (its lifecycle is described in Chapter 7). Its effect has been dramatic. It has reduced the dense stands of nodding thistle to scattered patches, surviving mainly in temporary disturbed sites along roadsides and around gravel pits. Fig. 248 shows its effect at a site in the prairies of Saskatchewan, the same view before and after control by the weevil. It has achieved this control on its own and is said to be the fourth most successful biocontrol agent in the world (including all agents, not just gall causers; Harris & Shorthouse, 1996). But this control has taken time. In Virginia, control was achieved in about six years, but in Saskatchewan it has taken about 15 years. This is probably because in the very dry prairies of central Canada there are always plentiful patches of bare soil–and dormant seeds in the seed bank survive for many years and readily germinate to fill these patches.

FIG 248. Carduus nutans in prairie grassland in southern Saskatchewan, Canada, illustrating (a) pre-and (b) post-biocontrol by the gall weevil Rhinocyllus conicus (the cattle trail in the foreground and the farm track top right can be seen in both photographs). (Peter Harris)

In attacked heads, R. conicus larvae feed in the receptacle and on developing seeds. The larvae sever the vascular strands supplying the seeds and this stimulates the growth of callus tissue on which the larvae also feed. Heads containing many larvae produce no healthy seeds and in heads with only one or two larvae, seed number is reduced. Fewer flowering heads develop on attacked plants and those heads that escape attack also produce fewer viable seeds. Adult weevils produce large numbers of eggs and are relatively long-lived, and so can infest large numbers of heads. They are good dispersers too and readily find isolated patches of thistles, up to 19 km away from an infested site. So, the effect on the plants can be considerable. In native areas in Europe, parasitism of the larvae and interference by other thistle head insects often cause high mortality, so that R. conicus densities are relatively low and its effect on the plant is less. In Canada, less than 3 per cent of larvae are parasitised and weevil numbers can be very high, the weevils infesting a large proportion of plants. The outcome has been a 95 per cent reduction in seed production in Virginia and 50 per cent reduction in western Canada, with stands in pastures reduced by up to 99 per cent. The thistle was controlled more rapidly in the US mainly because the seed bank in the soil was reduced more rapidly, but the eventual outcome has been the same–Carduus nutans has been controlled, the native vegetation has returned, and the weed is no longer a serious problem in North America. The thistle still survives in disturbed sites where there is little or no competition from other plants and as occasional scattered individuals. In North America, nodding thistle has been controlled by just one agent, which is unusual; more often, several agents are required to complete the job.

C. nutans is also a serious weed in Australia and New Zealand. In 1988, following its success in N. America, R. conicus was released in New South Wales, the first biocontrol agent to be introduced there. But R. conicus has not controlled the thistle in Australia or in New Zealand. It effectively reduces the number of seeds produced by the flower heads early in the season but has little impact on later heads, and these produce enough seeds to maintain the density of the weed. The problem is that the flowering season is longer in Australasia than in North America and longer than the active period of the adult weevils, and so seeds produced by later flowering heads survive. A second biocontrol agent has been introduced to complement R. conicus–the tephritid fly Urophora solstitialis, which induces a hard woody gall in the flower head. It has a partial second generation that attacks the later heads.

Full-grown larvae of U. solstitialis overwinter inside their woody galls and pupate in the spring, and adults emerge over a six-week period starting in early spring. They mate and the female lays eggs into young heads, drilling through the bracts with her long ovipositor; she lays each egg into a developing floret (in the same way as described for U. stylata on spear thistle, see Chapter 7). The hatchling burrows down the floret into the receptacle, eating through the developing ovule, and gall tissue forms around it. Galls of adjacent larvae coalesce and eventually form a hard multiple gall, a powerful nutrient sink that affects more of the plant than the flower head alone. The progeny of the first eggs laid in the spring produce adults that produce a second generation in later heads, and these cause galls that overwinter. U. solstitialis was released in 1992 in the same area as R. conicus had been released four years earlier (Woodburn, 1996). By 1996, R. conicus had spread more than 15 km from its release point and had reached all patches of the thistle. U. solstitialis’ spread was more limited but it found the later heads missed by R. conicus. It seems that the two insects compete for the same thistle heads early in the season. If U. solstitialis gets there first, its gall will start to develop successfully but, if R. conicus attacks the same head later, its larvae cut the vascular strands supplying the developing seeds and the U. solstitialis’ gall aborts, leading to the death of its larvae. But, if R. conicus is uncommon in early spring, U. solstitialis does well and many survive to produce the second generation, with the result that seed production may be reduced by 70 per cent. So, although R. conicus is the dominant competitor, U. solstitialis can reinforce its effect. U. solstitialis does even better when R. conicus is absent, when it can attack over 90 per cent of heads. But it disperses less actively and does not reach all thistle patches. In contrast to North America, the biocontrol of nodding thistle in Australia has required two agents.

Besides controlling or contributing to the control of C. nutans, R. conicus also attacks other introduced thistles, e.g. Carduus acanthoides, Cirsium vulgare and Cirsium arvense. In Canada, it is less effective against C. acanthoides than C. nutans because, as in Australia, C. acanthoides has a longer flowering period and later heads escape the weevil. R. conicus can reduce the introduced Cirsium species too but only if the weevils originate from the same part of Europe as the weed. Weevils from Mediterranean Europe are more successful on Carduus thistles while those from further north in France prefer Cirsium. In 1968 and 1969 when the first weevils were introduced into Canada and the USA, it was fortuitous that they belonged to the Carduus strain and so successfully controlled C. nutans.

Although biocontrol by Rhinocyllus conicus has been a success story, there are drawbacks. After about 20 years in the US and in Canada, the larvae have been found in the heads of several native thistles, including some relatively rare and endangered species, and they have reduced seed production in all of them. The weevil’s preferred host is still Carduus nutans, in which its larvae grow faster and survival is higher, but it performs well enough in the native thistles and it does reduce their abundance. R. conicus also adversely affects native insects that feed in the flower heads of the native thistles. All of the native thistles in North America are Cirsium species and so the Cirsium strain of R. conicus is more of a problem than the Carduus strain. But after many decades, the behaviour of R. conicus from any source may change and they may shift to related hosts. A change in behaviour can occur as the preferred host C. nutans becomes less common and harder for R. conicus to find amongst the variety of other thistle species present. The weevils may then deposit their eggs on other species, native as well as introduced, and some will survive. Native thistles with lifecycles and flowering periods similar to C. nutans are likely to be the most vulnerable to attack by R. conicus. This long-term problem is a danger in all biological control programmes and, in order to reduce the problem as far as is possible, all related native species must be tested thoroughly before biocontrol agents are released.

Other thistles

Other thistles besides Carduus nutans have become weeds in North America, common enough to require control. Several gall insects and a gall-causing fungus have been introduced to control spear thistle Cirsium vulgare and creeping thistle C. arvense (known as Canada thistle in North America). C. vulgare is a biennial with large flower heads and, like Carduus nutans, it reproduces only by seed. C. arvense has smaller heads and is perennial, spreading by rhizomes as well as by seed, and is a serious problem in many parts of Canada, in wheat crops as well as in pastures.

The European tephritid fly Urophora stylata induces large woody galls in the flower heads of C. vulgare that reduce its seed production (its biology is described in Chapter 7). Insects from Germany and Switzerland were released in 1973 and have become established in British Columbia and in parts of eastern Canada. The gall reduces seed production by 60 per cent or so but the insect dies out when thistle stands become sparse, and this allows the thistle to recover. The seeds can remain viable in the soil for 36 years, too. The gall is a strong nutrient sink that draws nutrients from neighbouring flower heads as well as the attacked one, and heavily attacked stands can be reduced to scattered plants. But this only happens if the insect is well synchronised with development of the plant; timing is more critical than for Rhinocyllus conicus. The first spear thistle heads to flower are usually the largest and produce the largest number of seeds, and it is these heads that need to be galled. A cold spring can delay emergence of the adult flies so that they miss the first heads when their size is suitable for oviposition. Lack of synchrony is common and this prevents U. stylata from controlling C. vulgare on its own. R. conicus also attacks C. vulgare and adds to the decline in seed production. Although spear thistle is not completely controlled in Canada, it has not been as serious a weed as nodding thistle.

A second tephritid fly, Urophora cardui, also native to Europe, causes galls in the stems of C. arvense (see Chapter 8) and has been introduced to Canada as part of the programme to control creeping thistle. U. cardui was released in 1974 and is well established in eastern Canada but it is sporadic in the prairies, surviving only near rivers and ponds. Large galls reduce the growth of the shoots and roots and prevent or delay flowering but, like U. stylata, only if synchronised with early growth of the shoots. Although U. cardui has spread in eastern Canada, its effect is not sufficient to control the weed. Several other agents have been introduced and together have increased the stress on the weed. The root-galling weevil Cleonis pigra was accidentally introduced from Europe before 1933. A galling rust fungus Puccinia punctiformis appeared before 1912, also accidentally introduced from Europe. In addition, several non-galling insects have been introduced intentionally: a tephritid fly Terellia ruficauda whose larvae feed in the flower heads; and two leaf-feeding beetles, Cassida rubiginosa and Lema cyanella. And R. conicus and U. stylata sometimes spill over on to C. arvense and reduce its growth. But despite the number of agents, creeping thistle is not completely under control in Canada.

Knapweeds: Centaurea species and Acroptilon repens

Several knapweeds are notorious weeds in the prairies of Canada and northern USA. Particularly noxious are the non-British diffuse and spotted knapweeds Centaurea diffusa and C. biebersteinii (originally thought to be C. maculosa), native to southeast Europe and southwest Asia. The related Russian knapweed Acroptilon repens adds to the problem in the hotter, drier prairies of southwest Canada. C. diffusa is normally biennial but occasionally develops in a year or is a short-term perennial. The other species are perennials. All were probably introduced with Turkestan alfalfa seed from the Caspian Sea area. The knapweeds thrive in the dry prairie grasslands of southwest Canada and reduce forage for livestock by up to 90 per cent. C. diffusa and C. biebersteinii infested nearly 30,000 hectares in British Columbia in 1972, an area that had increased to 83,000 hectares by 1984 and, in western Canada as a whole, threatened an estimated 8.4 to 10.7 million hectares. In 1979 alone, the loss to farmers was calculated as $58 million (Harris & Shorthouse, 1996; Harris & Shamoun, 2002). A. repens thrives in the driest parts of British Columbia and can form stands of 64 stems per m². These densities displace the native vegetation completely and the knapweeds cannot be used as fodder; Russian knapweed in particular is toxic to livestock. Control by herbicides would be prohibitively expensive on this marginal grazing land. Fortunately there are several insects specific to these knapweeds and most are gall causers, and these are being used in biocontrol programmes.

Three Urophora species induce galls in the flower heads of the Centaurea species: U. quadrifasciata, U. affinis and U. jaceana (their biology is described in Chapter 7). U. affinis was the first to be released in 1970, introduced from France, and became established in Alberta and British Columbia. More introductions followed, from Ukraine. U. quadrifasciata was introduced in 1972 from Ukraine and also became established in British Columbia. U. jaceana was accidentally introduced from Europe sometime before 1986 and survived in eastern Canada but not in the west; it has not played a part in reducing abundance of these weeds. U. affinis and U. quadrifasciata complement each other and together they have a greater effect on the knapweeds than either does alone. U. affinis causes a more complex gall that is a more powerful nutrient sink than that of U. quadrifasciata; its gall sequesters nutrients from other parts of the plant as well as from the galled head and does more damage than the simpler U. quadrifasciata gall, especially in the larger heads of the spotted knapweeds. U. affinis can be very common, too, sometimes exceeding 3,000 galls per m² (Harris & Shorthouse, 1996). U. quadrifasciata is a useful addition, however, because its adults are more active and quickly find remote patches of the knapweeds. This species also has two generations a year so it increases faster than U. affinis. Once established at a site, these two species reduce the number of knapweed flower heads and therefore numbers of seeds produced by up to 90 per cent. But even this high figure is not enough for complete control; so additional biocontrol agents have been introduced.

Sphenoptera jugoslavica is a beetle (belonging to the Buprestidae) whose larvae gall the roots of diffuse and spotted knapweeds and whose adults eat the leaves (see Chapter 8). The gall prevents small knapweed rosettes from growing upwards to form a flower stalk and thence flowers and seeds, and large rosettes produce fewer seeds if they do manage to flower. The beetle is native to the steppes of southeastern Europe and in 1976 it was introduced to Canada from northern Greece. It is established in British Columbia and the northwest US and adds to the seed reduction caused by the Urophora species. It is particularly successful on C. diffusa, whose seed production has been reduced by 95 per cent using the Urophora species and S. jugoslavica together, and the ground covered by the weed has reduced from 50 per cent to 14 per cent. This reduction is below the replacement level for C. diffusa, and it has declined further since the mid-1990s. At least in hot dry prairie sites, C. diffusa has been controlled using three agents (Fig. 249) and the native grassland species have returned. C. diffusa is still present but now covers only about 10 per cent of the ground. But the transition from dense stands to scattered patches and isolated plants has taken more than 13 years to achieve because, as for Carduus nutans, it takes time for the seed bank in the soil to be reduced. Although S. jugoslavica has less effect on C. biebersteinii, these knapweeds have also declined in dry sites mainly because U. affinis affects them to a greater extent than C. diffusa. As the native grasses return and increasingly compete with the spotted knapweeds, these weeds should also reduce to acceptable levels, controlled with three agents plus plant competition.

FIG 249. Centaurea diffusa in prairie grassland near Penticton, British Columbia, Canada, illustrating (a) pre-and (b) post-biocontrol by three gall insects, Urophora diffusa, U. quadrifasciata and Sphenoptera jugoslavica ((a) is a close-up of part of the site shown in (b)).

Although the knapweeds have been controlled successfully in hot dry prairies where drought is common, in damper more mesic sites, further biocontrol agents have been required. The root weevil Cyphocleonus achates has successfully helped to reduce spotted knapweeds on light soils, and the root moth Agapeta zoegana has helped reduce all of the knapweeds on more mesic sites generally. Neither of these insects causes a gall. The larvae burrow into the roots, the weevil in the centre and the caterpillar in the outer layers, and deprive the plant of its storage tissues. The root enclosing the weevil larva does become swollen due to accumulation of callus tissue, but the larva is almost fully fed by this time and does not use the callus as food. So, the swelling is not a true gall. C. achates was first introduced into Canada in 1987 and A. zoegana in 1982.

The Centaurea knapweeds have finally been controlled in the prairies of Canada and northern USA although this has taken time as well as several agents. As well as those insects described here, other species have been tested for their suitability for biocontrol. But most have either not passed the rigorous testing procedure, because they also attack desirable or harmless species, or they have been released but have failed to survive.

Acroptilon repens is another knapweed that still causes problems, particularly in the hot, dry interior of British Columbia. A gall-causing nematode worm Subanguina picridis, introduced from Armenia and Tadzhikistan, is having some effect. S. picridis causes galls in the stems, buds and leaves of the plant and these induce a strong nutrient sink for about half of the growing season. In addition, rates of transpiration from the galls are nearly twice as high as from the ungalled stems, and the attacked plants tend to dry up rapidly in dry sites (the gall is described in Chapter 8). The worms survive best in prairies with hot dry summers but with moist conditions in spring. In such areas, a continuous attack for three years will solve the problem of this weed. The worms were first released in 1976 but failed to become established. Now they are grown in tissue culture in the laboratory and sprayed on in water each year as a bioherbicide, an example of inundative control.

The long-leaved wattle: Acacia longifolia

In the 1830s, Acacia longifolia was intentionally introduced from Australia into South Africa to bind and stabilise sand dunes. But it soon became an uncontrollable weed. It produces large numbers of seeds that are long-lived and accumulate in the soil, where they can reach densities of 34,000 per m². The seeds are fire-adapted; fires stimulate them to germinate en masse, producing seedlings in abundance and displacing the local vegetation. An Australian chalcid wasp Trichilogaster acaciaelongifoliae belongs to the Pteromalidae, a family in which gall-causing is unusual, and is famous for being one of the most successful biocontrol agents in the world, after Rhinocyllus conicus. T. acaciaelongfoliae was introduced into South Africa in 1982 and 1983 (Dennill, 1988). Its galls develop in the flower buds and prevent pod formation. The galls are spherical and woody with a central chamber that contains the chalcid wasp larva. They fuse together in clusters in the inflorescences preventing development of the seeds. If attacked by inquilines, the galls develop additional chambers that form knobbly outgrowths. The galls can be exceedingly common and can reduce seed production by 99 per cent. Each gall forms a strong nutrient sink that particularly affects the early growth of the reproductive organs. The wasp induces the tree to produce more galls per branch than its normal number of inflorescences and this diverts large quantities of nutrients away from normal growth. Nearly a quarter of the mass of the tree above ground can be in the form of galls, so heavy that they sometimes cause substantial branches to break off. Overall, galling by T. acaciaelongifoliae reduces both reproduction and vegetative growth of this weedy tree, which explains why the wasp is such an effective biocontrol agent. Following the success of this chalcid, a second Trichilogaster species has been introduced into South Africa to control golden wattle Acacia pycnantha and it seems to be becoming as successful as T. acaciaelongifoliae.

The chestnut gall wasp: Dryocosmus kuriphilus

Gall wasps do not usually become pests, even those that have accidentally or intentionally been introduced to new countries. The one exception is Dryocosmus kuriphilus, a global pest of sweet chestnut Castanea. It is native to China, and since the 1950s it has spread to Japan, Nepal, Korea, the USA and Italy, France and Slovenia in Europe. Although it is not a pest in China, it severely damages the chestnut crop in the invaded countries.

Dryocosmus kuriphilus belongs to the cynipid tribe Cynipini, most of which gall oaks Quercus species. This species is unusual in being the only member of its tribe to attack sweet chestnut. It induces many-chambered galls in buds and has a single generation each year, producing only females (it is parthenogenetic). Its effect on the trees is severe: fruit yield is reduced by 50–75 per cent and heavy attack also reduces timber production and can kill the tree (Aebi et al., in Osaki et al., 2006). Chestnut growers exchanging nursery stock and young trees that were not subjected to quarantine procedures have spread it around the world. In 1941, D. kuriphilus was accidentally introduced to the Okayama area of Japan and within 25 years it had spread throughout the country. In Korea, it was first recorded in 1958 and spread across the country in 37 years. It reached Georgia in the USA in 1974 and, two years later, it had spread and was becoming a major pest. It reached Europe more recently; it was first noticed in 2002 in Piemonte, Italy, and was soon found over an area of 160 km². It probably had been introduced in 1995 or 1996 when eight chestnut cultivars were imported from China. By 2005, it had spread throughout Italy and was recorded for the first time in neighbouring regions of France. Despite knowledge of the virulence of D. kuriphilus, 1250 chestnut plants were imported into Slovenia from Piemonte, with no quarantine precautions. However, after a warning from the phytosanitary service of Piemonte, Slovenian workers traced nearly half of the imported plants and destroyed those infested. But there is little doubt that the species has survived in Slovenia and it will probably spread further in Europe.

After its introduction to Japan, attempts were made to control D. kuriphilus. Chemical pesticides proved to be ineffective because the gall protects the young stages of the wasp and the adult stage is brief. For about 20 years, a breeding programme that produced resistant varieties of Castanea kept the pest under control, but around 1960 a strain of D. kuriphilus appeared that could successfully attack these varieties. It was apparent, in the areas in China where the cynipid was native, that it did not seriously affect chestnut production and so perhaps its enemies controlled it naturally there. After a search in China, 11 parasitoid species in five chalcid families were found (Aebi et al., in Osaki et al., 2006). One species, Torymus sinensis, proved to be effective; it was host-specific and had a lifecycle closely synchronised with its host’s lifecycle. A biocontrol programme started in Japan in the late 1970s. In 1979 and 1981, 5,000 D. kuriphilus galls were imported from China to Japan and reared in cages and, in 1982, 260 mated T. sinensis females were released at one site, a fruit tree research station in Osaka. The parasitoid population increased and the adults dispersed efficiently, and soon affected the rate of attack of the gall wasp pest: this reduced from 40 per cent in 1983 to only 3 per cent in 1988. This is well below 30 per cent, the level at which the gall wasp reaches pest status. In Japan, 20 years later, D. kuriphilus is attacked by a rich community of parasitoids, 24 species in seven chalcid families and one braconid, and these include Chinese species introduced with the galls as well as native Japanese species that also parasitise oak cynipids. One inquiline Synergus species has been found in the chestnut galls but it is rare and does not add significantly to the parasitoid community in the gall (in contrast to the knopper gall on oak, Andricus quercuscalicis; see Chapters 10 and 12). The community in chestnut galls seems to be richer in Japan than in China, but the gall is certainly under-studied in China. In Japan, the parasitoid community on D. kuriphilus seems to have stabilised and species shift easily from chestnut galls to oak galls and back again; host-shifting is not a problem for these parasitoids.

In Korea, although T. sinensis has never been introduced deliberately, it is found throughout the country. It is, though, a different strain from the Chinese species and is probably native. Originally it specialised on oak cynipids but, as in Japan, it now attacks both chestnut and oak gall wasps. In Italy, and after only five years, 15 chalcids in six families have been recorded on D. kuriphilus and all of these also attack oak cynipids. They may all be native species derived from oak galls. T. sinensis has also been introduced into Italy from Japan but has failed to become established. There have been problems of synchronisation between the parasitoid and the host, and trials are continuing. It is possible, however, that the native parasitoids shifting from oak gall wasps may control the chestnut gall wasp naturally, with no further intervention.

Despite the harmful effects of a few over the course of time, galls have mainly proved to be beneficial, most importantly in their widespread use for making permanent writing-inks. Galls continue to be of value to humans, now and probably in the future, mainly for their use in helping to control weeds without the use of chemical herbicides.

CHAPTER 14

Galls in History

LIKE ALL MODERN NATURALISTS, cecidologists ‘stand on the shoulders of giants’, using the discoveries of those who came before them, and often taking them for granted. The history of cecidology is littered with examples of confusion and superstition, but also with inspiring examples of meticulous observation and the application of rigorous scientific method.

Throughout most of history, until well into the nineteenth century, the word ‘gall’ meant the Aleppo gall, which probably involved more than one species. This was the gall of commerce, important for dyeing cloth, tanning leather and for making ink (see Chapter 13). In Classical Greek, the Aleppo gall is or ; as mentioned in Chapter 1, this comes from a root meaning something gushing or bubbling forth, ooze or outgrowth; it meant the dye or ink and, by association, the gall too (Arthur Cain, pers. com.). The word ‘gall’ is derived from the Latin galla, an excrescence produced on oak trees, and this has led to galle (French), Galle (German) and gal (Dutch) (Redfern et al., 2008). The taxonomy of the Aleppo gall has recently been clarified and the species has been renamed Andricus infectorius (previously known as A. gallaetinctoriae or A. tinctoriusnostris; Stone et al., in press).

The Classical Greeks in the fourth century BC knew considerably more about galls and cecidology than did European peoples throughout the next millennium and until the nineteenth century, apart from a few more enlightened pockets of knowledge in the seventeenth century, led by John Ray in England and Marcello Malpighi in Italy. Malpighi was the first to understand that galls, although formed of plant tissue, were caused by insects. But, in general, galls remained subjects of superstition for the next two centuries. They have occasionally entered the limelight of major scientific debate, especially in the long-running controversy over spontaneous generation. Knowledge of types of galls increased during the late nineteenth century and throughout the twentieth century as more species were described and their structure became more clearly understood. But our understanding of how they develop is still obscure and is the basis of scientific work continuing today.

This chapter describes how knowledge of galls advanced, from the time of the Classical Greeks until the end of the nineteenth century. Knowledge also has advanced through the twentieth century and continues today. These advances are considered too recent to be ‘history’ and so are not included here; many aspects are discussed in other chapters.

THE SCIENTIFIC GREEKS: THEIR KNOWLEDGE OF GALLS

As with much else in western science, the first references to galls that we know of come from Classical Greece. The earliest reference was to the Aleppo gall and its use in medicine, made by the physician Hippocrates (about 460–377 BC; see Chapter 13).

Surprisingly, the great naturalist Aristotle (384–322 BC) did not mention galls. His fame as a naturalist rests mainly on his Historia animalium, which is the oldest existing monograph of general zoology. In accounting for the development of scientific thought and method, Moore (1993) stated that Aristotle ‘defined the field, outlined the major problems, and accumulated data to provide answers’. It is hard to accept that Aristotle was unaware of galls. He did know about the ‘caprification’ of the fig, that fig insects were essential for figs to ripen, but he did not realise that galls were involved. Fig flowers are enclosed inside the fig ‘fruits’ (i.e. the syconia) and only fig insects can transfer pollen from the male to the female flowers (see Chapter 7 and Fig. 104 in Box 7.2). The fig insect induces galls inside the female flowers and her larvae develop inside them. This is one of the few examples of a gall and its inducer being essential to the survival of both plant and insect. Aristotle may not have been unfamiliar with galls, however. His book on plants is lost (Arthur Cain, pers. com.) and, as galls were regarded as being parts of plants, this is where he would have included them. Aristotle was important in the history of cecidology, as in other branches of natural history, because he taught his students the art of detailed and accurate observation, and the importance of collecting information, asking questions and formulating hypotheses. One of his pupils was Alexander the Great; another, his favourite, was Theophrastus (371–286 BC).

BOX. 14.1. The galls of Theophrastus on deciduous oaks

Theophrastus’ galls are interpreted by Senn (1941), including the host oaks that he recognised, and including the reasoning behind his identifications. Many of the names Senn used for the gall and for the oak species are old ones, and are updated in the figures (these are modern images, with the photographer of each acknowledged). Senn included a plate of line drawings but all, except Cynips Quercus-Tozae, are from secondary sources and are not included here.

FIG 250. Andricus infectorius asexual Aleppo galls on Quercus infectoria. (Gyuri Csóka)

FIG 251. Andricus caliciformis asexual galls on Quercus sp. (Michael Chinery)

The following notes were made by Senn on identification of Theophrastus’ galls (with their native distribution and up-to-date names from Stone et al., in press).

1. & 2. These are both common galls; they were well known to everyone in the eastern Mediterranean and were regularly used in trade. Theophrastus compared them to each other and did not describe them in full detail because they were so familiar. The Aleppo gall (caused by Andricus infectorius) was not the smallest gall known to him, but the smaller of the two galls familiar to the people. It has biggish warty protuberances and ridges especially towards its distal end and is not as smooth as another gall (probably the marble gall, see 10). It apparently grows very fast. Theophrastus said: ‘after this gall has grown up in a single day, and been subject to the heat (of the day), it dries up and grows no more’. Its native distribution is in southern central and eastern Europe (Hungary and the Balkans). It was imported all over western Europe for dyeing cloth, tanning leather and for ink-making (see Chapter 13).
      The Bassorah gall (caused by Andricus quercustozae f. insana) was the largest of the galls known to Theophrastus, reaching a diameter of 4–5 cm., sometimes as big as an apple. It is blackish red in colour with a resin covering its surface like a transparent varnish and has pointed or rounded structures set around its middle. Unlike the Aleppo gall, it remains fresh for several days and takes longer to reach its full size. In lamps, it burns well once lighted without having to be soaked in oil; the resinous envelope burns with a bright but sooty flame while the spongy interior glows or smoulders. It is native in Asia Minor, Iran and the Levant and in southern Europe from southern Italy eastwards.

3. The mulberry gall (caused by Andricus caliciformis) was described very briefly by Theophrastus, but the comparison with a mulberry is precise. It is smaller than the Aleppo gall, only 1.2 cm across, and is rare; it would not have been noticed by ordinary people. Böhner (1935, quoted by Senn) identified it as Andricus aestivalis (now A. lucidus, the sexual generation), which induces galls on the catkins of Quercus cerris and Q. suber; but the individual parts of this gall are cup-shaped, quite unlike the convex individual fruits of a mulberry. Senn had no hesitation in identifying it as Cynips caliciformis. It is native in southern and eastern Europe from Italy eastwards.

4. The phallus-like gall (caused by Andricus quercustozae) has created many problems in the translation of Theophrastus’ original Greek. Senn stated that Böhner, in 1935, translated it thus: ‘it is said that in an early stage it resembles the male organ but when it has got ripe and hard it resembles a bull’s head’. These two structures are so different that the seventeenth century botanist Caspar Bauhin thought they referred to two different galls. Hort (1916, referred to by Senn) translating the Greek literally, also described it as resembling a bull’s head. According to Senn, though, the Greek words do not describe the shape of the structure (‘like the male organ’) but the thing in action (‘during its erection’). So, during its growth, the gall becomes erect and hard, behaving like a bull. Senn translated the specific passage, with no mention of bulls:

‘Besides this the oak-tree bears another growth, which resembles in its behaviour the privy part of a man; for in the course of its development it rises up, grows hard, and becomes provided with an orifice, and in form also comes to resemble the said organ.’

After the insect has left, the gall becomes very hard with an irregular ridge or circle of projections running round it behind the apex. The orifice is the exit hole of the insect, which if it happens to be near the tip of the gall, makes its resemblence to a penis obvious. It is clear that Theophrastus studied the internal structure of the galls as well as the external: ‘when broken open, this structure shows in its interior a something the size of an olive-stone’. Thus, the structure is hollow, containing a small inner gall inside which the gall insect passes its larval and pupal stages. Theophrastus does not mention an organism inside the ‘olive-stone’ so presumably he did not break it open. This gall is native in central and southern Europe from France eastwards and across North Africa.

5. The felt gall (caused by Andricus theophrastea) contains a hard globular unilocular structure, 6–8 mm long, which tapers at both ends, with the surface covered with a felt of long yellowish white or reddish brown hairs 6–10 mm long. The whole structure looks like a dense woolly ball, 2–3 cm across. It has been referred to Andricus quercusramuli (the sexual gall), but this contains several distinct and separate larval chambers and does not fit Theophrastus’ description. Senn identified it with conviction as Andricus theophrastea. Pliny said it burns well without any oil but Trotter (1903, quoted by Senn) found that it must be soaked well in oil first, unlike the Bassorah gall. It is native around the eastern Mediterranean, locally common in southern Turkey.

6. The hairy honey gall (caused by Andricus lucidus) also presented problems of identification. Böhner (1935, quoted by Senn) thought it might be Cynips (= Andricus) caput medusae or Andricus multiplicatus. A. caputmedusae is covered by distinct and separate hairs, as is required, not like the hairs of the felt gall, which are dense and matted together. A. multiplicatus is a leafy gall (see 8), not like that described. Neither, however, produces a honey-like sap. Andricus lucidus, however, fits Theophrastus’ description well. It is native to central and southern Europe from southern France eastwards to Iran.

7. Theophrastus provided a detailed description of the axillary gall (caused by Andricus foecundatrix), which had not been identified before Senn recognised it. The Greek is difficult to interpret and has led to some odd translations (e.g. Hort, 1916). The first is that it had a ‘hollow stalk’. No oak gall has a hollow stalk; what is meant is a hollow chamber, which is present in the hop or artichoke gall. The second is that it had knobs arising on it ‘like the navel of a man’. Senn explained that this was a mistranslation arising because the ancient Greek for navel in medical terminology is the same as for umbilical cord. Thus, instead of knobs like a navel, Theophrastus was describing the inner scales of the gall, which are narrower and longer than the normal bud scales and pale in colour (an umbilical cord is yellow-white with a ribbon-like shape). A third problem is the colour: ‘the knobs are whitish or black-spotted and the inside of the gall is bright red’. The tips of the outer scales of the artichoke gall often do show dark spots or lines, but the interior is greenish white, not red. Senn suggested ‘the colour might be brighter in Greece where strong sun may bring colour to parts which further north remain green or colourless’. This is true of some species of gall in Britain; a gall in the sun becomes flushed with red, while deep in the tree canopy it is pale green. The final difficulty is that it grows ‘in an axil of the branches’. The artichoke gall, however, develops from a bud in the axil of a leaf stalk. Senn suggested that Theophrastus used a general word that could mean a ‘stalk’ or ‘branch’ and that, here, it should be translated as stalk . A. foecundatrix is widely distributed in Europe, including Britain, southern Scandinavia and central, south and eastern Europe to Iran.

8. Theophrastus’ description of this gall (caused by Andricus multiplicatus) also presented translation problems. One suggestion was that it was not a gall at all but the leaf roll of a tortrix caterpillar. Senn, however, discovered a good illustration and description of Andricus multiplicatus in Darboux & Houard’s Galles de Cynipides (1907) and was convinced that this was the gall meant by Theophrastus. It is native in central and southern Europe from Italy eastwards to Iran.

9. Theophrastus’ description fits the currant gall (caused by the sexual generation of Neuroterus quercusbaccarum) well, except that it does not normally grow near the midrib. It has been identified previously as the cherry gall Cynips quercusfolii. But the cherry gall is not translucent, and Senn suggested that this character was more reliable than its position on the leaf. Theophrastus may have relied on a single observation of a gall in an unusual position, or perhaps he mixed up two galls. The currant gall is a common and widespread species, occurring throughout Europe south of central Sweden, and North Africa to Iran.

10. Theophrastus did not formally describe the marble gall (caused by Andricus kollari) but he compared the Aleppo gall (1) to a smoother gall that is of no use for tanning. Senn deduced that he was referring to another common species similar to the Aleppo gall, and marble galls can easily be mistaken for Aleppo galls (Connold, 1901); he thus tentatively identified Cynips Kollari. It is native to central, south and eastern Europe, from Italy eastwards.

FIG 252. Andricus theophrastea asexual galls on Quercus sp. (Gyuri Csóka)

FIG 253. Andricus multiplicatus sexual gall on young leaves of Quercus cerris. (Graham Stone)

Theophrastus was unusual amongst Aristotle’s followers in focusing his attention away from humans. In his Enquiry into Plants, he did for botany what Aristotle had done for zoology, and he has good claim to the title of ‘Father of Botany’ (Moore, 1993). He was the first to classify plants and described more than 500 species, recognising many as weeds. He also studied galls. In the Enquiry, he described the normal annual structures on trees, such as leaf, flower, catkin, fruit and tendril, and then followed this with a description of ‘others’. The section on oak galls begins ‘The oak bears more things besides its fruit than any other tree’. Apart from other growths, such as fungi and mistletoe, he described ten galls, of which Senn (1941) was able to identify nine with some certainty and with a good guess at the last, which was mentioned only in passing. Box 14.1 compares Theophrastus’ descriptions with Senn’s interpretation of species of the gall wasps and host oaks. For Theophrastus, the word ‘gall’ () meant a growth on oak and, in particular, one on a deciduous oak.

The galls in Box 14.1 are all galls of deciduous oaks. Theophrastus gave different names to growths on evergreen oaks, such as kermes oak Quercus coccifera, cork oak Q. suber and holm oak Q. ilex, names like ‘scarlet berry’ or ‘globule’. It was not until the nineteenth century that the word ‘gall’ was universally applied to similar growths on all plants. There was one major exception, Malpighi, who, in De Gallis (1679), used the term for growths on many other plants (see below).

Theophrastus was familiar with growths on other trees and shrubs, besides oaks. On elm, beech, rose and pistacia, he described ‘sacs, pouches or pockets’ but it seems that he did not regard them as objects similar to the galls on deciduous oaks. He noted that the sac-like structures on Pistacia contained creatures like fleas (he was probably referring to the aphid galls of Baizongia pistaciae; David Wool, pers. com.; Figure 77f, Chapter 5). This aphid gall on pistacia is large and obvious and common in Greece and southeast Europe. He was also familiar with a sac on elm, probably the one caused by the pemphigid Eriosoma ulmi (Fig. 54, Chapter 4). He knew the ‘little spongy sort of thing’ on rose (the bedeguar gall of Diplolepis rosae, Fig. 141e, Chapter 9). He knew too that ‘flies’ often came out of galls; he noted them from the bedeguar and the aphid galls and also from currant galls on oak. He did not realise, though, that the insect was essential for the development of the gall. If he considered the origins of the insects at all, he would have assumed that they arose by spontaneous generation, a notion recurring throughout the writings of both Aristotle and Theophrastus (Arthur Cain, pers. com.; Redfern et al., 2008).

FROM 0 TO AD 1500: NOTHING NEW IN CECIDOLOGY

Along with most of the rest of western science, cecidology went into decline for more than a millennium after the Greeks. Although galls continued to be used as medicines and for dyeing, tanning and ink-making, very little new information was gathered and previous knowledge became interwoven with superstition (see also Chapter 13). Even the Roman chronicler Pliny the Elder showed symptoms of this decline. Pliny was born in AD 23 and died in the eruption of Vesuvius which destroyed Pompeii and Herculaneum on 24 August AD 79. He is important in the history of biology for his Historia naturalis, a compendium covering the physical universe, geography, anthropology, mineralogy, biology, medicine and the fine arts. It was a rather uncritical compilation, but is valuable because it is the only record of what was known at the time (Moore, 1993). But in it, fact and fable tend to be included indiscriminately.

Pliny was familiar with ‘gall-nuts’, i.e. the Aleppo galls of commerce [caused by Andricus infectorius, Fig. 250], which were widely used for dyeing and the tanning of leather and for ink-making. He thought they grew as rapidly as fungi:

Pliny affirms, That the Galls break out altogether in one Night, about the beginning of June, and arrive to their full Growth in one Day…

(From John Evelyn in Sylva, 1644, quoted by Swanton, 1912.)

He assumed that the gall-nut was the fruit of the oak, produced in alternate years to acorns; it developed in a day and shrivelled up immediately if struck by heat (Cosens, 1916). His source was clearly Theophrastus’ description of the Aleppo gall (see Box 14.1). Like the Greek naturalists, Pliny had noticed holes in the galls and had seen ‘culices’ (gnats) in watery pustules in oak leaves (perhaps the currant galls of Neuroterus quercusbaccarum, Fig. 150a, Chapter 9) but he did not connect the insects with the formation of the gall. He also did not think of the galls on different trees as being similar structures; ‘galls’ were the familiar Aleppo galls, and growths on other plants were different. Pliny also recognised the bedeguar on rose (Diplolepis rosae, Fig. 141e, Chapter 9) and the gall of Mikiola fagi on beech (Fig. 81e, Chapter 9). Most of Pliny’s knowledge of the use of galls in healing and in commerce was based on the writings of Aristotle and Theophrastus, and would also have been due to his contemporary Dioscorides, a Greek surgeon who was physician to the Roman army at the time of the emperors Nero and Claudius. In De Materia Medica, Dioscorides described the properties of 600 medicinal plants, including oak galls, and also included animal products of dietary and medicinal value. This work was to be the leading pharmacological text for the next 16 centuries.

After the Roman period, much of the knowledge gained in science and medicine was lost in Christian Europe, but was retained and improved upon in the Islamic world. It then re-entered the west. Early medieval references to galls are few indeed. The Aleppo gall was imported for tanning, dyeing and ink-making (Chapter 13), and there is an early recipe for making ink provided by Rabbi Rashi (of Troyes, France, 1040–1105), an unusual Jewish teacher who broke with a tradition of oral transmission and wrote everything down (Meyer, 1987). It is symptomatic of the age that, following Albertus Magnus (1193–1280), galls were seen in Europe as objects for prognostication and superstition. No progress had been made since Pliny, more than 1,000 years before, in understanding what galls were. They were still regarded as fruits of the oak, like acorns. In fairness, we should admit that confusion between galls and fruits of trees has continued until very recent times. The spinose gall of a scale insect (Fig. 98a, Chapter 6) was thought to be the fruit of the tropical tree Hopea ponga until 1960, when 277 years of misidentification was ended (described in Chapter 6).

THE SIXTEENTH, SEVENTEENTH AND EIGHTEENTH CENTURIES

By the late sixteenth century, very little scientific progress had been made in the study of galls, although their uses and identification were more clearly described. The physician and herbalist John Gerard (1542–1612) described several sorts of gall in The Herball or General Historie of Plants (1597). He followed Theophrastus in restricting the word ‘gall’ to growths on foreign oaks, particularly the Aleppo gall. The quotations and figures that follow are all from his Herball (images from the original provided by Sheffield University Library).

On native oaks, in the section ‘Of the Oke’, Gerard described ‘…a fungous excrescence, which we cal Oke Apples’. He described their uses in treating many diseases, including recipes for their preparation, and indicated their prognostication abilities (see Chapter 13). Although his ‘Oke Apples’ were found on native British oaks (these must have been those caused by Biorhiza pallida, Fig. 145a, Chapter 9), they probably also included Aleppo galls imported from the Middle East. Konrad von Megenberg (1309–37) is said to have coined the term ‘oak apple’ to replace the older ‘gall-nut’ used for the Aleppo gall (Cosens, 1916), and the term probably covered several species. Gerard lists the names used in different languages for these galls:

The Gal Tree is called Quercus, Robur, and Galla arbor: the Gal is called in Greeke : the apothecaries and Italians, keepe the name Galla for the fruite: in high Dutch [i.e. German] Galopfel: in low Dutch [i.e. Dutch] Galnoten: in Spanish Agalla, Galha, and Bugalha: in French Noix de Galle: in English Gaules and Gals.

Perhaps he derived ‘oke apple’ from the German.

Galls according to Gerard are produced by ‘the Gall tree’. He described two kinds of tree ‘The great Gall tree Galla, sine Robur maius’ and ‘The little Gall tree Galla minor’ (Fig. 254).

Of trees that bring foorth Gals, there be divers sorts, as may appeere by the divers formes and sorts of Gals set foorth in this present Chapter, which may serue for their seuerall distinctions, whereof some bring foorth Acornes, likewise, and some nothing but Gals: the figures of some fewe of the trees shall give you sufficient knowledge of the test, for all the Acorne, or Maste trees bring foorth Gals: but those trees whose figures we have set foorth do beare those Gals fit for medicine, and to thicken skins with [i.e. for tanning].

Dioscorides and Galen make but two sorts of Gals, the one little, yellow, full of holes, and more spungie in the inner part [1]: both of them rounde, having the form of a little ball; and the other smooth and eeuen on the outside [probably 10]: since, the later writers have founde moe [more], some having certaine little knobs sticking foorth, like in forme to the Gall, which doth also cleaue and growe without stalke to the leafe [1 or 2]. There is also founde a certaine excrescence of a light greene colour, spungie, and waterie, in the middle whereof nowe and then is found a little flie or worme [9]: which soft ball in hot countries, doth many times become harde, like the little smooth Gall [10], as Theophrastus saith.

In the second paragraph, Gerard describes three or four further galls (the last called an ‘excrescence’) included in Theophrastus’ Enquiry into Plants (the numbers in square brackets are those in Box 14.1), but he describes them less clearly. The first seems to be Theophrastus’ ‘Little Gall’, the Aleppo gall (Andricus infectorius, Fig. 250), which would have holes in it after the emergence of the gall causer and any parasitoids and inquilines. The second may be the Aleppo gall again, which has ridges and bumps on its surface, or it may be Theophrastus’ ‘Dark-coloured Resinous Gall’, the Bassorah gall (Andricus quercustozae, Fig. 237, Chapter 11), which has little knobs sticking forth, although it is much larger (Gerard did not mention its size). Both arise directly from a leaf stalk. Both of these were imported into England and would have been familiar to Gerard. The description, a little ball, smooth and even (10 in Box 14.1), is more appropriate for the marble gall (Andricus kollari, Fig. 156, Chapter 6), which Gerard would not have seen growing in Britain (though it may have been imported; it did not survive in Britain until the 1830s), but he may be noting it from Theophrastus’ description only without having seen specimens. It was not generally used for ink-making in Gerard’s time. The description of the excrescence is very similar to Theophrastus’ ‘Translucent, Watery Leaf Gall’, the currant gall (Neuroterus quercusbaccarum, Fig. 150a, Chapter 6). This is native in Britain and would have been common on British oak trees. Gerard included figures of five galls in his Herball (Figs 255–7) and one of ‘The Gall tree with fruite’ (Fig. 257). The galls could all be the Aleppo gall, which is quite variable, or ‘Galla orbiculata’ may be the marble gall Andricus kollari. From his description, the tree is clearly not a native British oak. He noted, too, that ‘The Gals are founde in Italy, Spaine, and Bohemia, and most of the hot regions’.

FIG 254. The great Gall tree Galla, sine Robur maius and the little Gall tree Galla minor (Gerard, 1597; image provided by Sheffield University Library).

FIG 255. Galla orbiculata, the round gall and Galla oblonga, the long gall (Gerard, 1597; image provided by Sheffield University Library).

FIG 256. Galla viridis, siue omphactis, greene or unripe gal and Galla Asinina, asses gals (Gerard, 1597; image provided by Sheffield University Library).

FIG 257. Galla Galericulata, hooded gals and Galla cum fructa, the gall tree with fruite (Gerard, 1597; image provided by Sheffield University Library).

Reading the second of the two paragraphs above:

The spungie bals which are found upon the branches, are most aptly and properly called Spongiole sylvestris Rosae, the little spunges of the wilde Rose: the shops mistake it by the name of Bedeguar: for Bedeguar among the Arabians is a kind of Thistle, which is called in Greek, that is Spina alba, the white Thistle: not the white Thorne, though the word do import no lesse.

Gerard was familiar with the bedeguar gall on wild roses. He noted that these ‘spungie bals’ were remedies for several conditions (e.g. cures for baldness or for kidney stones), quoting Pliny and Galen (about AD 129–200) as supporting authorities. Gerard objects to the word ‘bedeguar’ for this gall, though it is commonly used today and probably has been used during the four centuries since Gerard’s time. He might have preferred the alternative name ‘robin’s pincushion’. The origin of ‘bedeguar’ is Persian, quoted in an Arabic text (see also Chapter 9), and literally means ‘wind-brought’, referring to thistles; perhaps it related to the wind-blown thistledown as well as the spininess of thistles, reminiscent of the hairiness of bedeguar galls. The Arabic source suggests that Pliny and Galen’s knowledge was transmitted via the Moslem world.

Gerard was more concerned with the uses of galls (Chapter 13) than with their origins or biology. He had noticed the association with insects, and took the view that both the galls and the insects (‘little flie or worme’) arose by spontaneous generation. In this view, he reflected the general belief of his time, expressed cogently by the respected sixteenth-century physician Mathiolus, who not only believed in the spontaneous generation of galls and gall insects but also in their use for prognostication. The disproof of the notion of spontaneous generation is popularly associated with Louis Pasteur (1822–95) and his work on bacteria, but the idea had been controversial for two centuries, particularly because it could be interpreted as contradicting the religious view of creation.

In general ‘like breeds like’, and the notion of a fixed set of species created by God and breeding true thereafter was both comprehensible and in accordance with scripture. There was, however, a contrary observation, that flies, moulds and other creatures appeared spontaneously from dirt and putrefaction. From Aristotle onwards, therefore, a notion of spontaneous generation was held alongside the traditional Genesis account, encompassing lice in hair, mites and maggots in cheese, intestinal worms, snails and frogs falling from the sky, fossils, and galls and the insects associated with them (Moore, 1993). At first sight, belief in spontaneous generation would appear to be blasphemous, contradicting a literal interpretation of Genesis. The great theologian St Augustine of Hippo (AD 354–430) provided an ingenious escape clause (Moore, 1993):

Just as the seed contains all that is necessary in the course of time to grow into a tree, so the universe must be conceived as having had at the same time all things that were to be made in it since God created all things at the same time. [Included here are] also those things which earth and water produced potentially and causally prior to the time they came into being in the form they are now known to us.

But this did not satisfy everyone, and disproof of spontaneous generation in the case of gall insects was sought, in part to affirm the truth of the Genesis account.

It was William Harvey (1578–1657), the discoverer of the circulation of the blood, who first made clear the proposition that ‘ex ovo omnia’–that all come from eggs. His treatise on embryology Exercitationes de Generatione Animalium (1651) carries a frontispiece with Zeus opening an egg from which emerges a variety of animals and plants; the egg is inscribed ex ovo omnia. In fact, Harvey was far from proving his case and continued to follow Aristotle in believing that filth and carrion gave rise to maggots and that intestinal worms arose spontaneously. Even Francesco Redi (1626–98), who demonstrated that meat protected from flies did not generate maggots, continued to believe that intestinal worms and gall insects arose spontaneously. He did not witness oviposition, and assumed that ‘the plant had a vegetable soul which presided at the origin of galls, with their eggs, larvae and imagines’ (see Connold, 1901; Cameron, 1882–93). Redi’s work was in turn followed by Anton van Leeuwenhoek (1632–1723), one of the first naturalists to use a microscope (magnifying up to 200 times). He found that covered meat putrified, even though lacking maggots; he could see bacteria but could not establish for certain how they got there.

The problem of the generation and development of galls, and the relationship between galls and gall insects, was eventually solved by Marcello Malpighi (1628–94), who observed and recorded oviposition by gall insects. His work represents the first major advance in cecidology since Theophrastus 1900 years earlier and he can certainly be regarded as the Father of Cecidology. Before considering Malpighi’s work in detail, it is worth looking at the contribution made by his English contemporary, the theologian, scholar and naturalist John Ray (1627–1705).

Ray took up the new passion for ‘natural theology’, that the understanding of God could be approached by study of his handiwork in the creation, and not through Scripture alone. In 1691, he produced The Wisdom of God manifested in the Works of the Creation…, a work which was frequently revised and reprinted (a facsimile of the 1826 edition was published by the Ray Society in 2005). Its aims were set out in the full title:

The Wisdom of God manifested in the Works of the Creation…viz the Heavenly Bodies, Elements, Meteors, Fossils, Vegetables, Animals (Beasts, Birds, Fishes, and Insects), more particularly in the Body of the Earth, its Figure, Motion, and Consistency; and in the Admirable Structure of the Bodies of Man and other Animals; as also in their Generation, etc. with Answers to some Objections.

There follows in the Preface (Ray, 1826):

That by the Works of the Creation in the Title, I mean the Works created by God at the first, and by him conserv’d to this Day in the same State and Condition in which they were at first made; for Conservation (according to the Judgement both of Philosophers and Divines) is a continu’d Creation.

Ray took a harder line than St Augustine, seeing in the notion of spontaneous generation support for atheism and materialism. Dealing with those insects whose mode of generation remained unknown, he wrote (the quotations which follow come from the fourth edition, 1704, of The Wisdom…, for which I am indebted, with the notes in square brackets, to Arthur Cain):

The Discovery of the manner of the generation of these sorts of Insects I earnestly recommend to all ingenious Naturalists, as a matter of great moment. For if this point be but cleared, and it be demonstrated that all Creatures are generated Univocably [only, unambiguously] by Parents of their own kind, and that there is no such thing as Spontaneous generation in the World, one main prop and support of Atheism is taken away, and their strongest hold demolished: They cannot then exemplifie their foolish Hypothesis of the generation of Man and other Animals at first by the like of Frogs and Insects at this present Day.

Perhaps because of this concern, Ray considered spontaneous generation in great detail, bringing against it his own observations, theoretical considerations and evidence from contemporary authorities, including Malpighi, Redi (with whom he disagreed), Swammerdam (1637–80), Leeuwenhoek, Harvey and Martin Lister (1638–1712). Lister was the physician of Queen Anne, and clearly understood that insects were the cause of galls. As an example of Ray’s theoretical objections, the following quotation illustrates (at some length!) the blend of theology and science involved:

Indeed to me it seems unreasonable that Plants being of a lower Form or Order of Being, should produce Animals; for either they must do it out of indisposed Matter; and then such Production would amount to a Creation [i.e. possible only to God]; or else they must prepare a fit Matter, which is to act beyond their strength, there being required to the preparation of the Sperm of Animals a great apparatus of Vessels, and many Secretions, Concoctions, Reflexions, Digestions, and Circulations [i.e. chemical processes] of the Matter, before it can be rectified and exalted into so noble a Liquor: And besides, there must be an Egg too, for we know ex ovo omnia, to the perfection whereof, there are as many Vessels, and as long a Process required. Now in Plants there are no such Vessels, and consequently no such Preparation of Eggs or Sperm, which are the necessary Principles of Animals.

Ray described the process of egg-laying by gall insects and noted the strict association of particular species with particular plants. He also included a detailed précis of Malpighi’s work, published as De Gallis in Latin in 1679 (see Redfern et al., 2008). Ray noted particularly that Malpighi included as galls all ‘Preternatural and Morbose Tumors and Excrescencies of Plants’ and that such growths can be caused in several ways: ‘excited or raised up either by some Venenose [poisonous] Liquor’ injected with the eggs; or by ‘boring with their terebrae [ovipositors]’ or by larvae hatched from the eggs ‘making their way with their teeth into the Buds, Leaves or Fruit, or even Wood’ of the plant on which the eggs were laid. Ray summarised Malpighi’s conclusion:

We conclude therefore, that Galls and other Tumors of Plants are nothing else but morbose Excrescencies, raised up by the force of the Egg there laid, disturbing the Vegetation and Temper of the Plants, and perverting the Motion of their Humors and Juices; wherein the enclosed Eggs and Animalcules are cherished, nourished, and augmented, till their proper parts being manifested, explicated [unfolded], and hardned or strengthned, they are as it were new-born, affecting to come forth into the open Air.

Ray also noted that Malpighi had watched an insect laying eggs into a bud of an oak, which he then opened and saw that the eggs in the bud were exactly the same as those remaining inside the insect.

Like others of his time, Ray could not solve the problem of intestinal worms by direct evidence, but he expressed the strong conviction that their eggs are ingested. He made the shrewd observation:

I am at present, till better informed, of Opinion, that their Eggs are swallowed with the Meat we eat; and I am rather induced to think so, because Children in their first Infancy, and as long as they are constantly confined to a Milk Diet, are seldom troubled with them.

Marcello Malpighi broke new ground with De Gallis, a chapter devoted to the study of galls in Anatomes Plantarum, an important part of his Opera Omnia; this was published in 1679. In De Gallis (On Galls; the title page is shown as Fig. 258), he described the galls of Italy and Sicily in fine detail. This is the first systematic and scientific study of galls and includes many detailed and accurate illustrations (one plate is reproduced here as Fig. 259). Malpighi was a physician. He graduated in medicine in 1653 at Bologna University and taught there before being appointed to the Chair of Theoretical Medicine at Pisa. In 1660, he returned to Bologna and spent the next 30 years in medical practice there, except for four years (1662–66) as Professor of Medicine at Messina in Sicily. In 1691, he moved to Rome and became physician to Pope Innocent XII. He died in Rome in 1694.

FIG 258. The title page of Anatomes Plantarum, the work containing De Gallis, 1679 (facsimile in Redfern et al., 2008, with permission from the Ray Society).

FIG 259. A plate, Tab. XIII, from De Gallis, depicting the development of the artichoke gall of Andricus foecundatrix, with sections illustrating its constituent parts. On the three lower drawings, additional galls are shown, attached to the artichoke gall; R is a typical ramshorn gall of Andricus aries, and S is the same gall distorted by inquilines (facsimile in Redfern et al., 2008, with permission from the Ray Society).

Malpighi is best known for the major contributions he made to medical science, particularly to human anatomy and physiology, and to embryology, which he studied in domestic hens. His reputation as a founding father of modern medicine is well deserved. But he had other interests too and, using microscopes that were improving rapidly throughout the seventeenth century, he extended his powers of observation to insects and plants. He studied the development and anatomy of the silkworm Bombyx mori and the microscopic structure of plants, and it is this interest that included galls. All of his work was written in Latin and most was published by the Royal Society of London; he was elected a Fellow of the Royal Society in 1669. His medical treatises were translated into English by Adelmann (1966). De Gallis has been translated recently (Redfern et al., 2008); it includes a facsimile of the first edition and an interpretation of the 60 or so galls that Malpighi described, identifying them and including photographs of most of them. Although he concentrated on galls on oaks and particularly those caused by cynipid wasps, he looked at other galls too, on other trees and on herbaceous plants. He described, for example, gall midge galls on beech and dogwood (caused, respectively, by Mikiola fagi and Craneiobia corni) and an eriophyoid gall on poplar, probably that of Aceria populi, containing ‘almost innumerable little orange-yellow worms and eggs’. He noticed nodules (caused by bacteria) on the roots of beans and was puzzled because there were no recognisable creatures inside. For more detail of these and other galls that Malpighi described, see Redfern et al. (2008).

Although galls were well known in the seventeenth century, having been used since antiquity as herbal medicines and for dyeing cloth, tanning leather and for making ink (see Chapter 13), there was no understanding of what they were or how they were formed. A few contemporary naturalists, e.g. John Ray and Martin Lister (see above), realised that insects were associated with galls, but the prevailing view was that they, the galls as well as the creatures, arose by spontaneous generation (a view not supported by Ray and Lister). Malpighi also discounted spontaneous generation and went further by proving that the gall wasps emerging from galls developed from eggs like other animals. He described the development of galls from their smallest beginnings to the emergence of the adult insects and he clearly expressed their dual nature–involving stimulation by the insect, which injected an irritant or poison (‘ichor’) with the eggs, and a reaction by the plant. He dissected many of the galls to discover their internal structure. He watched gall wasps laying eggs into buds and then dissected the bud and the female and found that the eggs inside each were identical. In this he was well ahead of his time, in demonstrating that the emergence of adult insects from the galls had nothing to do with spontaneous generation. Even so, the belief that flies and germs spontaneously arose from putrescence and decay was not finally discarded until the experiments of Louis Pasteur in the nineteenth century.

Malpighi was most familiar with gall wasps; in these, the females lay eggs directly into the plant tissue that will develop into the gall. He also suspected that, in other groups, the eggs may be laid some distance away and that it is the larvae that insert themselves into the plant and induce the gall. But Malpighi did not fully appreciate how important a living, feeding larva is for the complete development of the gall, even in cynipids. Otherwise, his understanding of how galls develop was much the same as it is now. Exactly how the insect manipulates the plant is a subject of active current research.

During the eighteenth century, more species of gall-causing insects and their galls were described but there was little advance in understanding how galls developed. René Antoine Ferchault de Réaumur (1683–1759) noted that the ovipositors of different insects varied and suggested that they might cause different wounds and result in galls of different shapes and sizes (see Cameron, 1882–93). He agreed with Malpighi and others that irritation of the plant by the ovipositor played a part in gall development but he thought, too, that feeding by the larva together with ‘heat’ from the egg might be involved. Réaumur also described galls caused by eriophyoid mites, but he regarded the tiny, worm-like creatures he observed as insect larvae.

William Derham, a contemporary of Ray who wrote his biography, confirmed Malpighi’s observation that not all insects found in galls are themselves gall causers (the quotation comes from Swanton, 1912):

I apprehend we see many vermicules, towards the outside of many oak-apples, which I guess were not what the primitive insects laid up in the germ from which the oak-apple had its rise, but from some supervenient additional insects, laid in after the apple was grown, and whilst it was tender and soft.

Thus starts the study of gall communities (see Chapter 10). Derham also noted that more galls were recorded by Malpighi in Italy and Sicily than he could find in southern England and that many were not familiar to him. This was the first observation that distribution of galls might be influenced by geography (see Chapter 12).

A major advance in the eighteenth century was the organisation of biological classification into a natural system, started by Linnaeus (1707–78) and continued by his student Fabricius (1745–1808). In his Systema Naturae (1758), Linnaeus gave each species a generic and a specific name (binomial nomenclature) and classified them into natural groups in which the affinities between species were recognised. This is the basis of the classification system used today. He named over 20 species of gall insects (for 14, he used the word ‘gall’ for the rest, a term like ‘growth’ or ‘sac’), although most are now placed in different genera to those that he used. They included gall-causing aphids, tephritids, sawflies and cynipids infesting a variety of host plants, herbaceous plants as well as trees. His knowledge of how galls developed was limited, though, and it seems that neither he nor Fabricius was aware of Malpighi’s works (Adler & Straton 1894).

The curate Gilbert White (1720–93) is the best-known English naturalist of the eighteenth century. Although he corresponded with foreign naturalists of his day, including Linnaeus, he spent his whole life in and around the village of Selborne in Hampshire. It is surprising that, despite his meticulous observations on birds, mammals and other creatures, he rarely mentioned galls in The Natural History and Antiquities of Selborne (1813 edition, first published 1789). There is just one paragraph:

The stalks and ribs of the leaves of Lombardy poplar are embossed with large tumours of an oblong shape, which by incurious observers have been taken for the fruit of the tree. These galls are full of small insects, some of which are winged, and some not. The parent insect is of the genus cynips. Some poplars in the garden are quite loaded with these excrescences.

He is clearly referring to Pemphigus galls (see Chapter 5 and Fig. 76) and it is odd that his identification is so inaccurate. It seems unlikely that he discussed the galls with Linnaeus; Linnaeus had named Pemphigus bursarius and was familiar with the gall, although he called it a sac. Gilbert White makes no reference to oak galls, which are usually the first to be noticed by casual observers. Perhaps this was because oaks are relatively uncommon on the Chalk Downs around Selborne; the local woods are dominated by beech.

It seems that noone in most of the eighteenth century made a special study of galls, though botanists must have come across them on collecting trips. The British botanist Withering, in British Plants (1776–96), briefly mentions galls on oak and he noted that they were used in dyeing cloth. A footnote added by a Mr Marsham describes the lifecycle of the oak cherry gall insect (Fig. 260):

FIG 260. Cherry galls of Cynips quercusfolii on Quercus robur. (Michael Chinery)

The balls or galls upon the leaves, are occasioned by a small insect with four wings, called Cynips quercusfolii, which deposits an egg in the substance of the leaf, by making a small perforation on the under surface. The ball presently begins to grow; and the egg in the centre of it changes to a worm; this worm again changes to a nymph [i.e. pupa], and the nymph to a flying insect with four wings.

Throughout the whole period from Hippocrates to the end of the eighteenth century, the only galls known were caused by insects (and eriophyoid mites mistaken for insect larvae). Although it was known that fungi caused diseases of plants and sometimes distorted them and caused swellings, these were not known as galls.

GALL-CAUSING FUNGI: KNOWLEDGE UP TO THE EIGHTEENTH CENTURY

Fungi as gall inducers were completely neglected. Before the late seventeenth century and the development of the microscope, they provided evidence for spontaneous generation, arising without seed from decaying substances. Caesalpinus (1519–1603) in De Plantis (1583) was uncertain whether they could be regarded as living (quoted in Ramsbottom, 1953, rev. 1989):

Some plants have no seed; these are the most imperfect, and spring from decaying substances; and they therefore have to feed themselves and grow, and are unable to produce their like; they are a sort of intermediate existence between plants and inanimate nature. In this respect fungi resemble zoophytes, which are intermediate between plants and animals.

Robert Hooke (1635–1703) observed a variety of structures with his new compound microscope, observations that he published in 1665 in Micrographia, or some Physiological Descriptions of Minute Bodies made by Magnifying Glasses, with Observations and Inquiries thereupon. In 1663, he had examined pieces of cork and he described box-like structures that he called ‘cells’. This led eventually (in the nineteenth century) to the realisation that the fundamental units of all living organisms are cells. He investigated a wide variety of objects, including parasitic fungi: ‘…of a Plant growing in the blighted or yellow specks of Damask-rose-leaves, Bramble leaves and some other kinds of leaves’. The ‘plant’ on the rose is a rust fungus, probably Phragmidium tuberculatum, and Hooke’s drawing (Fig. 261) is the first known illustration of a microscopic fungus, which also happens to be a gall inducer. Hooke’s drawing is shown alongside a modern electromicrograph. The swellings on the leaves are telia, with teliospores supported on stalks (see Chapter 3 for a description of the lifecycles of some rust species). Hooke assumed that the fungus (a ‘plant’) arose by spontaneous generation but thought that it probably spread by ‘seeds’ (quoted in Ramsbottom, from Micrographia):

FIG 261. Galls of the rust fungus, probably Phragmidium tuberculatum, on leaves of cultivated roses, Rosa spp. The mounds are telia with teliospores growing from them on stalks. (a) Hooke’s 1665 drawing (Sheffield University Library); (b) a scanning electromicrograph (from Preece & Hick, 1990, with permission).

Though the original cause, or seminal principle from which this minute Plant on Rose leaves did spring, were, before the corruption caused by the Mill-dew, a component part of the leaf on which it grew, and did serve as a coagent in the production and constitution of it, yet might it be so consummate, as to produce a seed which might have a power of propagating the same species…So that the little cases which appear to grow on top of the slender stalks, may, for aught I know, though I should suppose them to spring from the perverting of the usual course of the parent Vegetable, contain a seed, which, being scatter’d on other leaves of the same Plant, may produce a Plant of much the same kind.

FIG 262. Electromicrograph of telia and teliospores of T. anemones on a leaf of A. nemorosa (from Preece & Hick, 1990, with permission).

Facsimile of Fig. 1 in Tab. III of Ray’s Synopsis Methodica Stirpium Britannicum (third edition, 1724). It shows the ‘Conjurer of Chalgrave’s Fern’, which is a leaf of wood anemone galled by Tranzschelia anemones (from Stearn, 1973, with permission from the Ray Society).

It is surprising that Hooke did not see the spores of the moulds he examined. In 1679, Malpighi in De Gallis figured the spores of common moulds growing on second-rate cheese, on the putrescent skin of melons, and on lemons and oranges, on wood and on bread. Hooke regarded the spores as florets borne on an inflorescence (the sporangiophore) (Ramsbottom, 1953, rev. 1989).

Another gall-causing rust fungus was illustrated in Ray’s Synopsis Methodica Stirpium Britannicum (third edition, 1724), prepared and illustrated after Ray’s death by the German botanist Dillenius (1687–1747), although it was not then recognised as a fungus. In Fig. 262, Tab. III of the Synopsis is reproduced, with the mysterious ‘Conjurer of Chalgrave’s Fern’ illustrated. This is not a fern but a leaf of wood anemone Anemone nemorosa distorted by the rust Tranzchelia anemones forming swellings scattered over its lower surface. The telia of this species cause galls, mistaken for the sori of the fern, specifically on A. nemorosa.

NINETEENTH-CENTURY ADVANCES

While the century as a whole saw immense scientific advances in many spheres of biology, the first half of the century was a slack period for cecidology, and knowledge regressed. Connold (1901), who was often vague about sources, quoted suppositions such as ‘…ova were deposited by the parent insect in the ground, and from thence drawn up with the sap’. He also quoted from Insect Architecture (1857) by J. Rennie, whose knowledge of the formation of galls was less than that of Malpighi, 170 years before:

…the parent fly ejects into the cavity [made by her ovipositor] a drop of corroding liquor, and immediately lays an egg or more there; the circulation of the sap being thus interrupted, and thrown, by the poison, into a fermentation that burns the contiguous parts, and changes the natural colour. The sap, turned from its proper channel, extravasates and flows round the eggs, while its surface is dried by the external air, and hardens into a vaulted form.

Until the mid-nineteenth century, it was generally accepted that each adult gall insect species produced a specific poison which resulted in a specific gall structure. Darwin (quoted in Cameron, 1882–93) says:

As the poisonous secretion of insects belonging to various orders has the special power of affecting the growth of various plants; as a slight difference in the nature of the poison suffices to produce widely different results; and, lastly, as we know that the chemical compounds secreted by plants are eminently liable to be modified by changed conditions of life, we may believe it possible that various parts of a plant might be modified through the agency of its own altered conditions.

In 1803, a gall on a filamentous alga was discovered by Vaucher; the alga was later named after him as Vaucheria (Spooner, in Williams, 1994). The alga and the gall had been illustrated earlier by Vahl in Flora Danica (1787), using Danish material of Ulva (now Vaucheria) dilatata (see Fig. 13, Chapter 3). Vahl considered the growth to be a normal part of the alga, and this led to two varieties of V. dilatata being described, one based on galled and one on the ungalled filaments (Roth 1800, 1806, quoted in Spooner, in Williams, 1994). Vaucher realised that the galls were caused by an ‘insect’ Cyclops lupula and, in 1807, similar galls from Britain were illustrated (by Smith; see Spooner, in Williams, 1994). In 1834, the causer was recognised as a rotifer and was described by Ehrenberg as a new species, Notommata (now Proales) werneckii. This, however, was overlooked during the next 40 years, which led to muddles over its identification.

From the 1850s onwards, many new discoveries were made in cecidology, some of them reflecting the theoretical concerns of the time, including the theory of evolution by natural selection and its consequences. Towards the end of the nineteenth century and as the lifecycles of cynipid wasps were investigated in detail, it became clear that many species of oak gall wasp had two generations a year, one producing males and females and the other females only, with each causing a different type of gall, often on a different part of the host plant. Often this led to confusions over nomenclature, with a single species being given two distinct names. As the relationships were worked out, pairs of species originally thought to be distinct were combined as one. This was done via a series of careful experiments, mainly by Adler in Germany (Adler & Straton, 1894). Sorting out the lifecycles of oak gall wasps continues today, particularly for species in eastern Europe, Asia and North America, where the fauna of oaks is less well known than in western Europe.

The phenomenon of alternating sexual and asexual stages in animals was first discovered in 1819 by De Chamisso for ‘Salpae’ (coelenterates, Cnidaria, for example, jellyfish) in which polyps and medusae alternate. Throughout the century, evidence accumulated that showed that many types of animals required two or more generations to complete their lifecycles (Adler & Straton, 1894, summarised the information). Normally, one generation was sexual and the other(s) asexual involving parthenogenesis, with the generations differing in form and living under different conditions. Alternation of generations in oak cynipids was first suspected in the middle of the century when, in 1840, parthenogenesis was demonstrated by Theodor von Hartig; in breeding experiments, he showed that no males existed in some generations. Leuckart in 1858, followed by Lichtenstein in 1881, regarded the parthenogenetic generation of the cynipids as the larvae of the sexual generation. Spangle galls were given as an example: individuals of Neuroterus lenticularis (the agamous form of N. quercusbaccarum) were the larval forms of Spathegaster baccarum (the sexual form of N. quercusbaccarum) even though they had wings and ovipositors, and laid eggs. In 1861, Osten-Sacken, who studied oak galls in North America, proposed that agamous species were actually sexual with the males developing in different galls that had yet to be recognised. In 1864 Walsh, also an American, questioned whether parthenogenesis did, in fact, exist in the cynipids. From galls of the same type, Walsh observed males and females emerging from some and females only, of a quite different form, from others, and he suggested that both types of female could mate with the males. But there was no evidence to support either of these views and they had to be abandoned.

The first clear description of alternating sexual and asexual stages in oak cynipids was published in North America in 1873 by Bassett on Quercus bicolor (and reported by Adler, see Adler & Straton, 1894; Straton translated Adler’s work into English):

Bassett had repeatedly found enormous numbers of a gall belonging to a species of Cynips, on a small oak (Quercus bicolor). These galls appeared with the leaves, causing shapeless swellings of the petiole and midrib; they contained a large number of larvae, and in June there emerged flies of each sex in nearly equal proportion. Late in the summer there was formed on the points of the young shoots of the same oak tree, a differently shaped gall in which the flies passed the winter. This latter species occurred in the female sex only. It closely resembled the former species, but was somewhat larger. From this observation Bassett arrived at the conclusion that each species of Cynips which is found occurring exclusively in the female sex is succeeded by another generation which is bisexual…Bassett concluded by saying that it would not surprise him if it were proved that every species of the genus Cynips had each year two generations differing in the manner indicated by him.

At about the same time (1875), unaware then of Bassett’s work, Adler himself was investigating the lifecycles of oak cynipids in Germany, and by the end of the century he had largely solved the problems of their lifecycles. Given that different generations form galls of very different character, and often on different parts of the plant, proving that both are the same species was no small task. Adler investigated the lifecycles with experiments under controlled conditions and his work is a fine example of meticulous observation and sound scientific method. He emphasised the importance of rearing adult flies from individual isolated galls so that there was no doubt which species he was dealing with:

A lucky chance led me to select the species of Neuroterus for my first experiments: these galls are easily collected in large numbers, and there is little difficulty in rearing the flies. In every case I made a point of breeding the flies from the galls, so that I might be absolutely certain of the species. The flies emerge in March and April from Neuroterus galls which had matured in autumn, and they proceed at once to lay eggs in the buds of the oak. It struck me as remarkable that although the egg was laid so early, the gall did not develop until July: and it was the strangeness of this circumstance, added to a desire to investigate the method of gall formation, that led me to undertake my first direct experiments in breeding. These afforded me the surprising result that from the eggs laid by Neuroterus there appeared a totally different generation, one so wholly unlike its parent that it had been described hitherto as of another genus (Spathegaster) [Adler’s emphasis]. This fact was published by me in 1877, and what Bassett had only thrown out as a conjecture in 1873 was now proved and demonstrated in one species at least.

Adler then proceeded to investigate the lifecycles of about 40 oak cynipids then recorded for northern Germany. His aim was to prove that many or all species had two alternating generations, one sexual and one agamous. His investigations involved careful breeding experiments which ensured that the female instigating each type of gall could be identified without any doubt:

In order that the results which I obtained might be unquestionable it was necessary to select a method which provided against every possible source of error. This could only be satisfactorily accomplished by watching the development of the gall of each species, from the time when the egg was laid until the gall reached maturity. Unfortunately, however, this peculiar difficulty exists, that the most important phase of development, viz. that during which the eggs of the fly are buried in the bud or tissues of the oak, must unavoidably be hidden from direct observation. Indirect observation alone is available, for any actual examination of the eggs when laid must necessarily end in their destruction. Thus if a gallfly lays its egg in a bud, one can predict with certainty what gall will be produced, so long as care is taken that the same bud is not pricked either before or afterwards by another fly. Breeding experiments must therefore be so arranged as to enable each species to be isolated and watched while actually depositing the egg.

These are ideal instructions for anyone investigating lifecycles and gall formation. Adler described his methods for rearing the species that gall leaf buds or bark, for which he used four-to six-year-old saplings of Quercus sessiflora (now Q. petraea) planted in pots and covered to prevent insects leaving or entering. For species that gall the catkins, it was necessary to use adult trees growing outside; for these, he sleeved parts of branches in muslin bags to isolate them.

Adler considered the advantages of parthenogenesis to the insect in order to explain how alternating generations might have evolved. The production of parthenogenetic eggs in arthropods is widespread and appears at particular seasons whenever such reproduction is more efficient, perhaps, for example, to take advantage of a temporary glut of food. But this explanation does not apply to most oak cynipids in which the parthenogenetic generation develops in the autumn and overwinters, and adults emerge and lay eggs in the spring. Adler suggested that mating would be less successful in early spring when the weather is likely to be colder and wetter than later in the year, and stressed that parthenogenesis allows an increase in numbers despite the adverse conditions. He considered that sexual reproduction was the primitive type, with parthenogenesis evolving later and perhaps becoming dominant. In Cynips (now Andricus) kollari, he thought that the sexual stage had been lost altogether (its sexual generation had not then been discovered). Today, in Britain and western Europe, species originally thought to be distinct have been linked, so that their lifecycles are complete, but there remain a few oak cynipids in which a sexual generation is unknown and others with bisexual galls only.

Adler, in the last quarter of the nineteenth century, was using evolutionary arguments at a time when Darwinian ideas were firmly entrenched but were not universally accepted. Two distinguished biologists, G. J. Romanes and St George Mivart, both considered that galls constituted an obstacle to the theory of evolution by natural selection. Mivart’s argument (1889, quoted in Adler & Straton, 1894) ran:

Now surely it is too much to ask us to believe that the germ-plasm of the plant, in the first instance, before even, say, a single cynips had visited it, had in the complex collocation of its molecules, an arrangement such as would compel the plant which was to grow from it, to grow those cells and form a gall…It would be very interesting to know how natural selection could have caused this plant to perform actions which, if not self-sacrificing (and there must be some expenditure of energy), are at least so disinterested.

In other words, the plant is providing benefit for the insect without gaining any advantage in return. Romanes took the same line, but he stressed that it was ‘the one and only case’ to show such apparent altruism (Cockerell, 1890). Adler tackled these arguments thus:

It is next of interest to inquire how the various structures of the gall came to be evolved. It may be taken as perfectly certain that the tree does not form them in a disinterested manner for the sake of the Cynips. Darwin says: ‘If it could be proved that any part of the structure of any one species had been formed for the exclusive good of another species, it would annihilate my theory, for such could not have been produced through natural selection’. So far as galls are concerned, Darwin’s theory is perfectly safe. The ‘excitatory emanations’, as Professor Romanes aptly calls them, which lead to gall-growth, can only have arisen by gradual and increasing improvements in the initial stages of their formation, acting through natural selection, over an unlimited period of time, and through numerous consecutive species.

Adler classified oak galls into five groups of increasing structural complexity that illustrate an evolutionary sequence (the names of most of these insects have changed (see Appendix A; the structure of some of the galls is described in Chapter 9):

1. Simple galls, consisting of nutritive tissue enclosed in thin-walled parenchyma with vascular bundles: Neuroterus ostreus, Spathegaster albipes, S. baccarum, S. Aprilinus

2. Galls similar to these, but having the nutritive tissue first enclosed in sclerenchyma, which forms an ‘inner gall’: Neuroterus lenticularis, N. laeviusculus, N. numismatis, N. fumipennis, Aphilothrix Sieboldi, A. autumnalis, A. radicis, A. globuli, Andricus curvator, Biorhiza renum, B. aptera

3. Galls possessing an inner gall like the last, but having it surrounded by thick-walled parenchyma: Dryophanta longiventris, D. divisa

4. Galls with the inner gall enclosed in a spongy layer of branched parenchyma with wide intercellular spaces, and having the surface covered with differentiated epidermis: Unilocular: Dryophanta scutellaris. Multilocular: Teras terminalis

5. Galls which have the inner gall enclosed in thick-walled parenchyma, and then in spongy tissue; and which have a differentiated epidermis: Cynips Kollari.

He suggested that this sequence shows gradual improvements in providing favourable conditions for the larvae and protection against parasites, with each change arising by natural selection. Thus, there is nothing altruistic in the formation of the gall. The insect is exploiting the normal response of the plant to an injury by converting the wound tissue so formed into nutritive and protective tissue. Adler summarised existing theories of gall formation from Malpighi to Darwin:

Darwin and all writers before him held that force calling out gall formation was due to a chemical secretion injected by the gall mother…Darwin speaks of galls as produced ‘by a minute atom of the poison of a gall-insect’, and compares them to the specific local processes of zymotic [infectious] diseases.

He continued that ‘the most reasonable, if the only reasonable theory, is that each insect infects or inoculates the leaf or other structure of the chosen plant with a poison peculiar to itself’. This is generally accepted today. Adler showed in addition that the larva and particularly its salivary secretion is necessary for the gall to develop. The debate on why the elaborate structure of galls develops continues today (see Chapter 11).

Adler was concerned only with cynipid galls. Cameron (1882–93) stressed that gall insects were so diverse that gall formation could not be explained by just one process. Cameron noted that in sawflies the gall forms before the egg hatches, while in cynipids the larva must hatch and start to feed before the gall begins to form (although the fluid injected with the egg does cause a small swelling). By the end of the century, therefore, the stage had been set for investigating mechanisms of gall formation by recognising that a single answer might not apply to all cases.

Two centuries earlier, Malpighi and Derham had realised that cynipid galls contained animals in addition to the gall causer. The relationships between them remained obscure until the mid-nineteenth century and the work of von Hartig, Über die Familien der Gallwespen (1840–43). He arranged gall inhabitants into three classes (described in Adler & Straton, 1894):

The gall makers, which he compared to householders.

Inquilines, guest-flies, cuckoo-flies or lodgers, who take up their quarters uninvited, live on the food provided, but do not aim to kill the gall maker.

Parasites, who deposit their eggs on the larvae of the gall maker or lodgers with the object of destroying them, who are therefore murderers.

Adler observed that, although parasites were not able to initiate galls, some could prolong gall growth after the death of the gall causer, and so modify the gall.

In the late nineteenth century, more information about particular groups of gall-causing insects accumulated, with the appearance of large monographs: Mayr’s Die Mitteleuropäischen Eichengallen (1872–82) (parts of which were translated into English in the Entomologist, 1870–71); Buckton’s British Aphides (1876–83) in four volumes, which included a few aphid galls; and Cameron’s Monograph of British Phytophagous Hymenoptera (1882–93), with Volume IV devoted to cynipids. Other gall inducers were also recognised, including fungi, nematodes and eriophyoid mites; and galls were recognised from more plants and parts of plants, including the roots (Connold, 1901). The first half of the twentieth century was notable for the publication of major catalogues and keys for the identification of galls (e.g. Houard, 1908–13; Rübsaamen & Hedicke, 1925; Felt, 1940; Barnes, 1948–56; Docters van Leeuwen, 1957, see Roskam, 2009; Buhr, 1964–5).

Although galls have been known for 2,500 years, their great variety did not begin to be appreciated until little more than 100 years ago. In the second half of the twentieth century, structure and development were investigated and described for more and more galls. But the explanation of how they develop was not much clearer than it was in the seventeenth century, and remains obscure. Until the seventeenth century, galls were of interest for their useful properties, as herbal remedies for a wide range of illnesses and for ink-making, dyeing and tanning. Some of these uses continued until the mid-twentieth century, although for the previous 100 years or so, galls were also studied for their own sake. Their main contribution to the history of biology was the part they played in the controversy about spontaneous generation, which ran throughout the seventeenth and eighteenth centuries. Most of the history of the study of galls involves insect galls only, particularly cynipid galls, and these are still the ones that are best understood.

Endnotes

In the past, the family Psyllidae included all members of this group and ‘psyllid’ was the acceptable common name. Now, the bugs are classified into several related families, united into the superfamily Psylloidea, so the common name has become ‘psylloids’ (see Box 4.4). The same rule applies to the gall mites, Eriophyoidea (see Box 4.1).

Types of gall are described in different chapters, as follows (see Fig. 38): Erinea on leaf, without (1–5) and with (6–8) associated swelling of the leaf–Chapter 4; Fold galls (11–12)–Chapter 4; Rolls of leaf margin (13–15)–Chapter 4; Blister galls (9–10)–Chapter 5; Pouch galls in leaf (16–24) and stem (25)–Chapter 5; Big bud gall (26)–Chapter 6; Virescence (27)–Chapter 2.

Except, apparently, in New Zealand where the scale has two generations a year; the spring generation has females only and males are produced in the autumn (Parr, 1940).

The terms, bottom-up and top-down, are based on the idea of a simple food chain: plant-herbivore-predator. The numbers of the herbivore, i.e. the gall causer, may be affected by the host plant and factors that affect it (climate, soil and other environmental conditions), i.e. bottom-up effects. Or, the gall causer may be affected by its enemies (its parasitoids, predators and inquilines), i.e. top-down effects.

Eurytoma brunniventris is part of the Eurytoma rosae–complex, a group of closely related species that parasitise gall wasps in rose and oak galls. Recent work suggests that E. brunniventris specialises on Andricus and Synergus species in the centre of large galls, while the species attacking Andricus species in small galls and Synergus species in outer chambers of large galls is Eurytoma rosae, the same species that parasitises Diplolepis rosae and its inquiline in bedeguar galls (Alex Aebi and Graham Stone, pers. com.).

Different parts of large plants are often found as separate fossils and have been given different names: these are ‘form-genera’ or ‘form-species’. Rhachis fossils of Psaronius species belong to the form-genus Stipitopteris while foliage fossils are known as form-genus Pecopteris; and other names are used for fossils of the spores and sporangia. Reconstruction of the whole-plant tree-fern Psaronius chasei combines species of these form-genera (Labandeira & Phillips, 2002).

The phenotype of an organism describes what it looks like (its structure) and how it functions and behaves. Its genotype, or genome, is its genetic make-up, dictated by the genes carried on the chromosomes of the cell and inheritable by its offspring, and the genes largely govern the phenotype. So, the genome of the gall-causing insect controls the appearance and development of the gall even though it (the gall) is made entirely of plant tissues (and is, therefore, the product of the plant’s genome). Thus, the gall is the extended phenotype of the insect, superimposed on the plant.

The Earth’s crust is divided into a number of plates on which the continents sit. Over geological time, the plates move in relation to each other, a movement caused by the formation of new crust along some plate margins and destruction of crust along other margins. The direction and speed of movement is different for each plate and has changed during the course of evolution.

In the USA, Q. laevis is known as turkey oak with a small ‘t’, because of the shape of its leaf, like a turkey’s foot.

‘Astringent’ (according to the Shorter Oxford English Dictionary, 1983): having power to draw together or contract the organic tissues; binding, constrictive, causing constipation; also, blood is staunched by astringents.

These volumes include useful papers by many authors, relevant to Chapters 1–14; the papers are not quoted individually. Note that titles relevant to particular chapters may be found under the General heading.

Glossary

(bold = defined elsewhere in the Glossary)

achene: dry fruit containing a single seed, e.g. in Asteraceae (daisies, thistles, dandelions, etc.).

agamic: (see asexual).

alata (plural alatae): winged aphid (c.f. aptera).

alkaloids: organic compounds produced by many plants that are toxic to many herbivores.

alleles: different forms of a gene.

allopatric speciation: the formation of two species from an ancestral species whose range has been divided geographically by a barrier, so that the populations on each side evolve independently and eventually are unable to interbreed (c.f. sympatric speciation).

allozymes: enzymes derived from alleles; the enzymes exist in more than one form but all forms function in the same way (see Box 12.1, Chapter 12).

amber: fossilised resin produced by conifers.

ambrosia gall: cavity of the gall of a gall midge lined with fungal hyphae on which the larva may feed; the fungus is necessary for the development of the gall as well as the plant and the larva.

amorphous: without distinct shape or structure.

analogous: structures that may have a similar function but are derived independently (c.f. homologous; see Box 11.1, Chapter 11).

anamorph: asexual fruitbody of a fungus; species in which no sexual spores are known are anamorphic.

androphore: an asexual female of a gall wasp that lays only haploid male eggs (see Box 9.1, Chapter 9), so reproducing by arrhenotokous parthenogenesis (c.f. gynephore).

angiosperm: flowering plant.

anholocycle: lifecycle of an aphid species in which the sexual stage is lost (see Box. 4.5, Chapter 4).

anther: male part of flower, produces pollen.

apical bud: bud at apex of shoot, the terminal bud (c.f. axillary bud).

aptera (plural apterae): wingless aphid (c.f. alata).

arrhenotoky: parthenogenetic reproduction in which males develop from unfertilised haploid eggs and females develop from fertilised diploid eggs (see also haplodiploidy; c.f. thelytoky).

artichoke gall: leafy gall formed of overlapping leaves, gall length longer than width, originating from a bud; due to failure of internodes to lengthen (c.f. rosette gall).

asexual: without sex.

autoecious: a plant parasite that completes its development on one host plant.

auxin: a plant hormone promoting or regulating growth.

axil: the junction between leaf and stem.

axillary bud: bud in axil of a leaf, a lateral bud.

big bud: swollen bud gall, containing small distorted leaves, caused by gall mites (Eriophyoidea).

biological control (biocontrol): the control of a pest or weed by an introduced enemy of the pest (rather than by chemical pesticides).

biotroph: a parasite that gains nutrients from a living host.

bivoltine: a lifecycle with two generations each year (c.f. multivoltine, univoltine).

blister gall: rounded thin-walled swelling with a cavity, raised on one or both sides of a leaf; or a bulge raised on one side of a leaf, the concave surface lined with fungal spores.

boreal forests: coniferous forests of cold temperate climates in the northern hemisphere.

bottleneck: a new population that arises from a small number of founders with genes that are a small subset of a larger parental gene pool has passed through a genetic bottleneck; the genetic variation of the new population is less than in the old (see Box 12.1, Chapter 12).

bottom-up effects: based on the idea of a simple food chain: plant herbivore predator; numbers of the herbivore, i.e. the gall causer, are affected by the host plant and factors that affect it (climate, soil and other environmental conditions) (c.f. top-down effects; see Chapter 10).

bracts: scale-like modified leaves protecting young leaves in bud; or enclosing flower head, especially in Asteraceae (daisy, thistle family).

brooming: (see witches’ broom).

callus: wound tissue produced by plant after damage by a generalised herbivore or a simple gall causer.

cambium: layer of young unspecialised cells forming a meristem; cells divide rapidly, responsible for increase in girth of stems and roots.

canker: a solid growth or swelling on a branch or trunk of a tree, caused by bacteria or fungi (and so a type of gall); or due to wounding (so may not be a gall).

caterpillar: larva of a butterfly, moth or sawfly; typically with prolegs (false legs) on the abdomen.

cell vacuole: fluid-filled sac filling most of a fully developed plant cell.

cellulose: substance that forms plant cell walls.

chelicerae: the jaws of mites and spiders.

chitin: material that forms the exoskeleton of arthropods.

chloranthy: transformation of petals, sepals and other flower parts into leaf-like structures (phyllanthy has the same meaning).

chloroplast: organelle in plant cells that contains chlorophyll; the site of photosynthesis.

chromosome: long molecule of DNA often wrapped in proteins; carries the genes of the cell.

clade: a group of related species in a cladogram.

cladogram: a family tree diagram that illustrates the evolutionary relationships within a group using cladistics, a rigorous method for determining the status of characters (see Box 11.1, Chapter 11).

class (see phylum).

cleptoparasite: species that fight and displace gall causers and take over their gall; common in thrips (see Box 4.3, Chapter 4).

cline: the linear pattern of spread of a species from one area to another; a species that spreads away from its endemic range loses genetic diversity along the way (see Box 12.1).

clone: juveniles that are genetically identical to their mother, produced parthenogenetically.

coevolution: synchronous development of two species where each drives the evolution of the other, e.g. in figs and fig wasps (see Chapter 7); cospeciation is similar.

collenchyma: a type of supporting tissue in plants.

companion cells: cells that drive the functioning of phloem vessels.

conserved evolution: all galls in a clade have the same character(s) and so have a common origin.

convergent evolution: two or more species with different origins evolve the same character independently, a similarity that does not reflect common ancestry (see Box 11.1, Chapter 11).

coppicing: cutting trees or shrubs to ground level to encourage growth of many young stems.

crown gall: a growth that develops in the crown of a plant, at the junction of the stem and root or elsewhere on a tree or shrub; caused by the bacterium Agrobacterium tumefaciens.

cultivar: a variety of plant produced and maintained by cultivation.

cuticle: the waxy layer on the outside of the epidermis of a stem or leaf.

cytokinin: a plant hormone that stimulates cell division.

cytoplasm: the substance of the cell excluding the nucleus and the cell wall.

dehiscence: the shedding of plant parts or of galls from a leaf.

detritivore: an animal that feeds on detritus (or on the microorganisms that grow on it).

dicot, dicotyledon: a plant that germinates with two seed leaves (cotyledons); normal leaves with a midrib and network of veins (c.f. monocotyledon).

differentiate, differentiation: cells specialised to perform particular functions (c.f. unspecialised cells of the meristem).

dikaryotic: each cell contains two haploid nuclei (see Box 2.4, Chapter 2).

dioecious: plant in which pollen and seeds are produced by separate individuals (c.f. monoecious).

diploid: with two sets of chromosomes in the cell (c.f. haploid).

distal: part of a structure furthest from the base or point of attachment (c.f. proximal).

DNA, deoxyribonucleic acid: the genetic material of an organism (see Box 11.1, Chapter 11).

dormant bud: undeveloped bud on trunk, branch or in a leaf axil.

ectoparasitoid: a parasitoid that lays its eggs on the outside of the host or nearby in the gall chamber; the larva feeds on the host from the outside.

endemic: a species that has evolved in a particular geographical region, found nowhere else (c.f. native).

endoparasitoid: a parasitoid that lays its eggs inside the host and whose larva feeds within it.

endophytic: living permanently inside a host plant.

endopterygote: an insect with a pupa in its lifecycle.

endosperm: the food store of a seed, used by the germinating seedling.

epidermis: outermost layer of cells in a plant.

erineum (plural erinea): a simple gall consisting of a patch of glandular hairs, caused by gall mites.

ergot: black or purple gall replacing grain of grasses (Poaceae) and some sedges (Cyperaceae), caused by the fungus Claviceps.

eukaryote: an organism whose cells include nuclei bounded by membranes (c.f. prokaryote).

family: (see phylum).

fasciation: abnormal multiplication of plant parts, often with thickening; usually due to physiological malformation, occasionally due to bacteria or viruses (when the growth would be classed as a gall).

fecundity: the number of eggs or offspring that a female can produce.

flagellum (plural flagella): long whip-like appendage of a cell that beats to propel the cell forwards.

floret: a small individual flower, a number of which make up a flower head or inflorescence, e.g. in Asteraceae (daisy family) and Poaceae (grasses).

frass: dry granular faeces produced by caterpillars, beetle larvae, etc.

frond: the leaf of a fern.

fruitbody: part of a fungus that produces spores.

fundatrix (plural fundatrices): the founding female of all the parthenogenetic generations that develop in a gall, e.g. of aphids, thrips and gall mites.

fynbos: a shrubby heathland vegetation, with a rich variety of woody plants less than 1 m tall; soils are thin, dry and nutrient-poor and frequent fires are characteristic; found in the southwest Cape of South Africa.

gallicolae: individuals that develop inside a gall, used particularly for gall aphids (see Box 4.5, Chapter 4).

gametes: sex cells, e.g. eggs and sperm.

genome, genotype: the genetic make-up of an organism dictated by the genes carried on the chromosomes of the cell and inheritable by the offspring (c.f. phenotype).

genus: (see phylum).

glacial refuge: areas (e.g. in southern Europe) isolated by ice or cold climate where species survived during northern glaciations.

Gondwana: the southern supercontinent that existed 150 million years ago, incorporating South America, Africa, India, Australia, New Zealand and Antarctica; it began to break up into these present-day continents in the early Cretaceous.

gregarious parasitoid: several parasitic individuals live inside the host.

gynandrophore: an asexual female of a gall wasp able to produce both male and female offspring (see Box 9.1, Chapter 9) (c.f. androphore, gynephore).

gynephore: an asexual female of a gall wasp that lays only diploid female eggs (see Box 9.1, Chapter 9), so reproducing by thelytokous parthenogenesis (c.f. androphore).

haplodiploidy: females are diploid with two sets of chromosomes in each cell and develop from fertilised eggs; males are haploid developing from unfertilised eggs containing only one set of chromosomes from the mother (see arrhenotoky).

haploid: with one set of chromosomes in the cell (c.f. diploid).

haustorium (plural haustoria): (i) specialised hypha of a parasitic fungus that penetrates tissues of host; (ii) swelling of gall-causing plant where attached to its host; function in both to absorb nutrients from the host.

herbaceous: a non-woody plant.

heteroecious: a herbivore or parasite that requires two host species in which to complete its development.

heterogony: a sexual generation alternates with one or more asexual generations each year (also known as cyclical parthenogenesis and heterogyny; see Boxes 4.5 and 9.1, Chapter 4, Chapter 9).

Holarctic: the geographic region that includes the Nearctic and the Palaearctic (see Fig. 12.1, Chapter 12).

holocycle: the full lifecyle involving two host plants, e.g. of aphids (see Box 4.5, Chapter 4).

homologous: structures that have a common origin although their functions may be different (c.f. analogous; see Box 11.1, Chapter 11).

honeydew: sweet liquid produced from anus of aphids, psylloids and scale insects, often collected by ants (see also Box 2.4, Chapter 2).

host: (i) a plant that is parasitised by a gall causer; (ii) an animal that is parasitised by another animal (a parasitoid or a parasite).

host-switching: the ability of a species specialising on one host species to survive on another host; also organ switching.

hyperparasitoid: a parasitoid of a parasitoid (see also Box 10.1, Chapter 10).

hyperplasy: abnormal growth caused by multiplication of plant cells.

hypertrophy: abnormal growth caused by enlargement of plant cells.

hypha (plural hyphae): thread-like structures that make up the body (the mycelium) of a fungus.

idiobiont: a parasitoid that kills or paralyses its host when ovipositing so that it does not develop further (c.f. koinobiont; see Box 10.1).

inflorescence: flower head composed of two or more (often many) florets, e.g. in Asteraceae (daisy family), Apiaceae (umbellifers), Moraceae (figs), Poaceae (grasses).

inquiline: a mite, insect or other organism that lives in and feeds on the tissues of a gall caused by another species, and which may distort the gall; if it kills or is responsible for the death of the host, it is a lethal inquiline.

insectivore: an animal (and a few plants) that feeds on insects.

instar: stage of development between two moults of an insect or mite.

intercellular: the space between cells.

interglacial: warmer period between two glaciations.

internode: portion of plant stem between two joints (or nodes).

invaginate: turn inside-out.

kingdom (see phylum).

kleptoparasite: (see cleptoparasite).

koinobiont: a parasitoid that does not kill its host at oviposition allowing the host larva to grow until it is fully fed; then the parasitoid kills and feeds on it (c.f. idiobiont; see Box 10.1).

labium: mouthparts of insects (next to the maxillae).

lammas leaves: second flush of young leaves produced by some trees in summer.

larva (plural larvae): juvenile stage of an insect, often specifically used for those with a pupa in the lifecycle, e.g. moths, beetles, flies, wasps (c.f. nymph).

leaf initials: tiny structures formed by the meristem that develop into leaves.

leaflet: individual section of a compound leaf attached to the rhachis, e.g. in ash (Fraxinus), roses (Rosa), peas (Pisum), vetches (Vicia).

leaf mine: linear or blotch pattern caused by a larva feeding in the thickness of a leaf.

lenticels: pores in the surface of stems and tree branches and trunks that allow gases to enter and exit.

lerps: shelters constructed by a psylloid nymph from hardened honeydew.

lethal inquiline: (see inquiline).

lignin: woody tissue produced by plants.

mandibles: jaws of insects (and other animals).

maxillae: mouthparts of insects (next to the mandibles).

meiosis: division of a diploid cell to produce haploid daughter cells.

meristem: part of plant containing young unspecialised cells, able to divide rapidly and to develop into a variety of tissues; found in buds on shoots and at tip of roots, in cambium, and (in grasses) at the base of stems and leaf sheaths.

mesic: equable conditions; temperatures, rainfall, soil fertility, etc. are not extreme.

mesophyll: the green tissue of a leaf, divided into upper palisade and lower spongy layers.

microorganism: bacteria, fungi and other organisms that cannot be seen with the naked eye.

microsatellites: regions of a chromosome where the DNA is repetitive; the number of repeat units in a microsatellite varies between individuals (see Box 12.1, Chapter 12).

middle lamella: membranous layer between cells that binds them together.

midrib: the main vein of a leaf.

mitosis: division of a cell (haploid or diploid) to produce daughter cells with the same number of chromosomes.

molecular taxonomy: the use of genetic material DNA to compare the similarities and differences between organisms and thus to classify them (see Box 11.1, Chapter 11).

monocot, monocotyledon: a plant that germinates with one seed leaf (cotyledon); normal leaves usually with parallel veins (c.f. dicotyledon).

monoecious: plant in which pollen and seeds are produced by the same individual (c.f. dioecious).

monophagous: feeding on one species of plant (c.f. oligophagous, polyphagous).

monophyletic: a group of species with a unique common ancestor, which includes all descendant species of that ancestor (see Box 11.1, Chapter 11).

morph: a distinctive form of individual in a polymorphic population.

multicellular: organism with many cells.

multilocular: a gall with several chambers (c.f. unilocular).

multinucleate: cells with many nuclei.

multivoltine: a lifecycle with many generations each year (c.f. bivoltine, univoltine).

mutation: an alteration in the structure of DNA, which can be inherited by the offspring.

mutualism: a symbiosis in which two organisms live together and both benefit.

mycangia: special pockets at the tip of the ovipositor or abdomen of gall midges that carry spores of ambrosia fungi.

mycelium: the body of a fungus, made up of hyphae.

mycorrhiza: an intimate interaction between a fungus and plant root benefiting both species; an example of a mutualism.

native: found naturally in an area, i.e. not introduced (c.f. endemic).

Nearctic: the geographic region that covers North America (c.f. Palaearctic, Holarctic (see Fig. 12.1, Chapter 12).

necrosis: blackened patches of dead cells in plants.

nectaries: structures in flowers that produce nectar.

Neotropics: most of Central and South America (see Fig. 12.1, Chapter 12).

node: a joint in a plant stem (c.f. internode).

nodule: small solid gall without internal chambers, caused by bacteria on roots of legumes and other plants; may be woody.

nucleus: organelle in a eukaryote cell that contains the chromosomes.

nutrient sink: the movement of nutrients to the gall and its larva drawn from elsewhere in the plant.

nutritive tissue: a layer or patches of cells rich in nutrients (glucose, sucrose, amino acids, etc.) and enzymes required by the galling larva; the nutritive tissue and feeding larva act as a nutrient sink.

nymph: juvenile stage of an insect, used for those without a pupa in the lifecycle, e.g. aphids, psylloids, scale insects (c.f. larva).

oligophagous: feeding on a few related species of plant (c.f. monophagous, polyphagous).

omnipotent: the ability of cells to develop into any type of tissue, as in meristems.

order: (see phylum).

ostiole: the opening at the top of a fig syconium.

oviparous: egg laying (c.f. ovoviviparous, viviparous).

ovipositor: the egg-laying tube of female insects, prominent in many parasitic wasps.

ovoviviparous: the egg hatches inside the female’s body so that a larva is laid (c.f. oviparous).

Palaearctic: the geographic region that extends from western Europe to eastern Asia (c.f. Nearctic, Holarctic (see Fig. 12.1, Chapter 12).

palaeotropical: the tropical zones of Africa and Asia (see Fig. 12.1, Chapter 12).

palisade mesophyll: (see mesophyll).

pappus: thistledown, the hairs attached to the achene of thistle florets.

parasitoid: an insect parasite whose larva feeds on a single host insect and kills it.

parenchyma: tissue consisting of large unspecialised cells found in all parts of plants.

parsimonious: interpretation of evolutionary change that involves the smallest number of independent changes in the state of a character; the simplest interpretation of a number of possibilities (see Box 11.1, Chapter 11).

parthenogenesis: asexual reproduction by females, without mating.

pathogen: a parasitic microorganism, a virus, bacterium or fungus.

pedipalps: mouthparts next to the chelicerae of mites and spiders.

peduncle: flower stalk (c.f. petiole).

permeable: a membrane or skin that is not waterproof.

petiole: leaf stalk (c.f. peduncle).

phenology: the timing of plant development, when it produces its leaves, flowers, fruits, etc.

phenotype: describes the structure of an organism and its function (c.f. genotype).

pheromone: a chemical produced by one individual that affects the behaviour of another of the same species.

phloem: vessels in vascular tissue that transport sugars from leaves to the rest of the plant (c.f. xylem).

photosynthesis: the process whereby green plants synthesise glucose from carbon dioxide and water.

phyllanthy: transformation of petals, sepals and other flower parts into leaf-like structures (chloranthy has the same meaning).

phyllodes: leafy extensions of petioles in plants that have lost their leaf blades (e.g. in acacias).

phyllody: (see phyllanthy).

phylogenetic: a natural classification of organisms, explained in Box 11.1, Chapter 11.

phylum (plural phyla): a rank within the hierarchy of classification; organisms are classified into a series of groups (of decreasing size) that indicate their affinities (e.g. kingdom, phylum, class, order, family, genus and species).

phytomycetophagy: feeding on fungi and on plants.

phytophagy: plant-feeding.

pinnule: a segment of the frond of a fern.

pit gall: a small depression usually in a stem, often with a raised rim.

pith: tissue in the core of a stem.

plasmid: small ring of DNA in a bacterial cell separate from its chromosome (see Box 2.2, Chapter 2).

plasmodium: a microorganism (e.g. a slime mould) containing many nuclei, not divided into separate cells.

plate tectonics: movement over geological time of the plates on which the continents sit, caused by the formation and destruction of crust along the plate margins.

pollen tube: a tube that grows from a pollen grain on the stigma, down the style to the ovary of the plant prior to fertilisation.

pollination: the transfer of pollen from the anthers to the stigma of flowers.

polymorphism, polymorphic: a species with several forms (or morphs).

polyphagous: feeding on many species of plant (c.f. monophagous, oligophagous).

polyphyletic: a group of species that has more than one ancestral species (c.f. monophyletic, see Box. 11.1, Chapter 11).

polyploid: cells with several sets of chromosomes (c.f. haploid, diploid).

pouch gall: rounded or elongate finger-shaped gall raised on surface of leaf with an opening to outside; or a sac-like gall formed from part or all of leaf.

prepupa: inactive stage of a full-grown caterpillar prior to pupation.

procecidium: a small swelling in a plant caused by the female during egg-laying.

prokaryote: an organism with cells without nuclei bounded by membranes, i.e. all bacteria (c.f. eukaryote).

propagule: reproductive structure, e.g. seed, spore, cutting.

protozoa: a disparate group of eukaryotic microorganisms (c.f. prokaryotes).

proximal: part of plant nearer to the stem or point of attachment (c.f. distal).

ptilinum: a balloon-like structure used by ‘higher’ flies to break out of the puparium.

pupa: stage in the lifecycle of an endopterygote insect between the larva and adult.

puparium: hardened skin of last-larval stage of ‘higher’ flies (Diptera); the pupa develops inside.

receptacle: swollen top of stem that supports the florets in a thistle head.

rhachis: the central stalk of a fern frond or of a leaf divided into leaflets.

rhizome: underground stem.

RNA, ribonucleic acid: a molecule that governs protein synthesis in all types of cell.

root crown, root collar: the junction of the stem and root.

rosette gall: leafy gall formed of overlapping leaves, gall wider than long, originating from a bud; due to failure of internodes to lengthen (c.f. artichoke gall).

rostrum: the snout of a weevil.

saprotrophic, saprophagous: feeding on dead and decaying organisms.

sclerenchyma: woody tissue in plants.

scleromorphic: a heathland vegetation of low woody shrubs with small, tough, leathery, long-lived leaves, often resinous and aromatic.

sclerophyllous: plants with tough and leathery leaves.

senescence: the process of ageing.

septum: membrane or wall that divides one cell from the next; or separates a structure into sections.

setae: long hairs.

sexuales: males and females of aphids (see Box 4.5, Chapter 4).

sexuparae: the parents of aphid sexuales (see Box 4.5, Chapter 4).

sieve tubes: the vessels in phloem that transport sugars.

sorus (plural sori): structure that produces the spores of fungi.

speciation: the evolution of new species.

species: (see phylum).

species diversity: a community attribute that depends on both the number of species and their relative abundance.

species richness: the number of species present in an area.

spermatophore: structure in which sperm are stored.

spongy mesophyll: (see mesophyll).

spontaneous generation: the idea that some organisms (e.g. germs, maggots, lice, intestinal worms) arose spontaneously from filth and dirt, i.e. refuting creationism as well as evolution.

sternal spatula: structure on underside of the thorax of full-grown larvae of Cecidomyiidae, important in identification (it is absent in some species).

stigma: part of the flower that receives pollen.

stoma (plural stomata): hole in leaf or stem that allows exchange of gases in and out of the plant.

style: stalk that bears the stigma at its tip.

successori: species that live in the gall after the galler, parasitoids and inquilines have left.

syconium: the fruit (inflorescence) of a fig.

symbiosis: the living together of two organisms (symbionts).

sympatric speciation: production by an ancestral species of a population whose members cannot breed with the parent species despite remaining in the same geographical area, so giving rise to a new species (c.f. allopatric speciation).

symplesiomorphy: an ancestral character shared by all members of a group (see Box 11.1, Chapter 11); some members may since have lost the character (c.f. synapomorphy).

synapomorphy: a derived character shared by other members of a group (see Box 11.1, Chapter 11; c.f. symplesiomorphy).

syncytium: giant multinucleate cells in galls of some nematodes.

thelytoky: parthenogenetic reproduction in which females develop from unfertilised diploid eggs (c.f. arrhenotoky).

tannins: toxins in leaves (e.g. of oaks) that deter attack by fungi and caterpillars; used for hardening (tanning) leather.

tarsus: part of the ‘foot’ of an insect.

top-down effects: based on the idea of a simple food chain: plant herbivore predator; the numbers of the herbivore, i.e. the gall causer, are affected by its enemies (parasitoids, predators and inquilines) (c.f. bottom-up effects; see Chapter 10).

totipotent: (see omnipotent).

transpiration: the loss of water vapour through the stomata of the plant.

tripinnate: frond divided into three (in ferns).

tuber: swollen underground stem.

unicellular: organism with one cell.

unilocular: a gall with one chamber (c.f. multilocular).

univoltine: a lifecycle with one generation a year (c.f. bivoltine, multivoltine).

vagrant: a species of mite that wanders on the leaf surface without causing a gall.

vascular tissue, vascular bundle: tissue (e.g. in leaf veins) in which water and nutrients are transported around the plant (c.f. xylem, phloem).

vector: organism than transports a parasite from one host to another.

virescence: a growth consisting of distorted and multiplied parts of a plant, either greened and leafy or woody and twiggy (c.f. phyllanthy, witches’ broom); or a single misshapen plant part.

viviparous: producing live young, not eggs (c.f. oviparous, ovoviviparous).

wing venation: the pattern of veins in an insect’s wing.

witches’ broom: cluster of twigs forming an untidy growth on branch or trunk of trees (c.f. chloranthy, phyllanthy).

xylem: vessels in vascular tissue that transport water and nutrients from roots to the rest of the plant (c.f. phloem).

zygote: the fertilised egg.

ABBREVIATIONS

f.sp., formae speciales: forms of a species that are morphologically identical but are restricted to different related host plants.

micron (µm): one thousandth of a millimetre (mm).

s.s., sensu stricto: in a narrow sense (used of species and genera).

+,–: complementary strains, e.g. of rust fungi (see Box 2.4, Chapter 2).

Appendices

APPENDIX A: GALL INDUCERS

VIRUSES

Cocoa swollen shoot virus (CSSV)

Lettuce big vein virus (LBVV)

Wound tumour virus (WTV)

BACTERIA AND PHYTOPLASMAS

(alphabetical order of species, with order and family)

Agrobacterium tumefaciens (Smith & Townsend), crown gall, Rhizobiales: Rhizobiaceae

Agrobacterium rhizogenes (Riker et al.) Conn, Rhizobiales: Rhizobiaceae

Pseudomonas syringae van Hall. (= Xanthomonas syringae), Xanthomonadales: Xanthomonadaceae

Rhodococcus fascians (Tilford) Dowson (= Corynebacterium fascians), Actinomycetales: Nocardiaceae

Xanthomonas campestris (Pammel) Dowson, pathovar juglandis, Xanthomonadales: Xanthomonadaceae

Xanthomonas populi (Ridé) van den Mooter & Swings, Xanthomonadales: Xanthomonadaceae

Xanthomonas syringae (see Pseudomonas syringae)

FUNGI: all groups traditionally studied by mycologists

(alphabetical order of species, with order and family)

Albugo candida (Pers.) O.Kuntze, white blister, Albuginales: Albuginaceae

Claviceps purpurea (Fr.) Tul. & C.Tul., ergot, Hypocreales: Clavicipitaceae

Cronartium ribicola J.C. Fischer, white pine blister rust, Uredinales: Cronartiaceae Exobasidium myrtilli Siegm. (= E. vaccinii), Entomatales: Exobasidiaceae

Exobasidium rhododendri Cramer, Entomatales: Exobasidiaceae

Gymnosporangium clavariiforme (Pers.) DC., Uredinales: Pucciniaceae

Gymnosporangium cornutum Kern, Uredinales: Pucciniaceae

Gymnosporangium juniperi-virginianae Schwein, cedar–apple rust, Uredinales: Pucciniaceae

Lachnellula wilkommii (Hartig) Dennis, Heliotiales: Hyaloscyphaceae

Melampsorella caryophyllacearum Schroet., Uredinales: Melampsoraceae

Microbotryum violaceum (Pers.) G. Geml & Oberw. (= Ustilago violaceum), Ustilaginales: Microbotryaceae

Phragmidium tuberculatum J. Müller, Uredinales: Phragmidiaceae

Plasmodiophora brassicae Woronin, clubroot, Plasmodiophorida: Plasmodiophoraceae

Pucciniastrum geoppertianum (Kühn) Kleb., Uredinales: Pucciniastraceae

Puccinia graminis Pers., Uredinales: Pucciniaceae

Puccinia punctiformis (Strauss) Rohl., Uredinales: Pucciniaceae

Ramichloridium subulatum (de Hoog) Arzaulou, Gams & Crous, Capnodiales: Mycosphaerellaceae

Spongospora subterranea (Wallr.) Lagerh., potato powdery scab, Plasmodiophorida: Plasmodiophoraceae

Synchytrium anemones (DC.) Woronin, Chytridiales: Synchytriaceae

Synchytrium endobioticum (Schilb.) Percival, potato wart disease, Chytridiales: Synchytriaceae

Synchytrium mercurialis (Lib.) Fuckel, Chytridiales: Synchytriaceae

Synchytrium taraxaci de Bary & Woronin, Chytridiales: Synchytriaceae

Taphrina alni (Berk. & Br.) Gjaerum (= T. amentorum), Taphrinales: Taphrinaceae

Taphrina betulina Rostrup, birch witches’ broom, Taphrinales: Taphrinaceae

Taphrina deformans (Berk.) Tulasne, almond and peach leaf curl, Taphrinales: Taphrinaceae

Taphrina populina Fr., Taphrinales: Taphrinaceae

Taphrina pruni Tulasne, pocket plum, Taphrinales: Taphrinaceae

Taphrina tosquinetii (Westend.) Magn., Taphrinales: Taphrinaceae

Taphrina wiesneri (Rathay) Mix (= T. cerasi), Taphrinales: Taphrinaceae

Tranzschelia anemones (Pers.) Nannf., Uredinales: Uropyxidaceae

Tranzschelia discolor (Fuckel) Tranz. & Litv., Uredinales: Uropyxidaceae

Uromyces pisi-sativi (Pers.) Liro, Uredinales: Pucciniaceae

Ustilago maydis (DC.) Corda, sweetcorn smut, Ustilaginales: Ustilaginaceae

Ustilago scitaminea Sydow, Ustilaginales: Ustilaginaceae

PLANTS

(alphabetical order of species, with family)

Rhinanthus minor L., Scrophulariaceae

Thesium humifusum DC., Santalaceae

Viscum album L., mistletoe, Viscaceae

Nematodes

(alphabetical order of species, with family)

Anguina tritici (Steinbuch) Chitwood, Anguinidae

Aphelenchoides ritzemabosi (Schwartz) Steiner & Buhrer, Aphelencoididae

Ditylenchus dipsaci (Kühn) Filip’jev, Anguinidae

Subanguina millefolii (Löw) Chizhov & Subbotin, Anguinidae

Subanguina picridis (Kiranjova) Brzeski, Anguinidae

Rotifers (with family)

Proales werneckii (Ehrenberg) (= Notommata werneckii), Proalidae

Eriophyoidea, gall mites, and Tarsonemoidea

(alphabetical order of species, with family.

All belong to the superfamily Eriophyoidea except Steneotarsonemus, which belongs to the Tarsonemoidea.)

Acalitus brevitarsus (Fockeu), Eriophyidae

Acalitus lowei Manson, Eriophyidae

Acalitus phloeocoptes (Nalepa), Eriophyidae

Aceria campestricola (Nalepa) (= Eriophyes ulmicola), Eriophyidae

Aceria erinea (Nalepa), Eriophyidae

Aceria filiformis (Nalepa), Eriophyidae

Aceria genistae (Nalepa), Eriophyidae

Aceria hindali (a new species, not yet formally named), Eriophyidae

Aceria lycopersici (Wolffenstein) (= A. cladophthirus), Eriophyidae

Aceria macrorhyncha (Nalepa) (= Eriophyes macrorhynchus, Artacris macrorhynchus), Eriophyidae

Aceria nalepai (Fockeu) (= Eriophyes inangulis), Eriophyidae

Aceria nervisequa (Canestrini), Eriophyidae

Aceria populi (Nalepa), Eriophyidae

Aceria tenuis (Nalepa), Eriophyidae

Aceria tristriata (Nalepa) (= Eriophyes tristriatus), Eriophyidae

Aceria varia (Nalepa), Eriophyidae

Aculops macrotrichus (Nalepa) (= Aceria macrotrichus), Eriophyidae

Aculus convolvuli (Nalepa) (= Aceria convolvuli), Eriophyidae

Aculus hippocastani (Fockeu), Eriophyidae

Aculus laevis (Nalepa), Eriophyidae

Cecidophyopsis psilaspis (Nalepa), yew big bud mite, Eriophyidae

Cecidophyopsis ribis (Westwood), blackcurrant big bud mite, Eriophyidae

Colomerus vitis (Pagenstecher), Eriophyidae

Epitrimerus trilobus (Nalepa), Eriophyidae

Eriophyes arianus (Canestrini), Eriophyidae

Eriophyes dichrostachia Tuck., Eriophyidae

Eriophyes laevis Nalepa (= Phytoptus laevis), Eriophyidae

Eriophyes leiosoma (Nalepa), Eriophyidae

Eriophyes distinguendus Nalepa (= E. paderinus), Eriophyidae

Eriophyes pyri (Pagenstecher), Eriophyidae

Eriophyes similis (Nalepa), Eriophyidae

Eriophyes tiliae (Pagenstecher) (= Phytoptus tiliae), Eriophyidae

Eriophyes viburni (Nalepa), Eriophyidae

Phyllocoptes eupadi (Newkirk) (= Eriophyes padi), Eriophyidae

Phyllocoptes goniothorax (Nalepa), Eriophyidae

Phytoptus avellanae Nalepa (= Phytocoptella avellanae), hazel big bud mite, Phytoptidae

Phytoptus bursarius (Nalepa), Phytoptidae

Phytoptus tetratrichus (Nalepa) (= Phytocoptella tetratrichus), Phytoptidae

Steneotarsonemus nitidus Beer, Tarsonemidae

Steneotarsonemus phragmitidis (Schlechtendal), Tarsonemidae

Steneotarsonemus spirifex (Marchal), Tarsonemidae

Trisetacus pini (Nalepa), Phytoptidae

Vasates quadripedes (Shimer), Eriophyidae

Hemiptera: Psylloidea, psylloids

(alphabetical order of species)

Craspedolepta subpunctata (Förster)

Livia juncorum Latreille, tassel gall

Pachypsylla celtidisgemma Riley

Pachypsylla celtidismamma (Fletcher)

Spanioneura buxi (L.) (=Psylla buxi), box cabbage gall

Trichochermes walkeri Förster

Trioza alacris Flor

Hemiptera: Phylloxeridae, phylloxerids

Daktulosphaira vitifoliae (Fitch) (= Viteus vitifoliae = Phylloxera vastatrix), grape phylloxera

Hemiptera: Adelgidae, adelgids

Adelges abietis (L.)

Adelges laricis Vallot

Hemiptera: Aphididae, aphids

(alphabetical order of species, with subfamily)

Aploneura lentisci Passerini, Eriosomatinae

Asiphonella dactylonii Theobald, Eriosomatinae

Baizongia pistaciae (L.), Eriosomatinae

Ceratoglyphina bambusae van der Groot (= Astegopteryx styracicola), Hormaphidinae

Dysaphis devecta (Walker), Aphidinae

Dysaphis plantaginea (Passerini), Aphidinae

Eriosoma ulmi (L.), Eriosomatinae

Forda marginata Koch, Eriosomatinae

Forda formicaria von Heyden, Eriosomatinae

Forda riccobonii (Stefani), Eriosomatinae

Geoica wertheimae Brown & Blackman (= G. utricularia), Eriosomatinae

Paracletus cimiciformis Heyden, Eriosomatinae

Pemphigus betae Doanne, Eriosomatinae

Pemphigus bursarius (L.), Eriosomatinae

Pemphigus dorocola Matsumara, Eriosomatinae

Pemphigus populinigrae (Shrank) (= P. filaginis), Eriosomatinae

Pemphigus spyrothecae Passerini, Eriosomatinae

Schlechtendalia chinensis (Bell), Chinese gall, Eriosomatinae

Slavum wertheimae Hille Ris Lambers, Eriosomatinae

Smynthurodes betae Westwood, Eriosomatinae

Tamalia coweni (Cockerell), Drepanosiphinae

Tetraneura ulmi (L.), Eriosomatinae

Thecabius affinis (Kaltenbach), Eriosomatinae

Hemiptera: Coccoidea, scale insects

(alphabetical order of species, with family)

Apiomorpha munita (Schrader) (= A. egeria), Brachyscelidae

Asterodiaspis minus (Lindinger), Asterolecaniidae

Asterodiaspis quercicola (Bouché), Asterolecaniidae

Asterodiaspis variolosa (Ratzeburg), Asterolecaniidae

Cystococcus pomiformis (Froggatt), Eriococcidae

Gallacoccus anthonyae Beardsley, Beesoniidae

Gallacoccus secundus Beardsley, Beesoniidae

Mangalorea hopeae Takagi, Beesoniidae

Maskellia globosa Fuller (= Brachyscelis pedunculata), Diaspididae

Thysanoptera, thrips

(alphabetical order of species; all belong to the Phlaeothripidae)

Alocothrips hadrocerus (Karny)

Arrhenothrips ramakrishnae (Hood)

Austrothrips cochinchinensis (Karny)

Crotonothrips memecylonicus Ananthakrishnan

Gynaikothrips crassipes Karny

Gynaikothrips ficorum (Marchal)

Iotatubothrips crozieri Mound & Crespi

Liothrips karnyi (Bagnall)

Liothrips ramakrishnae Ananthakrishnan & Jagadish

Lygothrips jambuvasi (Ramakrishna)

Oncothrips rodwayi (Hardy)

Thilakothrips babuli Ramakrishna

Lepidoptera, moths (with family)

Heliozela sericiella (Haworth), Heliozelidae

Retinia resinella (L.) (= Petrova resinella =

Evetria resinella), Tortricidae

DIPTERA, flies

(alphabetical order of families and species)

Anthomyiidae

Chirosia grossicauda (Strobl) (= C. parvicornis)

Cecidomyiidae, gall midges

Asphondylia atriplicis Townsend

Asphondylia aucubae Yukawa & Ohsaki

Asphondylia baca Monzen

Anisostephus betulinus (Kieffer)

Asphondylia gennadii (Marchal)

Asphondylia itoi Uechi & Yukawa

Asphondylia morivorela (Naito)

Asphondylia pruniperda Rondani

Asphondylia sarothamni (Loew)

Asphondylia sphaera Monzen

Asphondylia ulicis Trail

Asphondylia yushimai Yukawa & Uechi

Contarinia manii Harris

Contarinia ramachandrani Mani

Craneiobia corni (Giraud), dogwood rivet gall midge

Cystiphora sonchi (Bremi)

Dasineura crataegi (Winnertz), hawthorn button-top gall midge

Dasineura mali (Kieffer), apple leaf midge

Dasineura odoratae Stelter (= Dasineura affinis)

Dasineura pustulans (Rübsaamen)

Dasineura pyri Bouché, pear leaf midge

Dasineura ulmaria (Bremi)

Dasineura urticae (Perris)

Didymomyia tiliaceae (Bremi) (= D. reaumuriana)

Drisina glutinosa Giard

Fabomyia medicaginis (Rübsaamen)

Geocrypta galii (Loew)

Giraudiella inclusa (Frauenfeld)

Harmandiola tremulae (Winnertz)

Hartigiola annulipes (Hartig)

Lasioptera arundinis Schiner

Lasioptera rubi (Schrank)

Massalongia betulifolia Harris

Massalongia ruber (Kieffer) (= M. rubra)

Mayetiola destructor (Say), Hessian fly

Mayetiola graminis (Fourcroy) (= M. poae)

Mikiola fagi (Hartig)

Monarthropalpus flavus (Schrank) (= M. buxi)

Orseolia oryzae Wood-Mason

Orseolia oryzivora Harris & Gagné

Physemocecis hartigi (Liebel)

Physemocecis ulmi (Kieffer)

Pseudasphondylia matatabi (Yuasa & Kumazawa)

Rabdophaga degeeri (Bremi) (= R. ramicola)

Rabdophaga heterobia (Loew)

Rabdophaga rosaria (Loew)

Rabdophaga salicis (Schrank)

Rabdophaga strobiloides (Osten-Sacken)

Rondaniola bursaria (Bremi), lighthouse gall

Semudobia betulae (Winnertz)

Semudobia skuhravae Roskam

Semudobia tarda Roskam

Sequoiomyia kraeuseli Möhn

Spurgia euphorbiae (Vallot) (= S. capitigena)

Taxomyia taxi (Inchbald), yew artichoke gall midge

Chloropidae, chloropid flies

Lipara lucens Meigen

Lipara pullitarsis Doskoil & Chvala

Lipara rufitarsis (Loew)

Tephritidae, tephritid flies

Eurosta solidaginis Fitch

Myopites eximia Séguy

Myopites inulaedyssentericae Blot

Stamnophora vernoniicola (Bezzi)

Terellia tussilaginis (F.) (= Cerajocera tussilaginis)

Tephritis bardanae (Schrank)

Tephritis conura (Loew)

Tephritis formosa (Loew)

Urophora affinis (Frauenfeld)

Urophora cardui (L.)

Urophora congrua Loew

Urophora cuspidata (Meigen)

Urophora dzieduszyckii Frauenfeld

Urophora jaceana (Hering)

Urophora jaculata Rondani

Urophora mauritanica Macquart

Urophora misakiana (Matsumura)

Urophora nigripalpis Hering

Urophora quadrifasciata (Meigen)

Urophora sirunaseva (Hering)

Urophora solstitialis (L.)

Urophora stylata (F.)

Urophora terebrans (Loew)

HYMENOPTERA: sawflies and wasps

(alphabetical order of families and species)

Agaonidae

Ceratosolen arabicus Mayr

Ceratosolen capensis Grandi

Ceratosolen galili Wiebes

Ceratosolen pilipes Wiebes

Cynipidae, gall wasps

Andricus aries Mayr, ramshorn gall wasp

Andricus caliciformis (Giraud) (= Cynips caliciformis), mulberry gall

Andricus caputmedusae (Hartig)

Andricus conficus (Hartig)

Andricus coriarius (Hartig)

Andricus coronatus (Giraud)

Andricus corruptrix (Schlechtendal)

Andricus curvator Hartig

Andricus curtisii (Müller)

Andricus dentimitratus (Rejiö)

Andricus foecundatrix (Hartig) (= A. fecundator), oak artichoke gall wasp

Andricus gallaeurniformis (Fonscolombe)

Andricus galeatus (Giraud)

Andricus gemmeus Giraud

Andricus grossulariae Giraud (= A. mayri, = A. panteli)

Andricus hartigi (Hartig)

Andricus hispanicus Pujade Villar, Iberian marble gall wasp

Andricus hungaricus (Hartig)

Andricus infectorius (Hartig) (= A. tinctoriusnostris, = A. gallaetinctoriae, = Cynips tinctoria)

Andricus inflator Hartig (= A. globuli)

Andricus kollari (Hartig) (= Cynips Kollari), marble gall wasp

Andricus lignicolus (Hartig), cola nut gall wasp

Andricus lucidus (Hartig) (= A. aestivalis), hedgehog gall wasp

Andricus malpighii (Adler)

Andricus mitratus (Mayr)

Andricus multiplicatus Giraud

Andricus paradoxus (Radoskowsky) (= A. albopunctatus)

Andricus polycerus Giraud

Andricus quadrilineatus Hartig

Andricus quercuscalicis Burgsdorf, knopper gall wasp

Andricus quercusradicis (F.) (= A. radicis)

Andricus quercusramuli (L.) (= A. autumnalis), cottonwool gall

Andricus quercustozae (Bosc)

Andricus quercustozae f. insana (Bosc) (= Cynips insana), Bassorah gall wasp

Andricus schroeckingeri Wachtl

Andricus seckendorffi (Wachtl)

Andricus sieboldi (Hartig) (= A. testaceipes,

Aphilothrix Sieboldi)

Andricus solitarius (Fonscolombe)

Andricus theophrastea (Trotter) (= A.

Theophrasteus), woolly or cotton gall

Andricus viscosus (see Andricus dentimitratus)

Aphelonyx cerricola Giraud

Aulacidea hieracii (Bouché)

Aulacidea phlomica Belizin

Aulacidea verticillica Belizin

Aylax minor Hartig

Aylax papaveris (Perris)

Biorhiza pallida (Olivier) (= B. aptera, Teras terminalis), oak apple gall wasp

Callirhytis erythrocephala (Giraud) (= C. glandium)

Cynips agama Hartig

Cynips disticha Hartig

Cynips divisa Hartig (= Dryophanta divisa), red pea gall wasp

Cynips longiventris Hartig (= Dryophanta longiventris), striped pea gall wasp

Cynips quercusfolii L. (= Dryophanta scutellaris), cherry gall wasp

Diastrophus kincaidii Gillette

Diastrophus rubi (Bouché)

Diplolepis bicolor (Harris)

Diplolepis eglanteriae (Hartig), rose pea gall wasp

Diplolepis fusiformis (Ashmead)

Diplolepis mayri (Schlechtendal)

Diplolepis nervosa (Curtis), rose pea gall wasp

Diplolepis nodulosa (Beutenmüller)

Diplolepis polita (Ashmead)

Diplolepis rosae (L.), robin’s pincushion, bedeguar gall wasp

Diplolepis spinosa (Ashmead)

Diplolepis spinosissimae (Giraud)

Diplolepis triforma Shorthouse & Richie

Disholcaspis eldoradensis (Beutenmüller), the honeydew gall wasp

Disholcaspis weldi (Beutenmüller)

Dryocosmus cerriphilus Giraud

Dryocosmus deciduus (Beutenmüller)

Dryocosmus kuriphilus Yasumatsu, chestnut gall wasp

Isocolus rogenhoferi Wachtl (may be a synonym of I. scabiosae)

Isocolus scabiosae (Giraud)

Liposthenes glechomae (Kieffer)

Neuroterus albipes (Schenck) (= Spathegaster albipes, = N. laeviusculus), smooth spangle gall wasp

Neuroterus anthracinus (Curtis) (= Andricus ostreus), oyster gall

Neuroterus numismalis (Geoffroy in Fourcroy) (= N. numismatis), silk button spangle gall wasp

Neuroterus politus Hartig (= N. petioliventris,

N. aprilinus, Spathegaster aprilinus)

Neuroterus quercusbaccarum (L.) (= Spathegaster baccarum, = N. lenticularis), common spangle gall wasp

Neuroterus saliens (Kollar)

Neuroterus tricolor Hartig (= N. fumipennis), cupped spangle gall wasp

Pediaspis aceris (Gmelin), sycamore gall wasp

Phanacis centaureae Förster

Phanacis hypochoeridis (Kieffer)

Plagiotrochus amenti Kieffer (= P. suberi)

Plagiotrochus australis (Mayr)

Plagiotrochus coriaceus (Mayr)

Plagiotrochus quercusilicis (F.)

Trigonaspis megaptera (Panzer) (= Biorhiza renum)

Xestophanes potentillae (Retzius)

Pteromalidae, parasitic wasps

Trichilogaster acaciaelongifoliae (Froggatt)

Tenthredinidae, sawflies

Blennocampa phyllocolpa (Viitasaari & Vikberg)

Eupontania aestiva (Thomson)

Eupontania arcticornis Konow

Eupontania collactanea (Förster)

Eupontania glabrifrons Benson

Eupontania hastatae Vikberg

Eupontania lapponica Malaise

Eupontania nivalis Vikberg

Eupontania pedunculi (Hartig)

Eupontania polaris Malaise

Eupontania pustulator Forsius

Eupontania reticulatae Malaise

Eupontania samolad Malaise

Eupontania viminalis (L.)

Euura amerinae (L.)

Euura atra (Jurine)

Euura lanatae Malaise

Euura lasiolepis Smith

Euura mucronata (Hartig)

Euura testaceipes (Brischke)

Euura venusta (Brischke)

Phyllocolpa anglica (Cameron)

Phyllocolpa coriacea (Benson)

Phyllocolpa excavata (Marlatt)

Phyllocolpa leucosticta (Hartig)

Phyllocolpa nudipectus Vikberg

Phyllocolpa puella (Thomson)

Pontania bridgmanii (Cameron)

Pontania dolichura (Thomson)

Pontania proxima (Audinet-Serville)

Pontania triandrae Benson

Coleoptera, beetles

(alphabetical order of species, with family)

Cleonis pigra (Scopoli) (= Cleonus piger), Curculionidae

Eutrichapion scutellare (Kirby), Apionidae

Rhinocyllus conicus (Fröhlich), Curculionidae

Saperda populnea (L.), Cerambycidae

Sphenoptera jugoslavica Obenberger, Buprestidae

Undetermined fossil holometabolous insect

Pteriditorichnos stipitopterii Labandeira & Phillips

APPENDIX B: HOST PLANTS ALGAE

Vaucheria dilatata (Roth) (Vaucheriales: Xanthophyceae)

PLANTS

(alphabetical order of species, with family)

Abies alba Mill., European silver-fir, Pinaceae

Acacia catechu Willd., Mimosaceae

Acacia ferruginea DC., Mimosaceae

Acacia leucophloea Willd., Mimosaceae

Acacia longifolia Paxt., Mimosaceae

Acacia pycnantha Benth., Mimosaceae

Acacia tortillis (Forssk.) Hague, Mimosaceae

Acer pseudoplatanus L., sycamore, Aceraceae

Acer saccharinum L., silver maple, Aceraceae

Achillea millefolium L., yarrow, Asteraceae

Acroptilon repens (L.) de Candolle, Russian knapweed, Asteraceae

Aesculus hippocastani L., horsechestnut, Hippocastanaceae

Allium cepa L., onion, Liliaceae

Alnus glutinosa (L.) Gaertn., alder, Betulaceae

Ammophila arenaria (L.) Link, marram grass, Poaceae

Anemone coronaria L., Ranunculaceae

Anemone nemorosa L., wood anemone, Ranunculaceae

Anogeissus latifolia (Roxb. ex DC.) ex Guill. & Perr., Combretaceae

Apodytes dimidiata E. Meyer ex Arn., Icacinaceae

Arctium minus (Hill) Bernh., lesser burdock, Asteraceae

Araucaria araucana (Molina) K. Koch, monkey-puzzle, Araucariaceae

Atriplex canescens (Pursch) Nuttall, Chenopodiaceae

Avena sativa L., oats, Poaceae

Berberis vulgaris L., common barberry, Berberidaceae

Betula pendula Roth., silver birch, Betulaceae

Betula pubescens Ehrh., downy birch, Betulaceae

Brachypodium sylvaticum (Huds.) Beauv., false brome, Poaceae

Brassica oleracea L., cabbage, cauliflower, Brassicaceae

Bromopsis erecta (Huds.) Fourr., upright brome, Poaceae

Buxus sempervirens L., box, Buxaceae

Calluna vulgaris (L.) Hull, heather, ling, Ericaceae

Calycopteris floribundus Lamk., Combretaceae

Capsella bursa-pastoris (L.) Medik., shepherd’s-purse, Brassicaceae

Carduus acanthoides L., Asteraceae

Carduus nutans L., nodding thistle, Asteraceae

Celtis occidentalis L., hackberry, Ulmaceae

Centaurea biebersteinii DC., spotted knapweed, Asteraceae

Centaurea diffusa Lam., diffuse knapweed, Asteraceae

Centaurea jacea L., brown knapweed, Asteraceae

Centaurea maculosa Lam., spotted knapweed, Asteraceae

Centaurea nigra L., black knapweed, Asteraceae

Centaurea scabiosa L., greater knapweed, Asteraceae

Chamerion angustifolium (L.) Holub, rosebay willowherb, Onagraceae

Cinnamomum osmophloeum Kaneh., Lauraceae

Cirsium arvense (L.) Scop., creeping thistle, Asteraceae

Cirsium setidens (Dunn) Nakai, Asteraceae

Cirsium vulgare (Savi) Ten., spear thistle, Asteraceae

Cornus sanguinea L., dogwood, Cornaceae

Corylus avellana L., hazel, Betulaceae

Crataegus monogyna Jacq., hawthorn, Rosaceae

Crataegus laevigata (Poir.) DC., midland hawthorn, Rosaceae

Cytisus scoparius (L.) Link, broom, Fabaceae

Dactylis glomerata L., cock’s-foot, Poaceae

Dichrostachys glomerata (Forssk.) Chiov., Fabaceae

Elytrigia repens (L.) Desv. ex Nevsky, couch, Poaceae

Erysimum cheiri (L.) Crantz, wallflower, Brassicaceae

Euphorbia cyparissias L., cypress spurge, Euphorbiaceae

Fagus sylvatica L., beech, Fagaceae

Ficus retusa L., fig, Moraceae

Filipendula ulmaria (L.) Maxim., meadowsweet, Rosaceae

Forsythia x intermedia Zabel, forsythia, Oleaceae

Fragaria x ananassa (Duchesne) Duchesne, garden strawberry, Rosaceae

Fraxinus excelsior L., ash, Oleaceae

Galium mollugo L., hedge bedstraw, Rubiaceae

Galium verum L., lady’s bedstraw, Rubiaceae

Glechoma hederacea L., ground-ivy, Lamiaceae

Hopea ponga (Dennst.) Mabberley, Dipterocarpaceae

Hordeum distichon L., two-rowed barley, Poaceae

Hypochaeris radicata L., common cat’s-ear, Asteraceae

Inula crithmoides L., golden-samphire, Asteraceae

Juglans regia L., walnut, Juglandaceae

Juncus articulatus L., jointed rush, Juncaceae

Juncus squarrosus L., heath rush, Juncaceae

Juniperus communis L., common juniper, Cupressaceae

Larix decidua Mill., European larch, Pinaceae

Larrea tridentata (DC.) Colville, creosote bush, Asteraceae

Laurus nobilis L., bay laurel, Lauraceae

Lycopersicon esculentum Mill., tomato, Solanaceae

Mahonia aquifolium (Pursh) Nutt., Oregon-grape, Berberidaceae

Malus domestica Borkh., cultivated apple, Roscaeae

Malus sylvestris (L.) Mill., crab apple, Rosaceae

Maytenus senegalensis (Lam.) Excell., Celastraceae

Medicago sativa L., lucerne, Fabaceae

Melilotus albus Medik., white melilot, white sweet clover, Fabaceae

Memecylon lushingtonii Gamble, Melastomataceae

Mercurialis perennis L., dog’s mercury, Euphorbiaceae

Mimusops elengi L., Sapotaceae

Oryza sativa L., rice, Poaceae

Papaver dubium L., long-headed poppy, Papaveraceae

Papaver rhoeas L., field poppy, Papaveraceae

Phragmites australis (Cav.) Trin. ex Steud. (= P. communis), common reed, Poaceae

Picea abies (L.) Karst., Norway spruce, Pinaceae

Picea sitchensis (Bong.) Carr.), sitka spruce, Pinaceae

Pinus cembra L., Pinaceae

Pinus contorta Douglas ex Loudon, lodgepole pine, Pinaceae

Pinus strobus L., Weymouth pine, Pinaceae

Pinus sylvestris L., Scots pine, Pinaceae

Piper nigrum L., pepper, Piperaceae

Pistacia atlantica Desf., Anacardiaceae

Pistacia lentiscus L., Anacardiaceae

Pistacia palaestina Boiss., Anacardiaceae

Pisum sativum L., garden pea, Fabaceae

Plantago lanceolata L., ribwort plantain, Plantaginaceae

Poa nemoralis L., wood meadow-grass, Poaceae

Populus x candicans (Aiton) Sm., grey poplar, Salicaceae

Populus angustifolia James, narrowleaf cottonwood, Salicaceae

Populus nigra L., black poplar, Salicaceae

Populus tremula L., aspen, Salicaceae

Prunus avium (L.) L., wild cherry, Rosaceae

Prunus cerasus L., dwarf cherry, Rosaceae

Prunus domestica L., wild plum, Rosaceae

Prunus dulcis (Mill.) D.A.Webb, almond, Rosaceae

Prunus padus L., bird cherry, Rosaceae

Prunus persica (L.) Batsch, nectarine, peach, Rosaceae

Prunus spinosa L., blackthorn, sloe, Rosaceae

Psaronius chasei Morgan, Marattiaceae

Pseudostuga menziesii (Mirb.) Franco, douglas fir, Pinaceae

Pteridium aquilinum (L.) Kuhn, bracken, Dennstaedtiaceae

Pulicaria dysenterica (L.) Bernh., common fleabane, Asteraceae

Pyrus communis L., pear, Rosaceae

Quercus bicolor Willd., swamp white oak, Fagaceae

Quercus cerris L., Turkey oak, Fagaceae

Quercus cerris x Q. suber Lam., Lucombe oak (= Q. hispanica, = Q. pseudosuber), Fagaceae

Quercus coccifera L., kermes oak, Fagaceae

Quercus copeyensis C.M. Müll., Fagaceae

Quercus frainetto Ten. (= Q. conferta), Fagaceae

Quercus ilex L., holm oak, Fagaceae

Quercus infectoria Olivier, Aleppo oak, Fagaceae

Quercus lobata Née, California white oak, Fagaceae

Quercus lusitanica Lam., Fagaceae

Quercus macrolepis Kotschy (= Q. aegilops), Fagaceae

Quercus montana Willd., chestnut oak, Fagaceae

Quercus petraea (Matt.) Liebl. (= Q. sessiflora), sessile oak, Fagaceae

Quercus pubescens Willd., downy oak, Fagaceae

Quercus pyrenaica Willd. (= Q. Toza), Fagaceae

Quercus robur L. (= Q. pedunculata), pedunculate oak, Fagaceae

Quercus suber L., cork oak, Fagaceae

Quercus trojana Webb (= Q. macedonius), Macedonian oak, Fagaceae

Rhamnus cathartica L., buckthorn, Rhamnaceae

Rhododendron ponticum L., rhododendron, Ericaceae

Rhus semialata Murray, sumach, sumac, Anacardiaceae

Ribes nigrum L., blackcurrant, Grossulariaceae

Ribes rubrum L., redcurrant, Grossulariaceae

Robinia pseudacacia L., false-acacia, Fabaceae

Rosa acicularis Lindl., Rosaceae

Rosa arvensis Hudson, field rose, Rosaceae

Rosa blanda Aiton, Rosaceae

Rosa canina L., dog rose, Rosaceae

Rosa pimpinellifolia L., burnet rose, Rosaceae

Rosa rubiginosa L., sweet-briar, Rosaceae

Rubus caesius L., dewberry, Rosaceae

Rubus idaeus L., raspberry, Rosaceae

Rubus fruticosus L. agg., brambles, Rosaceae

Rubus parviflorus Nutt., thimbleberry, Rosaceae

Rubus saxatilis L., stone bramble, Rosaceae

Saccharum L., sugarcane, Poaceae

Salix alba L., white willow, Salicaceae

Salix aurita L., eared willow or sallow, Salicaceae

Salix borealis (Fr.), Salicaceae

Salix caprea L., goat willow or sallow, Salicaceae

Salix cinerea L., grey willow or sallow, Salicaceae

Salix dasyclados Wimm., Salicaceae

Salix eriocephala Michx. (= Salix cordata), heart-leaved willow, Salicaceae

Salix fragilis L., crack-willow, Salicaceae

Salix glauca L., Salicaceae

Salix gracilistyla Miq., Salicaceae

Salix hastata L., Salicaceae

Salix lanata L., woolly willow, Salicaceae

Salix lapponum L., downy willow, Salicaceae

Salix lasiolepis Bentham, arroyo willow, Salicaceae

Salix myrsinifolia Salisb., dark-leaved willow, Salicaceae

Salix pentandra (L.), bay willow, Salicaceae

Salix phylicifolia L., tea-leaved willow, Salicaceae

Salix polaris Wahlenb., Salicaceae

Salix purpurea L., purple willow, Salicaceae

Salix repens L., creeping willow, Salicaceae

Salix reticulata L., net-leaved willow, Salicaceae

Salix schwerinii E. Wolf, Salicaceae

Salix triandra L., basket or almond willow, Salicaceae

Sambucus nigra L., elder, Caprifoliaceae

Schefflera racemosa Harms, Araceae

Secale cereale L., rye, Poaceae

Sequoia langsdorfi (Brong.) Heer, Taxodiaceae

Serratula tinctoria L., saw-wort, Asteraceae

Shorea curtisii Dyer ex King, Dipterocarpaceae

Solanum dulcamara L., bittersweet, woody nightshade, Solanaceae

Solanum nigrum L., black nightshade, Solanaceae

Solanum tuberosum L., potato, Solanaceae

Solidago altissima L., late goldenrod, Asteraceae

Solidago canadensis L. agg., goldenrod, Asteraceae

Solidago gigantea Aiton, tall goldenrod, Asteraceae

Sonchus arvensis L., perennial sow-thistle, Asteraceae

Sorbus aria (L.) Crantz, common whitebeam, Rosaceae

Sorbus aucuparia L., rowan, Rosaceae

Taraxacum officinale agg., dandelions, Asteraceae

Taxus baccata L., yew, Taxaceae

Tilia cordata Mill., small-leaved lime, Tiliaceae

Tilia x europaea L., common lime, Tiliaceae

Tilia platyphyllos Scop., large-leaved lime, Tiliaceae

Trifolium repens L., white clover, Fabaceae

Triticum aestivum L., wheat, Poaceae

Ulex europaeus L., common or European gorse, Fabaceae

Ulex minor Roth, dwarf gorse or furze, Fabaceae

Ulmus glabra Hudson, wych elm, Ulmaceae

Ulmus procera Salisb., English elm, Ulmaceae

Urtica dioica L., common nettle, stinging nettle, Urticaceae

Vaccinium myrtillus L., bilberry, Ericaceae

Vaccinium vitis-idaea L., cowberry, Ericaceae

Veronica chamaedrys L., germander speedwell, Scrophulariaceae

Viburnum lantana L., wayfaring-tree, Caprifoliaceae

Viola odorata L., sweet violet, Violaceae

Vitis vinifera L., grapevine, Vitaceae

Zea mays L., maize, Poaceae

APPENDIX C: INQUILINES AND ENEMIES (PARASITOIDS AND PREDATORS) OF GALL CAUSERS

Fungi

(alphabetical order of species, with order and family)

Apiognomonia errabunda (Roberge ex Desm.) Höhn. (= Discula quercina), Diaporthales: Valsaceeae

Discula quercina (see Apiognomonia errabunda)

Hirsutella thompsoni Fischer, anamorph of

Ophiocordyceps, Hypocreales: Ophiocordycipitaceae

Macrosiphoniella sanborni (Gillette)

Metarhizium anisopliae (Metsch.) Sorok., anamorph of

Metacordyceps, Hypocreales: Clavicipitaceae

Pileolaria terebinthi (DC.) Castagne, Uredinales: Pileolariaceae

Plasmopara viticola Berlese & Toni, Peronosporales: Peronosporaceae

Verticillium lecanii (Zimm.) Viegas, anamorph, Hypocreales: Cordycipitaceae

Gall mites

(alphabetical order of species, with superfamily)

Aceria ulmi (Garman), Eriophyoidea

Cecidophyopsis psilaspis (Nalepa) (also a gall inducer, see Appendix A), Eriophyoidea

Steneotarsonemus phragmitidis (Schlechtendal)

(also a gall inducer, see Appendix A), Tarsonemoidea

Hemiptera

(alphabetical order of species, with family)

Anthocoris minki Dohrn, Anthocoridae

Anthocoris nemoralis (F.), Anthocoridae

Anthocoris nemorum (L.), Anthocoridae

Orius minutus (L.), Anthocoridae

Tamalia inquilinus Miller, Aphididae

Thysanoptera, thrips (with family)

Phallothrips houstoni Mound & Crespi, Phlaeothripidae

Thaumatothrips frogatti Karny, Phlaeothripidae

Lepidoptera, moths

(alphabetical order of species, with family)

Acleris schalleriana (L.), Tortricidae

Aethes rubigana (Treitschke), Cochylidae

Alophia combustella H.S., Pyralidae

Archips podana (Scopoli), Tortricidae

Batrachedra striolata Zeller, Batrachedridae

Blastobasis lignea (Walsingham), Blastobasidae

Ditula angustoriana (Haworth) (= Batodes angustoriana), Tortricidae

Eucosma cana (Haworth), Tortricidae

Metzneria lappa (L.), Gelechiidae

Operophtera brumata (L.), winter moth, Geometridae

Palumbina guerrini (Stainton), Gelechiidae

Pammene amygdalana Duponchel, Tortricidae

Pammene argyrana Hübner, Tortricidae

Diptera

(alphabetical order of species, with family)

Clinidiplosis cilicrus (Kieffer), Cecidomyiidae

Dasineura fastidiosa Roskam, Cecidomyiidae

Dasineura interbracta Roskam, Cecidomyiidae

Giraudiella inclusa (Frauenfeld), Cecidomyiidae (also a gall inducer, see Appendix A)

Heringia heringi (Zetterstedt) (= Pipizella heringi, = Pipiza heringi), Syrphidae

Heringia senilis Sack, Syrphidae

Lestodiplosis sp. near

trifolii Barnes, Cecidomyiidae

Leucopsis grunini Tanasijtshuk, Chamaemyiidae

Leucopsis palumbi Tanasijtshuk, Chamaemyiidae

Leucopsis steinbergi Tanasijtshuk, Chamaemyiidae

Palloptera parallela (Loew), Pallopteridae

Palloptera umbellatarum (F.), Pallopteridae

Parallelodiplosis galliperda (Löw), Cecidomyiidae

Pipiza festiva Meigen, Syrphidae

Rhacochlaena toxoneura (Loew) (= Euphranta toxoneura), Tephritidae

Trotteria obtusa (Loew) (= T. sarothamni), Cecidomyiidae

Xenodiplosis laeviusculi (Rübsaamen), Cecidomyiidae

HYMENOPTERA

(alphabetical order of families and species)

Aphidiidae, ichneumon wasps

Monoctonia pistaciaecola Starý

Braconidae, ichneumon wasps

Bracon discoideus Wesmael

Bracon minutator (F.)

Bracon picticornis Wesmael

Colastes braconius Haliday

Diaparsis stramineipes (Brischke)

Polemochartus liparae (Giraud)

Polemochartus melas (Giraud)

Cynipidae, gall wasps

Ceroptres arator Hartig

Periclistus brandtii (Ratzeburg)

Periclistus caninae (Hartig)

Saphonecrus connatus (Hartig)

Synergus albipes Hartig

Synergus apicalis Hartig

Synergus gallaepomiformis (Boyer de Fonscolombe)

Synergus nervosus Hartig

Synergus pallicornis Hartig

Synergus pallidipennis Mayr

Synergus reinhardi Mayr

Synergus thaumacerus (Dalman)

Synergus umbraculus (Olivier)

Diapriidae, proctotrupoid wasps

Spilomicrus stigmaticalis Westwood

Eulophidae, chalcid wasps

Aprostocetus aethiops (Zetterstedt)

Aprostocetus sp. near

agrus (Walker)

Aprostocetus aurantiacus (Ratzeburg)

Aprostocetus cerricola (Erdös)

Aprostocetus clavicornis (Zetterstedt)

Aprostocetus constrictus Graham

Aprostocetus eriophyes (Taylor)

Aprostocetus escherichi (Szelényi)

Aprostocetus eurytomae (Nees)

Aprostocetus pallipes (Dalman)

Aprostocetus legionarius Giraud

Aprostocetus serratulae Graham

Aulogymnus arsames (Walker)

Aulogymnus gallarum (L.)

Aulogymnus obscuripes (Mayr)

Aulogymnus skianeuros (Ratzeburg)

Aulogymnus trilineatus (Mayr)

Baryscapus berhidanus (Erdös)

Baryscapus daira (Walker)

Baryscapus diaphantus (Walker)

Cirrospilus vittatus Walker

Paragaleopsomyia cecidobroter Gordh & Hawkins (= Tetrastichus cecidobroter)

Pediobius clita (Walker)

Pediobius lysis (Walker)

Pnigalio nemati (Westwood)

Pronotalia inflata Graham

Quadrastichus brevinervis (Zetterstedt)

Quadrastichus sajoi (Szelenyi)

Quadrastichus ventricosus (Graham)

Quadrastichus xanthosoma Graham

Sigmophora brevicornis (Panzer) (= Aprostocetus brevicornis)

Eupelmidae, chalcid wasps

Eupelmus urozonus Dalman

Eupelmus vesicularis (Retzius) (= Macroneura vesicularis)

Eurytomidae, chalcid wasps

Eurytoma aciculata Ratzeburg

Eurytoma brevitergis Bugbee

Eurytoma brunniventris Ratzeburg

Eurytoma cynipsea Boheman

Eurytoma dentata Mayr

Eurytoma gigantea Walsh.

Eurytoma hypochoeridis Claridge

Eurytoma mayri Ashmead

Eurytoma obtusiventris Gahan

Eurytoma robusta Mayr

Eurytoma rosae Nees

Eurytoma serratulae Latreille

Eurytoma tibialis Boheman

Sycophila biguttata (Swederus) (= Eudecatoma biguttata)

Sycophila submutica (Thomson)

Sycophila variegata (Curtis)

Ichneumonidae, ichneumon wasps

Ctenophora pygobarba (Roman)

Ephialtes similis Bridgman

Exeristes arundinis (Kriechbaumer)

Gelis formicarius (L.)

Lathrostizus euurae (Ashmead)

Lathrostizus lugens (Gravenhorst)

Mastrus deminuens (Hartig) (= M. castaneus)

Orthopelma mediator (Thunberg)

Scambus annulatus (Kiss)

Scambus phragmitidis (Perkins)

Scambus vesicarius (Ratzeburg)

Ormyridae, chalcid wasps

Ormyrus nitidulus (F.)

Ormyrus pomaceus (Geoffroy in Fourcroy)

Platygastridae, proctotrupoid wasps

Metaclistus phragmitidis Debauche

Platygaster betularia Kieffer

Synopeas ?nervicola Kieffer

Pteromalidae, chalcid wasps

Arthrolytus ocellus (Walker)

Caenacis inflexa (Ratzeburg) (= Habrocytus periclisti)

Caenacis lauta (Walker) (= C. divisa)

Cecidostiba fungosa (Geoffroy in Fourcroy) (= C. adana)

Cecidostiba semifascia (Walker)

Eulonchetron torymoides (Thomson)

Gastrancistrus hamillus Walker

Hobbya stenonota (Ratzeburg)

Homoporus subniger (Walker)

Leptomeraporus nicaee (Walker)

Mesopolobus amaenus (Walker)

Mesopolobus diffinis (Walker)

Mesopolobus dubius (Walker)

Mesopolobus fasciiventris Westwood

Mesopolobus fuscipes (Walker)

Mesopolobus mediterraneus Mayr

Mesopolobus sericeus (Förster) (= M. jucundus)

Mesopolobus tibialis (Westwood)

Mesopolobus xanthocerus (Thomson)

Ormocerus vernalis Walker

Pseudocatolaccus thoracicus (Walker)

Psilonotus adamas Walker

Psilonotus achaeus (Walker)

Psilonotus hortensia Walker

Pteromalus albipennis Walker

Pteromalus bedeguaris (Thomson)

Pteromalus dolichurus (Thomson)

Pteromalus elevatus (Walker)

Pteromalus vibulenus (Walker)

Stenomalina liparae (Giraud)

Stinoplus etearchus (Walker)

Torymidae, chalcid wasps

Glyphomerus stigma (F.)

Megastigmus dorsalis (F.)

Megastigmus stigmatizans (F.)

Thaumatorymus notanisoides Ferrière & Novicky

Torymus affinis (Fonscolombe)

Torymus auratus (Müller) (= T. nigricornis, = T. nitens)

Torymus bedeguaris (L.)

Torymus chloromerus (Walker)

Torymus cyaneus Walker (= Syntomaspis cyaneus)

Torymus fagopirum (Provaucher)

Torymus flavipes (Walker) (= T. auratus)

Torymus fuscicornis (Walker)

Torymus geranii (Walker) (= T. cingulatus)

Torymus sp. near

microstigma (Walker)

Torymus nigritarsus (Walker)

Torymus nitidulus (Walker)

Torymus nobilis Boheman

Torymus roboris (Walker)

Torymus rubi (Schrank) (= T. macropterus)

Torymus sinensis Kamijo

Torymus solitarius (Osten-Sacken)

Vespidae, wasps

Vespa crabro L., hornet

Coleoptera

(alphabetical order of species, with family)

Apion minimum Herbst, Apionidae

Curculio salicivorus Paykull, Curculionidae

Curculio villosus F., Curculionidae

Mordellistena convicta Le Conte (= M. unicolor), Mordellidae

Aves, birds

(alphabetical order of species, with family)

Aegithalos caudatus (L.), long-tailed tit, Aegithalidae

Carduelis chloris L., greenfinch, Frigillidae

Parus atricapillus L., black-capped chickadee, Paridae

Parus major L., great tit, Paridae

Picoides pubescens (L.) (= Dendrocopus pubescens), downy woodpecker, Picidae

Sitta europaea L., nuthatch, Sittidae

Mammalia, mammals

Sciurus carolinensis Gmelin, grey squirrel, Sciuridae

Sus scrofa L., wild boar, Suidae

APPENDIX D: OTHER SPECIES

VIRUSES

Blackcurrant reversion disease (probably a virus)

Maize streak virus (MSV)

Tobacco mosaic virus (TMV)

FUNGI

(with order and family)

Phytophthora infestans (Mont.) de Bary, potato blight, Peronsporales: Peronosporaceae

PLANTS

(alphabetical order of species, with family)

Chenopodium ambrosioides L., epazote, Chenopodiaceae

Coriandrum sativum L., cilantro, Apiaceae

Ribes uva-crispa L., gooseberry, Grossulariaceae

INSECTS AND MITES

(alphabetical order of orders and species, with family)

Acari: Eriophyoidea

Stenacis triradiatus (Nalepa), Eriophyidae

Coleoptera

Cassida rubiginosa Müller, tortoise beetle, Chrysomelidae

Cyphocleonus achates Fåhr, Curculionidae

Lema cyanella (L.), Chrysomelidae

Diptera

Culex pipiens L., gnat, Culicidae

Phytomyza ilicis Curtis, Agromyzidae

Lipara similis Schiner, Chloropidae

Paracantha gentilis Hering, Tephritidae

Terellia ruficauda (F.), Tephritidae

Xyphosia miliaria (Schrank), Tephritidae

Hemiptera

Philaenus spumarius L., cuckoo-spit froghopper, Aphrophoridae

Hymenoptera

Nematus capreae L., Tenthredinidae

Nematus salicis L., Tenthredinidae

Lepidoptera

Agapeta zoegana L., Tortricidae

Bombyx mori (L.), silkworm, Bombycidae

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CHAPTER 4

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CHAPTER 8

Brooks, S.E. & Shorthouse, J.D. (1998) Developmental morphology of the stem galls of Diplolepis nodulosa (Hymenoptera: Cynipidae) and those modified by the inquiline Periclistus pirata (Hymenoptera: Cynipidae) on Rosa blanda (Rosaceae). Canadian Journal of Botany 76: 365–81.

Freese, G. & Zwölfer, H. (1996) The problem of optimal clutch size in a tritrophic system: the oviposition strategy of the thistle gallfly Urophora cardui (Diptera, Tephritidae). Oecologia 108: 293–302.

Jones, J.T. & Perry, R.N. (2004) Plant-parasitic nematodes–small animals, big impact. Biologist 51: 155–9.

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CHAPTER 9

Askew, R.R. & Ruse, J.M. (1974) The biology of some Cecidomyiidae (Diptera) galling the leaves of birch (Betula) with special reference to their chalcidoid (Hymenoptera) parasites. Transactions of the Royal Entomological Society of London 126: 129–67.

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Borkent, A. & Bissett, J. (1985) Gall midges (Diptera: Cecidomyiidae) are vectors for their fungal symbionts. Symbiosis 1: 185–94.

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Randolph, S. (2005) The Natural History of the Rose Bedeguar Gall and its Insect Community. British Plant Gall Society.

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Schönrogge, K., Walker, P. & Crawley, M.J. (1999) Complex lifecycles in Andricus kollari (Hymenoptera, Cynipidae) and their impact on associated parasitoid and inquiline species. Oikos 84: 293–301.

Shorthouse, J.D. (1993) Adaptations of gall wasps of the genus Diplolepis (Hymenoptera: Cynipidae) and the role of gall anatomy in cynipid systematics. Memoirs of the Entomological Society of Canada 165: 139–63.

Sitch, T. A., Grewcock, D.A. & Gilbert, F.S. (1988) Factors affecting components of fitness in a gall-making wasp (Cynips divisa Hartig). Oecologia 76: 371–5.

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CHAPTER 10

Abrahamson, W.G., Blair, C.P., Eubanks, M.D. & Morehead, S.A. (2003) Sequential radiation of unrelated organisms: the gall fly Eurosta solidaginis and the tumbling flower beetle Mordellistena convicta. Journal of Evolutionary Biology 16: 781–9.

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Askew, R.R. (1961b) A study of the biology of species of the genus Mesopolobus Westwood (Hymenoptera: Pteromalidae) associated with cynipid galls on oak. Transactions of the Royal Entomological Society of London 113: 155–73.

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Askew, R.R. & Ruse, J.M. (1974) The biology of some Cecidomyiidae (Diptera) galling the leaves of birch (Betula) with special reference to their chalcidoid parasites. Transactions of the Royal Entomological Society of London 126: 129–67.

Björkman, C. (2000) Interactive effects of host resistance and drought stress on the performance of a gall-making aphid living on Norway spruce. Oecologia 123: 223–31.

Brooks, S.E & Shorthouse, J.D. (1997) Developmental morphology of stem galls of Diplolepis nodulosa (Hymenoptera: Cynipidae) and those modified by the inquiline Periclistus pirata (Hymenoptera: Cynipidae) in Rosa blanda (Rosaceae). Canadian Journal of Botany 76: 365–81.

Burstein, M. & Wool, D. (1992) Great tits exploit galls as a source of food. Ornis Scandinavica 23: 107–9.

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Gordh, G. & Hawkins, B.A. (1982) Tetrastichus cecidobroter (Hymenoptera: Eulophidae), a new phytophagous species developing within the galls of Asphondylia (Diptera: Cecidomyiidae) on Atriplex (Chenopodiaceae) in southern California. Proceedings of the Entomological Society of Washington 84: 426–9.

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Miller, D.G. (2005) Ecology and radiation of galling aphids (Tamalia: Hemiptera: Aphididae) on their host plants (Ericaceae). Basic and Applied Ecology 6: 463–9.

Mound, L.A. (2005) Thysanoptera: diversity and interactions. Annual Reviews of Entomology 50: 247–69.

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Randolph, S. (2005) The Natural History of the Rose Bedeguar Gall and its Insect Community. British Plant Gall Society.

Redfern, M. & Hunter, M.D. (2005) Time tells: long-term patterns in the population dynamics of the yew gall midge, Taxomyia taxi (Cecidomyiidae) over 35 years. Ecological Entomology 30: 86–95.

Schönrogge, K. (2001) Quantifying galls: studying the invasion of species-rich communities. Cecidology 16: 31–41.

Sitch, T.A., Grewcock, D.A. & Gilbert, F.S. (1988) Factors affecting components of fitness in a gall-making wasp (Cynips divisa Hartig). Oecologia 76: 371–5.

Shorthouse, J.D. (1993) Adaptations of gall wasps in the genus Diplolepis (Hymenoptera: Cynipidae) and the role of gall anatomy in cynipid systematics. Memoirs of the Entomological Society of Canada 115: 139–63.

Van Valen, L. (1973) A new evolutionary law. Evolutionary Theory 1: 1–30.

Williams, R. (2005) Oak Galls in Britain. Vanellus Publications, Wedmore, Somerset.

Wool, D. & Bar-El, N. (1995) Population ecology of the galling aphid Forda formicaria von Heyden in Israel: abundance, demography and gall structure. Israel Journal of Zoology 41: 175–92.

Wool, D. & Ben-Zvi, O. (1998) Population ecology and clone dynamics of the galling aphids Geoica wertheimae (Sternorrhyncha: Pemphigidae: Fordinae). European Journal of Entomology 95: 509–18.

Wool, D. & Burstein, M. (1991) Parasitoids of the gall-forming aphid Smynthurodes betae (Aphidoidea: Fordinae) in Israel. Entomophaga 36: 531–8.

Wool, D. & Manheim, O. (1986) Population ecology of the gall-forming aphid, Aploneura lentisci (Pass.) in Israel. Researches in Population Ecology 28: 151–62.

CHAPTER 11

Abrahamson, W.G., Blair, C.P., Eubanks, M.D. & Morehead, S.A. (2003) Sequential radiation of unrelated organisms: the gall fly Eurosta solidaginis and the tumbling flower beetle Mordellistena convicta. Journal of Evolutionary Biology 16: 781–9.

Cox, C.B. & Moore, P.D. (2000) Biogeography. Blackwell, Oxford. Cook, L.G. & Gullan, P.J. (2004) The gall-inducing habit has evolved multiple times among the eriococcid scale insects (Sternorrhyncha: Coccoidea: Eriococcidae). Biological Journal of the Linnean Society 83: 441–52.

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CHAPTER 13

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CHAPTER 14

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Ray, John. (1826 edition, facsimile 2005) The Wisdom of God manifested in the Works of the Creation. Ray Society, London.

Rübsaamen, E.H. & Hedicke, H. (1925) Die Cecidomyiden (Gallmücken) und ihren Cecidien. Zoologica 77: 1–112.

Senn, G. (1941) Oak galls in the Historia Plantarum of Theophrastus. Transactions of the Royal Society of Edinburgh 60: 343–54.

Stearn, W.T. (ed.) (1973) John Ray: Synopsis Methodica Stirpium Britannicum, Editio Tertia, 1724. Ray Society, London.

Swanton, E.W. (1912) British Plant-Galls. A Classified Textbook of Cecidology. Methuen & Co., London.

White, Gilbert (1813, third edition; facsimile, 1993) The Natural History and Antiquities of Selbourne. Ray Society, London.

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SPECIES INDEX

Abies alba, European silver-fir 41

Acacia ferruginea 151–2

Acacia longifolia, long-leaved wattle 464

Acacia pycnantha, golden wattle 465

Acacia tortilis 415

Acalitus brevitarsus 81, 110

Acalitus phloeocoptes 286

Acer 80–1, 124, 133–4, 240, 244, 261

Acer pseudoplatanus, sycamore 28, 124, 133–4, 244, 261

Acer saccharinum, silver maple 80

Aceria campestricola 81, 133–5

Aceria erinea 81, 108–9

Aceria filiformis 81

Aceria hindali 287

Aceria lycopersici 42–4, 81

Aceria macrorhyncha 81, 133–4

Aceria nalepai 81

Aceria nervisequa 108

Aceria populi 487

Aceria tenuis 54

Aceria tristriata 81–2

Aceria ulmi 128, 287

Aceria varia 81

Achillea millefolium, yarrow 215–16

Acleris schalleriana 322

Acroptilon repens, Russian knapweed 215–16, 461–2, 464

Aculops macrotrichus 81

Aculus convolvuli 81

Aculus hippocastani 81, 109

Aculus laevis 81

Adelges abietis 95, 162, 164

Adelges laricis 162, 164–6

adelgids 3, 7, 12, 153, 162–3, 167–70, 286, 360–1

Aegithalos caudatus, long-tailed tit 324

Aesculus hippocastanum, horsechestnut 51, 109

Aethes rubigana 302

Agaonidae 178–9, 182–4, 201, 203, 385, 400–2, 404

Agapeta zoegana 463

Agrobacterium rhizogenes 51

Agrobacterium tumefaciens 4–5, 9, 20, 24–6, 47, 49–52, 446

Albuginaceae 57

Albugo candida 57–8

Alder, see Alnus glutinosa

Aleppo oak, see Quercus infectoria

Allium cepa, onion 215, 444, 455

almond, see Prunus dulcis Alnus 26, 41, 63, 80–2, 110

Alnus glutinosa, alder 26, 41, 63, 80, 82, 110

Alophia combustella 289

Ammophila arenaria, marram grass 64

Andricus aries, ramshorn gall wasp

Andricus caliciformis, mulberry gall 469, 470–1

Andricus caputmedusae 399, 473

Andricus conficus 393, 395

Andricus coriarius 393, 394, 399

Andricus corruptrix 275, 333, 336, 391, 424–5, 429

Andricus curtisii 398

Andricus curvator 272, 273, 337–8, 393, 398, 498

Andricus dentimitratus 393, 398

Andricus foecundatrix, oak artichoke gall wasp 272, 338, 393–4, 398, 438, 474

Andricus galeatus 397

Andricus gallaeurniformis 393–4

Andricus gemmeus 275, 391, 393, 395, 424

Andricus grossulariae 250, 275, 336, 391, 393, 424

Andricus hartigi 393–4, 398, 399

Andricus hispanicus, Iberian marble gall wasp 430–1

Andricus hungaricus 393

Andricus infectorius 396, 397, 424, 435, 436, 438, 441, 467, 470–1, 476, 479

Andricus inflator 393

Andricus kollari, marble gall wasp 248, 250, 272, 274–7, 323, 332–6, 338, 342, 390, 391, 393–4, 399, 424–31, 436, 438–9, 443, 474, 479, 481

Andricus lignicolus, cola nut gall wasp 275, 333, 390–1, 396, 424–5, 429

Andricus lucidus, hedgehog gall wasp 248, 250, 275, 279, 391, 393–4, 398, 424, 469, 471, 473

Andricus malpighii 398

Andricus mitratus 398

Andricus multiplicatus 470, 473–4

Andricus paradoxus 250

Andricus polycerus 393, 395, 397

Andricus quadrilineatus 398

Andricus quercuscalicis, knopper gall wasp 13, 248, 252, 272, 275, 277–9, 331, 333, 335–7, 342, 390–1, 393–4, 396, 398, 424–9, 433, 438, 439, 466

Andricus quercusramuli, cottonwool gall 337, 429, 473

Andricus quercustozae 395, 393, 438–9, 469, 471, 479

Andricus quercustozae f. insana, Bassorah gall wasp 438, 471

Andricus schroeckingeri 398

Andricus seckendorffi 393

Andricus solitarius 398

Andricus theophrastea, woolly or cotton gall 438, 469, 472–3

Andricus viscosus (see Andricus dentimitratus)

Anemone coronaria 42

Anemone nemorosa, wood anemone 42, 61, 492–3

Anguina tritici 447

Anisostephus betulinus 235–6

Anogeissus latifolia 121

Anthocoris minki 291

Anthocoris nemoralis 291

Anthocoris nemorum 291

Anthomyiidae 114

Aphelenchoides ritzemabosi 37, 214

Aphelonyx cerricola 336, 391, 424

Aphididae 91–4, 98, 118, 137, 140, 359–62, 366, 368

Aphidiidae 289

Aphids 23, 76, 91, 93–4, 98, 359, 362, 397

Apiognomonia errabunda 342

Apiomorpha munita 138

Apion minimum 321

Aploneura lentisci 153, 290–1, 363, 364

apple (cultivated), see Malus domestica

apple leaf midge, see

Dasineura mali Aprostocetus aethiops 330–1, 342

Aprostocetus aurantiacus 328

Aprostocetus cerricola 331

Aprostocetus clavicornis 310, 312–13

Aprostocetus constrictus 310, 313

Aprostocetus eriophyes 286–8, 299

Aprostocetus escherichi 310, 313

Aprostocetus eurytomae 325, 328

Aprostocetus legionarius 296, 298–9, 304

Aprostocetus pallipes 310, 312–13

Aprostocetus serratulae 295, 298

Archips podana 319, 322

Arctium minus, lesser burdock 187–8, 191, 295

Arrhenothrips ramakrishnae 121

arroyo willow, see Salix lasiolepis

Arthrolytus ocellus 331

Asiphonella dactylonii 364

aspen, see Populus tremula

Asphondylia atriplicis 372, 405

Asphondylia aucubae 372

Asphondylia baca 372

Asphondylia gennadii 373

Asphondylia itoi 372

Asphondylia morivorella 372

Asphondylia pruniperda 238

Asphondylia sarothamni 238, 313, 314–15, 372, 373

Asphondylia sphaera 372

Asphondylia ulicis 315

Asphondylia yushimai 372

Asterodiaspis minus 126

Asterodiaspis quercicola 125

Asterodiaspis variolosa 125–7

Asterolecaniidae 96–7

Atriplex canescens 415

Aulacidea hieracii 251

Aulacidea phlomica 387

Aulacidea verticillica 387

Aulogymnus arsames 329, 341

Aulogymnus gallarum 329, 331, 337

Aulogymnus obscuripes 331

Aulogymnus skianeuros 325, 328, 330

Aulogymnus trilineatus 330–1, 332

Austrothrips cochinchinensis 136

Avena sativa, oats 67, 215

Aylax minor 253

Aylax papaveris 251, 253

Baizongia pistaciae 141, 144, 291, 365, 475

barley, two-rowed, see Hordeum distichon

Baryscapus berhidanus 331, 337

Baryscapus daira 295, 298

Baryscapus diaphantus 330, 337

basket or almond willow, see Salix triandra

bassorah gall wasp, see Andricus quercustozae f. insana

Batrachedra striolata 317–18

bay laurel, see Laurus nobilis

bay willow, see Salix pentandra

bedeguar gall wasp (robin’s pincushion), see Diplolepis rosae

beech, see Fagus sylvatica

Beesoniidae 96–7, 128, 153, 162, 169

beetles 3, 12, 23, 100, 178–9, 205–6, 237–8, 286, 292, 300, 319, 349, 352, 369, 380, 404–5, 414, 461

Berberis vulgaris, common barberry 67, 450

Betula pendula, silver birch 235

Betula pubescens, downy birch 235

bilberry, see Vacinium myrtillus

Biorhiza pallida, oak apple gall wasp 245, 248, 250, 252, 261–4, 272, 323, 328, 330, 333–9, 346, 429, 436, 438, 446, 477

bird cherry, see Prunus padus

bittersweet, woody nightshade, see Solanum dulcamara

black knapweed, see Centaurea nigra

black nightshade, see Solanum nigrum

black poplar, see Populus nigra

black-capped chickadee, see Parus atricapillus

blackcurrant big bud mite, see Cecidophyopsis ribis

blackcurrant, see Ribes nigrum

blackthorn, sloe, see Prunus spinosa

Blastobasis lignea 306, 308–9

Blennocampa phyllocolpa 99, 111

Bombyx mori, silkworm 486

box cabbage gall, see Spanioneura buxi

box, see Buxus sempervirens

bracken, see Pteridium aquilinum

Bracon discoideus 319–20

Bracon minutator 295, 300–1

Bracon picticornis 319–20

brambles, see Rubus fruticosus

Brassica oleracea, cabbage, cauliflower 21, 37, 57, 59, 145, 153–4, 157–8, 214, 363

broom, see Cytisus scoparius

brown knapweed, see Centaurea jacea

buckthorn, see Rhamnus cathartica

burnet rose, see Rosa pimpinellifolia

Buxus sempervirens, box 129–30, 157

cabbage, cauliflower, see Brassica oleracea

Caenacis inflexa 325, 327–8

Caenacis lauta 330, 332

California white oak, see Quercus lobata

Callirhytis erythrocephala 342

Calluna vulgaris, heather, ling 36, 53

Calycopteris floribundus 136

Capsella bursa-pastoris, shepherd’s-purse 57–8

Carduus acanthoides 459

Carduus nutans, nodding thistle 189–90, 196, 457–9, 460, 461, 463

Cassida rubiginosa, tortoise beetle 461

Cecidomyiidae 46, 101–2, 105, 123–4, 147, 230, 239, 294, 302–3, 335, 344, 355, 369–70, 394, 407, 410–1, 434

Cecidophyopsis psilaspis, yew big bud mite 154–7, 286–8, 306, 309

Cecidophyopsis ribis, blackcurrant big bud mite 82, 154–6, 288, 447

Cecidostiba fungosa 330–2

Cecidostiba semifascia 330–2

Celtis occidentalis, hackberry 137, 157–8

Centaurea diffusa, diffuse knapweed 206–7, 461, 463

Centaurea jacea, brown knapweed 196

Centaurea nigra, black knapweed 192–4, 196–7, 295

Centaurea scabiosa, greater knapweed 186, 242

Ceratoglyphina bambusae 292

Ceratosolen arabicus 401

Ceratosolen capensis 401

Ceratosolen galili 401–3

Ceratosolen pilipes 401

Ceroptres arator 332

chalcid wasps 3, 319, 326, 385, 405

Chenopodium ambrosioides 444

cherry gall wasp, see Cynips quercusfolii

chestnut gall wasp, see Dryocosmus kuriphilu

chestnut oak, see Quercus montana

Chirosia grossicauda 114

Chloropidae 153, 159, 294

cilantro, see Coriandrum sativum

Cinnamomum osmophloeum 127

Cirrospilus vittatus 310, 313

Cirsium arvense, creeping thistle 186, 192–3, 214–15, 223–4, 226, 295–6, 379, 419–21, 459, 460–1

Cirsium setidens 379

Cirsium vulgare, spear thistle 192, 194, 196–7, 295, 305, 377, 459–61

Claviceps purpurea 64

Clavicipitaceae 64

Cleonis pigra 461

Clinidiplosis cilicrus 310–11, 335

Coccoidea 88, 96, 123, 128, 137–8

Cocoa swollen shoot virus (CSSV) 20

cola nut gall wasp, see Andricus lignicolus

Colastes braconius 319

Coleoptera 3, 178–9, 205, 334, 352, 404

Colomerus vitis 81–2, 109–10, 287–8

common barberry, see Berberis vulgaris

common cat’s-ear, see Hypochaeris radicata

common fleabane, see Pulicaria dysenterica

common juniper, see Juniperus communis

common lime, see Tilia x europaea

common nettle, see Urtica dioica

common reed, see Phragmites australis

common spangle gall wasp, see Neuroterus quercusbaccarum

common whitebeam, see Sorbus aria Contarinia manii 151

Contarinia ramachandrani 151

cord grass, see Spartina

Coriandrum sativum, cilantro 444

cork oak, see Quercus suber

Cornus sanguinea, dogwood 487

Corylus avellana, hazel 80, 154–5, 286

cottonwool gall, see Andricus quercusramuli

couch, see Elytrigia repens

cowberry, see Vaccinium vitis-idaea

crab apple, see Malus sylvestris

crack-willow, see Salix fragilis

Craneiobia corni, dogwood rivet gall midge 487

Crataegus laevigata, midland hawthorn 92, 172

Crataegus monogyna, hawthorn 69–71, 92, 115–16, 171–2

creeping thistle, see Cirsium arvense

creeping willow, see Salix repens

creosote bush, see Larrea tridentata

Cronartiaceae 70

Cronartium ribicola 447

Crotonothrips memecylonicus 122

crown gall, see Agrobacterium tumefaciens 4–5, 9, 20, 24–6, 47, 50–2, 446

CSSV, see Cocoa swollen shoot virus

Ctenophora pygobarba 319

cuckoo-spit froghopper, see Philaenus spumarius

Culex pipiens, gnat 251

cupped spangle gall wasp, see Neuroterus tricolor

Curculio salicivorus 319, 321–2

Curculio villosus 328, 330, 334–5

Cynipidae 5, 103, 178, 184, 220, 228, 230, 239–41, 344, 355, 374, 385–6, 389, 421

Cynips agama 267

Cynips disticha 267, 397

Cynips divisa, red pea gall wasp 262, 266–7, 330, 333–4, 336, 338–9, 340–1

Cynips longiventris, striped pea gall wasp 267, 339, 341

Cynips quercusfolii, cherry gall wasp 267, 334, 341–2, 474, 489–90

Cyphocleonus achates 463

cypress spurge, see Euphorbia cyparissias

Cystiphora sonchi 129, 131

Cystococcus pomiformis 445

Cytisus scoparius, broom 28, 40–1, 230, 238, 313–15, 373

Daktulosphaira vitifoliae, grape phylloxera 88, 138–40, 141, 447, 452–3

dandelion, see Taraxacum officinale

dark-leaved willow, see Salix myrsinifolia

Dasineura crataegi, hawthorn button–top gall midge 171–3

Dasineura fastidiosa 310–11

Dasineura interbracta 310–11

Dasineura mali, apple leaf midge 116

Dasineura odoratae 119

Dasineura pustulans 124–5

Dasineura pyri, pear leaf midge 116

Dasineura ulmaria, 147–9

Dasineura urticae 146–8, 151

dewberry, see Rubus caesius

Diaparsis stramineipes 319, 321

Diastrophus kincaidii 324

Diastrophus rubi 220, 228, 243, 251, 253, 323, 328, 338

Dichrostachys glomerata 135

Didymomyia tiliaceae 147–50

diffuse knapweed, see Centaurea diffusa

Diplolepis bicolor 256, 391–2

Diplolepis eglanteriae, rose pea gall wasp 249, 252, 255, 391

Diplolepis fusiformis 329, 391

Diplolepis mayri 249, 252, 255–6, 328, 391

Diplolepis nervosa, rose pea gall wasp 249, 252, 255–6, 328, 391

Diplolepis nodulosa 260, 261, 391–2

Diplolepis polita 329, 391

Diplolepis rosae, robin’s pincushion, bedeguar gall wasp 243, 248, 249, 252, 255, 256–61, 266, 323, 325–8, 334, 338, 346, 436, 446, 475–6, 481

Diplolepis spinosa 13, 391–2

Diplolepis spinosissimae 249, 252, 256, 260, 391

Diplolepis triforma 391–2

Diptera 3, 46, 101–2, 114, 123, 159, 178, 180, 230, 251, 290–1, 294, 369–70, 372–3

Discula quercina (see Apiognomonia errabunda) 342

Disholcaspis eldoradensis, the honeydew gall wasp 445

Disholcaspis weldi 444

Distylium 92

Ditula angustoriana 306, 309

Ditylenchus dipsaci 213–14

docks, see Rumex

dog rose, see Rosa canina

dog’s mercury, see Mercurialis perennis

dogwood rivet gall midge, see Craneiobia corni

dogwood, see Cornus sanguinea

downy birch, see Betula pubescens

downy oak, see Quercus pubescens

downy willow, see Salix lapponum

downy woodpecker, see Picoides pubescens

Drisina glutinosa 124

Dryocosmus cerriphilus 397

Dryocosmus deciduus 445

Dryocosmus kuriphilus, chestnut gall wasp 13, 433, 447, 465–6

dwarf cherry, see Prunus cerasus

dwarf gorse or furze, see Ulex minor

Dysaphis plantaginea 113

eared willow or sallow, see Salix aurita

elder, see Sambucus nigra Elytrigia repens, couch 65

English elm, see Ulmus procera

Ephialtes similis 296

Epitrimerus trilobus 81, 115, 117

Eriophyes arianus 128–9

Eriophyes dichrostachia 81, 135

Eriophyes distinguendus 81

Eriophyes laevis 80–2

Eriophyes leiosoma 81, 109

Eriophyes pyri, pear leaf blister mite 128–30

Eriophyes similis 81

Eriophyes tiliae 81, 133–4

Eriophyes viburni 109

Eriophyidae 79–80

Eriosoma ulmi 118, 475

Eriosomatinae 91–2, 94, 118, 137, 140, 359–60, 362, 365–7

Erysimum cheiri, wallflower 54

Eucosma cana 302–3, 305

Eulonchetron torymoides 319, 320–1

Eulophidae 156, 286, 312–13, 316, 336, 405

Eupelmidae 338–9

Eupelmus urozonus 328, 338–9, 427, 433

Eupelmus vesicularis 324–5

Euphorbia cyparissias, cypress spurge 42, 171–2

Eupontania aestiva 381

Eupontania arcticornis 381

Eupontania collactanea 231, 233–4, 318, 319–22

Eupontania glabrifrons 381

Eupontania hastatae 381

Eupontania lapponica 381

Eupontania nivalis 381

Eupontania pedunculi 393, 321, 381

Eupontania polaris 381

Eupontania pustulator 381

Eupontania reticulatae 381

Eupontania samolad 381

Eupontania viminalis 321

European larch, see Larix decidua

European silver-fir, see Abies alba

Eurosta solidaginis 13, 186, 223–6, 228, 295–6, 299, 379–80

Eurytoma aciculata 319, 321

Eurytoma brevitergis 324

Eurytoma brunniventris 329–32, 334, 338, 340–2, 433

Eurytoma cynipsea 324

Eurytoma dentata 314–15

Eurytoma gigantea 295, 299–300, 380

Eurytoma hypochoeridis 324

Eurytoma mayri 324

Eurytoma obtusiventris 296, 298, 300, 304, 380

Eurytoma robusta 299, 304, 420

Eurytoma rosae 327, 334

Eurytoma serratulae 295, 297–8, 304

Eurytoma tibialis 295, 297–8

Eurytomidae 184, 315, 336, 401, 405

Eutrichapion scutellare 206, 208

Euura amerinae 211, 381

Euura atra 381

Euura lanatae 381

Euura lasiolepis 211, 317–19, 347

Euura mucronata 231, 234, 381

Euura testaceipes 381

Euura venusta 381

Exeristes arundinis 296, 302

Exobasidiaceae 74

Exobasidium myrtilli 8, 29, 74

Exobasidium rhododendri 8, 29

Fagus sylvatica, beech 92, 108, 147, 149, 151, 262, 360, 370, 416, 475–6, 487, 489

Ficus retusa, fig 181–3, 201–3, 400–4

field poppy, see Papaver rhoeas

field rose, see Rosa arvensis

fig, see Ficus retusa

Filipendula ulmaria, meadowsweet 124–5, 147, 149

Forda formicaria 141, 144–5, 291–2, 362

Forda marginata 141, 290

Forda riccobonii 363

Forsythia x intermedia, Forsythia 39

forsythia, see Forsythia x intermedia

Fragaria x ananassa, garden strawberry 37, 40, 214

fungi 9, 18, 26–31, 37, 54, 120, 490

Galium mollugo, hedge bedstraw 149

Galium verum, lady’s bedstraw 147

gall midges 3, 7, 10, 15, 46, 76–7, 101–7, 111, 115–17, 120, 123–4, 128–9, 135, 151–3, 173, 175–6, 178, 205, 217–18, 221–2, 230, 235–6, 239, 286, 294, 305, 310–12, 315, 319, 321, 344, 349, 355, 359, 369–73, 385, 407, 411–12, 415–16, 447, 455

gall mites 115, 406

gall wasps 3–5, 10, 15, 103, 178, 184, 202, 205, 221, 228, 230, 234, 240–2, 246, 249–53, 255, 261–2, 272, 274, 319, 321, 323, 327, 329, 334–6, 344–5, 349, 355, 357, 359–60, 369, 372, 374, 382, 385–6, 388–90, 392, 394–5, 399–400, 403, 405–6, 411–12, 421, 423–5, 429, 431–2, 434, 465–6, 475, 487, 494

Gallacoccus anthonyae 162, 167–8

Gallacoccus secundus 168

garden pea, see Pisum sativum

garden strawberry, see Fragaria x ananassa

Gastrancistrus hamillus 310–11

Gelis formicarius 331, 427

Geocrypta galii 147–9

Geoica wertheimae 141, 145, 290

germander speedwell, see Veronica chamaedrys

Giraudiella inclusa 219, 221–2, 296, 302

Glechoma hederacea, ground-ivy 146–7, 247, 253–4, 445

Glyphomerus stigma 325, 327

gnat, see Culex pipiens

goat willow or sallow, see Salix caprea

golden wattle, see Acacia pycnantha

goldenrod, see Solidago canadensis

golden–samphire, see Inula crithmoides

gorse (common or European), see Ulex europaeus

grape phylloxera, see Daktulosphaira vitifoliae

grapevine, see Vitis vinifera

great tit, see Parus major

greater knapweed, see Centaurea scabiosa

grey willow or sallow, see Salix cinerea

ground-ivy, see Glechoma hederacea

Gymnosporangium clavariiforme 69–70

Gymnosporangium cornutum 33, 69–70

Gymnosporangium juniperi-virginianae 70

Gynaikothrips ficorum 294

hackberry, see Celtis occidentalis

Hamamelis, 92–3, 366

Harmandiola tremulae 313

Hartigiola annulipes 147–50

hawthorn button-top gall midge, see Dasineura crataegi

hawthorn, see Crataegus monogyna

hazel big bud mite, see Phytoptus avellanae

hazel, see Corylus avellana

heart-leaved willow, see Salix eriocephala

heather, ling, see Calluna vulgaris

hedge bedstraw, see Galium mollugo

hedgehog gall wasp, see Andricus lucidus

Heliozelidae 209

Heliozella sericiella 209

Hemiptera 3, 22, 88–91, 93, 96–7, 118, 123–4, 153, 162–3, 170, 291, 355, 411

Heringia heringi 289, 290

Heringia senilis 289

Hessian fly, see Mayetiola destructor

Hirsutella thompsoni 288

Hobbya stenonota 330, 332

holm oak, see Quercus ilex

Homoporus subniger 324

honeydew gall wasp, see

Disholcaspis eldoradensis

Hopea ponga 169, 477

Hordeum distichon, barley, two-rowed 67

horsechestnut, see Aesculus hippocastani

Hymenoptera 3, 86, 98–9, 101, 156, 178–9, 182, 184, 220, 229–30, 240, 242, 246, 249, 251, 266, 284, 289, 381, 385, 404, 499

Hypochaeris radicata, common cat’s-ear 324

Iberian marble gall wasp, see Andricus hispanicus

Ichneumonidae 317, 320, 326

Inula crithmoides, golden-samphire 198–9

Iotatubothrips crozieri 294

Isocolus rogenhoferi (may be a synonym of I. scabiosae) 242–3

jointed rush, see Juncus articulatus

Juglans regia, walnut 82, 108–9

Juncus articulatus, jointed rush 44–5

Juniperus communis, common juniper 69–70

kermes oak, see Quercus coccifera

knopper gall wasp, see Andricus quercuscalicis

Lachnellula wilkommii 447

lady’s bedstraw, see Galium verum

large-leaved lime, see Tilia platyphyllos

Larix decidua, European larch 163–4, 166

Larrea tridentata, creosote bush 415

Lasioptera arundinis 218, 221–3

Lasioptera rubi 220

late goldenrod, see Solidago altissima

Lathrostizus euurae 317–19

Lathrostizus lugens 319

Laurus nobilis, bay laurel 113–14

Lema cyanella 461

Lepidoptera 3, 99, 178, 209, 351, 404

Leptomeraporus nicaee 324

lesser burdock, see Arctium minus

Lestodiplosis sp. 287

Lettuce big vein virus (LBVV) 20

Leucopsis grunini 289, 291

Leucopsis palumbi 289, 291

Leucopsis steinbergi 289, 291

lighthouse gall, see Rondaniola bursaria

Liothrips spp. 84–5, 135

Lipara lucens 154, 160, 296, 301

Lipara pullitarsis 160–1, 298, 304

Lipara rufitarsis 160–1, 298–9, 304

Lipara similis 159, 298–9

Liposthenes glechomae 247, 251, 253–4, 444

Livia juncorum, tassel gall 42, 44–5, 88–9

long-headed poppy, see Papaver dubium

long-leaved wattle, see Acacia longifolia

lucerne, see Medicago sativa

Lycopersicon esculentum, tomato 37, 44, 49, 57, 61

Lygothrips jambuvasi 121–2

Macrosiphoniella sanborni 288

Maize streak virus (MSV) 21

maize, see Zea mays

Malus domestica, apple (cultivated) 2, 37, 53, 70, 113, 115–17, 245, 248, 252, 262–6, 323, 330, 332, 439, 446, 471, 477–8, 488

Malus sylvestris, crab apple 116

Mangalorea hopeae 169–70

marble gall wasp, see Andricus kollari

marram grass, see Ammophila arenaria

Maskellia globosa 138

Massalongia betulifolia 235–6, 310–13

Massalongia ruber 235–6, 310, 312

Mastrus deminuens 331

Mayetiola destructor, Hessian fly 454

Mayetiola graminis 45–6

meadowsweet, see Filipendula ulmaria

Medicago sativa, lucerne 53

Megastigmus dorsalis 330–2

Megastigmus stigmatizans 331–2, 429, 430–2

Melampsorella caryophyllacearum 41

Melilotus albus, white sweet clover (white melilot) 49

Memecylon lushingtonii 121

Mercurialis perennis, dog’s mercury 61

Mesopolobus amaenus 330–2, 427

Mesopolobus diffinis 306–8, 348

Mesopolobus dubius 329–32

Mesopolobus fasciiventris 328–9, 330, 332, 341–2

Mesopolobus fuscipes 331–2, 342

Mesopolobus mediterraneus 324

Mesopolobus sericeus 328, 334, 342, 427

Mesopolobus tibialis 329–32, 341–2

Mesopolobus xanthocerus 330–2

Metaclistus phragmitidis 312

Metarhizium anisopliae 288

Metzneria lappa 295, 302–3

Microbotryaceae 73

Microbotryum violaceum 73

midland hawthorn, see Crataegus laevigata

Mikiola fagi 147–51, 370, 476, 487

Mimusops elengi 121

mistletoe, see Viscum album

Monarthropalpus flavus 129–30

Monoctonia pistaciaecola 289

Mordellistena convicta 299, 380

moths 3, 8, 12, 178, 205, 335, 349, 404–5, 433

mulberry gall, see Andricus caliciformis

Myopites eximia 198–9

Myopites inulaedyssentericae 198, 200

narrowleaf cottonwood, see Populus angustifolia

nectarine, see Prunus persica

Nematus capreae 381

Nematus salicis 381

net-leaved willow, see Salix reticulata

Neuroterus albipes, smooth spangle gall wasp 268–9, 271, 335, 337, 341, 342

Neuroterus anthracinus, oyster gall 339, 340–1

Neuroterus numismalis, silk button spangle gall wasp 268–71, 341–2, 429

Neuroterus quercusbaccarum, common spangle gall wasp 269, 271–2, 335, 337, 341–2, 438, 494

Neuroterus saliens 391, 424

Neuroterus tricolor, cupped spangle gall wasp 268–9, 271

nodding thistle, see Carduus nutans

Norway spruce, see Picea abies

nuthatch, see Sitta europaea

oak apple gall wasp, see Biorhiza pallida

oak artichoke gall wasp, see Andricus foecundatrix

oats, see Avena sativa

onion, see Allium cepa

Operophtera brumata 289

Orius minutus 291

Ormocerus vernalis 331, 337

Ormyridae 330–2

Ormyrus nitidulus 330–2

Ormyrus pomaceus 330, 433

Orseolia oryzae 454–5

Orseolia oryzivora 454–5

Orthopelma mediator 325–6

Oryza sativa, rice 221, 454–5

oyster gall, see Neuroterus anthracinus

Pachypsylla celtidisgemma 157–8

Pachypsylla celtidismamma 137

Palloptera parallela 295

Palloptera umbellatarum 295

Palumbina guerrini 289, 290

Pammene amygdalana 335

Pammene argyrana 335

Papaver dubium, long-headed poppy 253

Papaver rhoeas, field poppy 253

Paracantha gentilis 185

Paracletus cimiciformis 291–2

Paragaleopsomyia cecidobroter 316, 405

Parallelodiplosis galliperda 335

Parus atricapillus, black-capped chickadee 304, 324

Parus major, great tit 292, 321, 342

peach, see Prunus persica

pear leaf blister mite, see Eriophyes pyri

pear leaf midge, see Dasineura pyri

pear, see Pyrus communis

Pediaspis aceris, sycamore gall wasp 244

Pediobius clita 329, 331, 337

Pediobius lysis 337

pedunculate oak, see Quercus robur

Pemphigus betae 141, 145, 417

Pemphigus bursarius 142–3, 290, 292, 362, 489

Pemphigus dorocola 292

Pemphigus populinigrae 140

Pemphigus spyrothecae 142–4, 292, 367–8

pepper, black, see Piper nigrum

perennial sow-thistle, see Sonchus arvensis

Periclistus brandtii 325–6

Periclistus caninae 328

Phallothrips houstoni 294

Phanacis centaureae 249

Phanacis hypochoeridis 324

Philaenus spumarius, cuckoo-spit froghopper 8, 189

Phragmidium tuberculatum 491

Phragmites australis, common reed 153, 159–60, 218, 221–2, 294, 296

Phyllocolpa anglica 381

Phyllocolpa coriacea 381

Phyllocolpa excavata 381

Phyllocolpa leucosticta 381

Phyllocolpa nudipectus 381

Phyllocolpa puella 381

Phyllocoptes eupadi 81

Phyllocoptes goniothorax 81, 116–17

phylloxerids 138

Physemocecis hartigi 129–30

Physemocecis ulmi 129, 131

Phytomyza ilicis 8

Phytophthora infestans, potato blight 27, 28

Phytoptus avellanae, hazel big bud mite 80–1, 154–5, 286

Phytoptus bursarius 140, 142, 144, 362, 368

Phytoptus tetratrichus 81, 117–18

Picea abies, Norway spruce 95, 162, 164

Picea sitchensis, sitka spruce 164, 167

Picoides pubescens 303

Picoides pubescens, downy woodpecker 303–4

Pileolaria terebinthi 289, 292

Pinus cembra 71

Pinus strobus, Weymouth pine 70

Pinus sylvestris, Scots pine 209–10

Piper nigrum, black pepper 84, 87

Pipiza festiva 289, 290–1

Pistacia atlantica 141, 362–4

Pistacia lentiscus 141, 362–4

Pistacia palaestina 145, 292, 362

Pisum sativum, garden pea 39

Plagiotrochus amenti 391, 432

Plagiotrochus australis 432

Plagiotrochus coriaceus 432

Plagiotrochus quercusilicis 432

Plantago lanceolata, ribwort plantain 214

Plasmodiophora brassicae 27, 54–5, 57, 447

Plasmodiophoraceae 54, 59

Plasmopara viticola 288

Platygaster betularia 310, 312

Platygastridae 311–12

Pnigalio nemati 319

Poa nemoralis, wood meadow-grass 45–6

Polemochartus liparae 296–7, 298, 304

Polemochartus melas 296–8

Pontania bridgmanii 321

Pontania dolichura 382

Pontania proxima 231–4, 319–21, 382

Pontania triandrae 321

Populus angustifolia, narrowleaf cottonwood 141, 418

Populus nigra, black poplar 63, 140, 142, 145

Populus tremula, aspen 313

potato blight, see Phytophthora infestans

potato, see Solanum tuberosum

Proales werneckii 47–8

Proalidae 48

Pronotalia inflata 296

Prunus avium, wild cherry 41

Prunus cerasus, dwarf cherry 41

Prunus domestica, wild plum 64

Prunus dulcis, almond 61, 172–3

Prunus padus, bird cherry 109–10

Prunus persica, nectarine, peach 28, 31, 51, 61–3, 447

Prunus spinosa, blackthorn, sloe 8, 64, 286

Psaronius chasei 350

Pseudasphondylia matatabi 372

Pseudocatolaccus thoracicus 314–15

Psilonotus achaeus 310, 313

Psilonotus adamas 310, 313

Psilonotus hortensia 310, 313

psylloids 3, 7, 12, 43, 76–7, 88–90, 92, 98, 103, 107, 111, 113, 117, 123–5, 127–8, 132, 137, 153, 157, 286, 411

Pteriditorichnos stipitopterii (undetermined fossil holometabolous insect) 350

Pteridium aquilinum, bracken 114

Pteromalidae 183–4, 290, 307, 311, 313, 315, 337, 401, 464

Pteromalus albipennis 297–8, 301

Pteromalus bedeguaris 325, 327

Pteromalus dolichurus 319, 321

Pteromalus elevatus 295, 300–1

Pteromalus vibulenus 324

Puccinia graminis 67–8, 450

Puccinia punctiformis 461

Pucciniaceae 67, 69

Pucciniastrum geoppertianum

Pulicaria dysenterica, common fleabane 198–9

purple willow, see Salix purpurea

Pyrus communis, pear 34, 115–17, 128–9, 234

Quadrastichus brevinervis 310, 312–13

Quadrastichus sajoi 286

Quadrastichus ventricosus 310, 312

Quadrastichus xanthosoma 310, 313

Quercus bicolor, swamp white oak 495

Quercus cerris, Turkey oak 241, 248, 252, 274, 276–80, 331–2, 335, 390–1, 421, 423–5, 432–3, 471, 473

Quercus coccifera, kermes oak 475

Quercus copeyensis 412

Quercus ilex, holm oak 391, 432, 475

Quercus infectoria, Aleppo oak 436, 469–70

Quercus lobata, California white oak 445

Quercus macrolepis 443

Quercus montana, chestnut oak 126

Quercus petraea, sessile oak 125, 209, 241, 252, 263, 266, 268–9, 272–4, 277, 329–30, 332, 390, 421, 423–4, 427, 429–30, 496

Quercus pubescens, downy oak 469

Quercus robur, pedunculate oak 125, 209, 241, 248, 252, 263, 266–8, 272–5, 277–8, 329–32, 390, 421, 423–5, 427, 429–30, 489

Quercus suber, cork oak 391, 424, 429–32, 471, 475

Rabdophaga degeeri 222

Rabdophaga heterobia 172

Rabdophaga rosaria 171

Rabdophaga salicis 222–3

Rabdophaga strobiloides 172–3

Ramichloridium subulatum 218

ramshorn gall wasp, see Andricus aries

raspberry, see Rubus idaeus

red pea gall wasp, see Cynips divisa

redcurrant, see Ribes rubrum

Retinia resinella 210

Rhacochlaena toxoneura 319, 321–2

Rhamnus cathartica, buckthorn 119

Rhinanthus minor 36

Rhinocyllus conicus 179, 187, 189–90, 206, 457, 460, 464

Rhizobiaceae 50

Rhodococcus fascians 39

Rhododendron ponticum, rhododendron 74

Rhus semialata, sumach 436, 438

Ribes nigrum, blackcurrant 82, 119, 154–6, 286, 447

Ribes rubrum, redcurrant 119

Ribes uva-crispa, gooseberry ribwort plantain, see Plantago lanceolata

rice, see Oryza sativa

robin’s pincushion, bedeguar gall wasp, see Diplolepis rosae

Rondaniola bursaria, lighthouse gall 146–8, 150

Rosa acicularis 391–2

Rosa arvensis, field rose 257

Rosa blanda 261, 391–2

Rosa canina, dog rose 111, 248, 257, 259, 325

Rosa pimpinellifolia, burnet rose 260

Rosa rubiginosa, sweet-briar 257

rose pea gall wasp, see

Diplolepis eglanteriae, Diplolepis nervosa

rowan, see Sorbus aucuparia

Rubus caesius, dewberry 220

Rubus fruticosus, brambles 4, 220, 228, 243, 323, 338, 388

Rubus idaeus, raspberry 220

Rubus saxatilis, stone bramble 220

Rumex, docks 206

Russian knapweed, see Acroptilon repens

rye, see Secale cereale

Salix spp. 38, 81, 99, 101, 111–12, 171–2, 210–11, 222–3, 231–3, 316–17, 319, 321, 347, 371, 381, 384–5, 392, 412, 416–17

Salix alba, white willow 171, 231–2, 319, 384

Salix aurita, eared willow or sallow 234

Salix borealis 381, 384

Salix caprea, goat willow or sallow 234, 417

Salix cinerea, grey willow or sallow 223, 231, 234

Salix dasyclados 381, 384

Salix eriocephala, heart-leaved willow 172

Salix fragilis, crack-willow 231–3, 319, 381, 384

Salix glauca 381, 384

Salix gracilistyla 381, 384

Salix hastata 381, 384

Salix lanata, woolly willow 381, 384

Salix lapponum, downy willow 381, 384

Salix lasiolepis, arroyo willow 112, 211, 317, 347

Salix myrsinifolia, dark-leaved willow 381, 384

Salix pentandra, bay willow 211, 384

Salix phylicifolia, tea-leaved willow 381, 384

Salix polaris 381, 384

Salix purpurea, purple willow 222

Salix repens, creeping willow 231–2, 319, 321

Salix reticulata, net-leaved willow 381, 384

Salix schwerinii 381, 384

Salix triandra, basket or almond willow 172, 384, 417

Sambucus nigra, elder 115, 436, 476

Saperda populnea 206

Saphonecrus connatus 332

sawflies 3–4, 7–8, 76, 98–101, 107, 111, 178, 205, 209–11, 230–4, 286, 296, 316, 326, 346–7, 349, 351, 364, 369, 381–3, 385, 392, 412, 416–18, 488, 499

saw-wort, see Serratula tinctoria

scale insects 3–4, 7, 12, 20, 76, 88, 92, 96–8, 103, 123, 125, 127–8, 137, 162, 294, 349, 406

Scambus annulatus 319

Scambus phragmitidis 296, 302

Scambus vesicarius 319–21

Schlechtendalia chinensis 436, 438–9, 441

Scots pine, see Pinus sylvestris

Secale cereale, rye 64, 66–7, 69, 437, 447–50, 452, 454

Semudobia betulae 310–13

Semudobia skuhravae 310, 313

Semudobia tarda 310, 313

Sequoiomyia kraeuseli 370

Serratula tinctoria, saw-wort 193

sessile oak, see Quercus petraea

shepherd’s-purse, see Capsella bursa-pastoris

Shorea curtisii 162, 167–8

Sigmophora brevicornis 313, 315

silk button spangle gall wasp, see Neuroterus numismalis

silkworm, see Bombyx mori

silver birch, see Betula pendula

silver maple, see Acer saccharinum

sitka spruce, see Picea sitchensis

Sitta europaea, nuthatch 342

Slavum wertheimae 141, 144–5, 363, 365

smooth spangle gall wasp, see Neuroterus albipes

Smynthurodes betae 290, 364

Solanum dulcamara, bittersweet, woody nightshade 43

Solanum nigrum, black nightshade 61

Solanum tuberosum, potato 9, 27–8, 31, 44, 49, 56–7, 59–61, 212–13

Solidago altissima, late goldenrod 224–5, 295, 379–80

Solidago canadensis, goldenrod 13, 186, 223–5, 227, 295–6, 300, 303–4, 345, 379–80

Solidago gigantea, tall goldenrod 295

Sonchus arvensis, perennial sow-thistle 129, 131, 188–9

Sorbus aria, common whitebeam 128

Sorbus aucuparia, rowan 33, 69–70, 128–9

Spanioneura buxi, box cabbage gall 154, 157

Spartina, cord grass 64

spear thistle, see Cirsium vulgare

speedwells, see Veronica

Sphenoptera jugoslavica 206, 462–3

Spilomicrus stigmaticalis 331, 427

Spongospora subterranea 27, 56–7

Spurgia euphorbiae 171–2

Stamnophora vernoniicola 200

Stenacis triradiatus 38

Steneotarsonemus nitidus 82, 87

Steneotarsonemus phragmitidis 302

Steneotarsonemus phragmitidis 302

Steneotarsonemus spirifex 82

Stenomalina liparae 297–8, 304

stinging nettle, see Urtica dioica

Stinoplus etearchus 324

stone bramble, see Rubus saxatilis

striped pea gall wasp, see Cynips longiventris

Styrax 92–3, 292

Subanguina millefolii 215–16

Subanguina picridis 214–16, 464

sumach, see Rhus semialata

Sus scrofa, wild boar 342

swamp white oak, see Quercus bicolor

sweet violet, see Viola odorata

sweet-briar, see Rosa rubiginosa

sycamore gall wasp, see Pediaspis aceris

sycamore, see Acer pseudoplatanus

Sycophila biguttata 325, 330–2, 336–7, 341, 427, 433

Sycophila submutica 324

Sycophila variegata 336, 433

Synchytriaceae 59

Synchytrium anemones 61

Synchytrium endobioticum 27, 59–60, 447

Synchytrium mercurialis 61

Synchytrium taraxaci 61

Synergus albipes 329–30, 332, 340, 342

Synergus apicalis 329

Synergus gallaepomiformis 329–32, 333, 334, 428

Synergus nervosus 329–31, 333–4, 340

Synergus pallicornis 331–2, 334, 338, 341, 427

Synergus pallidipennis 332

Synergus reinhardi 332–4, 429

Synergus thaumacerus 329

Synergus umbraculus 330–2, 333–4, 429

Synopeas ?nervicola 310–11

tall goldenrod, see Solidago gigantea

Tamalia coweni 368

Tamalia inquilinus 368

Taphrina alni 41

Taphrina betulina 40

Taphrina deformans 61–3, 447

Taphrina populina 63

Taphrina pruni 64, 348

Taphrina tosquinetii 63

Taphrina wiesneri 41

Taphrinaceae 30, 61

Taraxacum officinale, dandelion 61

tassel gall, see Livia juncorum

Taxomyia taxi, yew artichoke gall midge 156, 171–6, 305–7, 309, 312, 347–8, 437

Taxus baccata, yew 154–6, 171–2, 174–6, 286–8, 305–6, 308–11, 347–8, 437

tea-leaved willow, see Salix phylicifolia

Tenthredinidae 99–100, 210, 230, 381

Tephritidae 178–80, 184, 190, 294, 346, 373–5, 419

Tephritis bardanae 187–90, 193, 201, 295–6, 298, 300

Tephritis formosa 188–9

Terellia ruficauda 187, 461

Terellia tussilaginis 186, 190–2, 295–6, 298, 300, 375

Tetraneura ulmi 146

Thaumatorymus notanisoides 324

Thaumatothrips frogatti 294

Thecabius affinis 145–6

Thesium humifusum 36

Thilakothrips babuli thrips 3–4, 7, 23, 76–7, 82–4, 86–7, 90, 98, 107, 111, 117, 121–4, 131–2, 135–7, 284, 286–7, 293–4, 346, 349, 357, 406

Thysanoptera 3, 82, 84–5, 120, 124

Tilia platyphyllos, large-leaved lime 133

Tilia x europaea, common lime 134

Tobacco mosaic virus (TMV) 20–22

tomato, see Lycopersicon esculentum

tortoise beetle, see Cassida rubiginosa

Torymidae 183–4, 305, 313, 339–40, 401, 405, 429

Torymus affinis 339–40, 396

Torymus auratus 341, 345

Torymus bedeguaris 325, 327

Torymus chloromerus 297, 300–1

Torymus cyaneus 337, 427

Torymus fagopirum 324

Torymus flavipes 329–30, 341, 345

Torymus fuscicornis 310, 313

Torymus geranii 330–2, 341

Torymus nigritarsus 305–8, 347–8

Torymus nitidulus 310, 313

Torymus nobilis 330

Torymus roboris 330

Torymus rubi 324–5, 328

Torymus sinensis 432, 466

Torymus solitarius 324

Torymus sp. 314

Tranzschelia anemones 492

Tranzschelia discolor 42

Trichilogaster acaciaelongifoliae 464

Trichochermes walkeri 119

Trifolium repens, white clover 39–40

Trigonaspis sp. 333

Trioza alacris 113–14

Trisetacus pini 81

Triticum aestivum, wheat 9, 31, 35, 67–9, 175, 445, 447, 450–2, 454, 460

Trotteria obtusa 314–15

Turkey oak, see Quercus cerris

Ulex europaeus, common or European gorse 36, 53, 206, 208, 315

Ulex minor, dwarf gorse or furze 53, 208

Ulmus 81, 91, 118, 128–9, 131, 133–4, 137–8, 140, 291

Ulmus glabra, wych elm 118, 131, 138

Ulmus procera, English elm 133–4

Uromyces pisi-sativi 42

Urophora affinis 192–6, 198, 377, 462

Urophora cardui 185–6, 192–3,197–8, 200, 223–4, 226–8, 295–6, 299, 304, 377–9, 419, 420–1, 445, 461

Urophora congrua 377–8

Urophora cuspidata 377

Urophora dzieduszyckii 377

Urophora jaceana 192–8, 200, 295–6, 377–9, 462

Urophora jaculata 377

Urophora mauritanica 377–8

Urophora misakiana 377, 379

Urophora nigripalpis 377

Urophora quadrifasciata 192–5, 197–8, 199, 200, 377–8, 462–3

Urophora sirunaseva 377

Urophora solstitialis 193, 378, 459

Urophora stylata 192–200, 295–6, 299, 303–5, 377–8, 459, 460–1

Urophora terebrans 378

Urtica dioica, common nettle, stinging nettle 146–8, 446

Ustilaginaceae 72

Ustilago maydis 72–3, 443

Vaccinium myrtillus, bilberry 74

Vaccinium vitis-idaea, cowberry 74

Vasates quadripedes 80

Vaucheria dilatata 48

Veronica chamaedrys, germander speedwell 214

Veronica, speedwells 206

Verticillium lecanii 288

Vespa sp. 342

Viburnum lantana, wayfaring-tree 109

Viola odorata, sweet violet 119

Viscum album, mistletoe 36, 47

Vitis vinifera, grapevine 50–1, 82, 109–10, 138–9, 287, 360, 453

wallflower, see Erysimum cheiri

walnut, see Juglans regia

wasps 2–5, 10, 15, 26, 87, 99, 103, 178–9, 181–4, 201–5, 212, 221, 228, 230, 234, 240–3, 246, 249–53, 255, 261–2, 272, wasps cont. 274, 284, 286, 292–3, 309, 319, 321, 323, 326–7, 329, 332, 334–6, 342, 344–5, 349, 355, 357, 359–60, 369, 372, 374, 382–3, 385–6, 388–90, 392, 394–5, 399–406, 411–13, 417, 421, 423–5, 429, 431–2, 434, 465–6, 475, 487, 494

wayfaring-tree, see Viburnum lantana

Weymouth pine, see Pinus strobus

wheat, see Triticum aestivum

white clover, see Trifolium repens

white sweet clover (white melilot), see Melilotus albus

white willow, see Salix alba

wild boar, see Sus scrofa

wild cherry, see Prunus avium

wild plum, see Prunus domestica

wood anemone, see Anemone nemorosa

wood meadow-grass, see Poa nemoralis

woolly or cotton gall, see Andricus theophrastea

woolly willow, see Salix lanata

wound tumour virus (WTV) 20, 22, 49–50

wych elm, see Ulmus glabra

Xanthomonas juglandis 82

Xanthomonas syringae (see Pseudomonas syringae)

Xenodiplosis laeviusculi 335

Xestophanes potentillae 249

Xyphosia miliaria 187

yarrow, see Achillea millefolium

yew artichoke gall midge, see Taxomyia taxi

yew big bud mite, see Cecidophyopsis psilaspis

yew, see Taxus baccata

Zea mays, maize 20–1, 31, 36, 72–3, 443–4

SUBJECT INDEX

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acaricides 38

acetic acid 13, 52

achene 188, 191–2, 195–6, 198–200, 298, 302–3, 376–7

Adelgidae, conifer woolly aphids 91–4, 98, 162, 163–7, 288, 359–61, 366

Adler, Hermann 250, 488, 494–9, 514, 525

Africa 4, 20, 97, 102, 129–30, 180, 193, 200, 263, 266, 269, 374, 376, 390, 403–5, 409, 412–14, 421, 429–30, 433, 447, 454–5, 457, 464–5, 472, 474

agamic, see asexual reproduction

Albuginales 27–8

Aleppo galls 436, 438–43, 445, 470, 474, 476–7

alkaloids 67, 437, 447

alleles 422–3, 425

allopatric speciation 358–9

allozymes 422, 501

almond and peach leaf curl 61

Alps 416, 420–1

alternation of generations 359, 388, 494

amber 370, 386

ambrosia galls 103–6, 213–14, 218, 220–1, 230, 235–6, 237, 238, 305, 313, 371–2, 405, 407

Andricus galls 272, 394–5, 399

androphore 246, 265

angiosperms 4, 36, 80, 100, 241, 370–1, 381

apical bud 37, 157, 160, 174, 215, 227, 261, 392

anthers 35, 59, 73, 277

de Bary, Anton 451

apterae 95, 498

Arctic, the 99, 316, 382, 413, 416

Aristotle 257, 468, 475–6, 482

arrhenotoky/arrhenotokous 43, 80, 83, 86, 101, 143, 229, 242–4, 249, 250–2, 256, 260, 266

artichoke galls 107, 156, 171, 173–5, 272, 274, 305–6, 308, 309, 437–8, 348, 469, 474, 486

Ascomycota 27–8

asexual reproduction 25, 30, 59–60, 66, 93, 167, 218, 237, 242–6, 249–52, 261–9, 271–7, 279–80, 323, 329–39, 341, 345, 391, 429, 453, 470, 472, 494–5, 501, 504, 506, 537

agamic 336, 387, 390–1, 393–6, 398–9, 424–6, 428–30

atmospheric nitrogen 2, 52–3

Australia 4, 83–4, 88, 97, 99–100, 102, 125, 127–8, 138, 213, 216, 294, 343, 390, 400, 403–5, 409, 414, 416, 445, 447, 453, 457–9, 464

autoecious lifecycle 32, 360–2

auxins 13, 24–5, 52, 56

IAA (indole acetic acid) 13, 52

axillary bud 136, 141, 227, 234

aylacine galls 323–4

bacteria and phytoplasmas 23–4, 39

bacterial galls 3–4, 9–10, 12, 25–6, 38

bacterial nodules 2

Balkans 241, 390, 421, 424, 426, 430, 433, 471

Bassett, H. F. 250, 495–6

Bauhin, Caspar 447, 471

bean galls 128, 178, 231–2, 321, 326, 382

bedeguar gall (robin’s pincushion) 248, 255–7, 259–60, 323, 325–8, 334, 346, 436, 445, 446, 475–6, 481

beetle galls 205–6

beetles 3, 12, 23, 100, 178–9, 205–6, 237–8, 286, 292, 300, 319, 349, 352, 369, 380, 404–5, 414, 461

galls 205–6

Belgium 425–6

beneficial galls 26, 466

big buds v, 7, 76, 80, 82, 153–7, 286–8, 309

biogeographical regions 409

biological control 51, 189, 193, 206, 215, 433, 435, 447, 455–6, 460, 462–6

of weeds 435, 455

biology of aphids 91

of eriophyoid mites 79

of fig wasps 182

of gall midges 102

of gall wasps 240

of nematodes 213

of plant viruses 21

of psylloids 89

of sawflies 99

of scale insects 96

of tephritid flies 180

of thrips 84

biotroph 502

birch leaf gall midges 235, 310

bivoltine 373

black stem rust 67, 450

blister galls v, 6, 7, 9, 30, 32, 58, 61, 71, 76, 82, 101, 123–4, 128–9, 130–1, 235–6, 269, 342

bloodwood apples 445

blue-green bacteria, see cyanobacteria

boreal forests 418, 502

bottleneck 422, 502

bottom-up effects 283, 347

bracts 41, 164, 179, 187–9, 191, 193–4, 200–1, 302, 394, 459

Brazil 97, 405, 414

Britain ii, ix, 3, 36–8, 41–2, 45, 47, 53, 56, 59, 61, 67, 69, 73–4, 80–1, 83, 93, 97–9, 104, 125, 127, 137, 140, 145, 151, 157, 160, 180–1, 184–6, 190, 193, 198, 201, 206–7, 209, 213, 215, 222–3, 226, 232, 235, 238–9, 241, 244–5, 249–50, 253, 255–7, 261, 263, 266–9, 272, 274–7, 279, 288–90, 311–12, 319, 321, 324, 326–8, 330, 333, 336–8, 343, 345–6, 374, 390–1, 419–21, 424–30, 432–4, 436–7, 440, 442–4, 447, 449, 451–2, 474, 479, 481, 494, 497

brooming 4, 37, 39, 82

buds v, 2–3, 5, 7–10, 37–41, 43–4, 76–7, 80, 82–5, 88, 99–101, 103–6, 108, 110, 112, 117, 119, 130, 132, 141, 151, 153–8, 162, 166, 169–70, 173–4, 178, 187, 194, 210, 214, 216, 218, 220, 224, 228, 230–1, 233–5, 237–9, 246–8, 253, 255–7, 259–60, 262, 265–7, 270, 273, 276–80, 286–8, 305, 308–9, 316, 318, 338, 347, 357, 363, 371–3, 375, 383–4, 390, 392–3, 416–18, 426, 428–9, 464–5, 484, 487, 496

Caesalpinus 490

callus 8, 109, 130, 179–80, 185, 187–90, 192, 197, 201, 206–7, 209–211, 227–8, 234, 296, 351, 374, 404, 458, 463

cambium 2, 10, 50, 71, 149, 210, 221, 227, 247, 265

canker 70, 447,

caterpillars 4, 8, 12, 77, 78, 98, 99, 111, 122, 197, 209–10, 218, 232, 284–5, 290, 292–4, 301–3, 305, 308–9, 311, 313, 316–17, 319–23, 331–2, 335–6, 342, 345, 351, 374, 379, 404–5, 414, 438, 463

cell enlargement 74, 129

cell multiplication 118, 125, 129

cell vacuole 12, 109, 502

Central America 37, 411, 415, 444

cereals 6, 31, 35, 50, 64, 66–7, 82, 447, 450, 452, 454

changing ranges of galls and gall insects 418–34

chelicerae 79, 83, 109

China 241, 389, 391, 416, 432–3, 436, 439, 442–3, 465–6

chitin 27, 29

chloranthy 18–19

chloropid galls 153, 161, 292, 295–301

chloroplast 12, 49, 54, 172, 175

chromosomes 24, 93–4, 243, 422

Chytridiales 27–8, 355

Chytridiomycota 27–8

cigar galls 76, 153, 159, 302

caused by Lipara species 159

clades 364, 376, 382–3, 388–9, 398–9, 402

cladograms 352–6, 363–4, 372, 375–8, 381, 386–7, 398, 400–1, 407

Agaonidae 400

Andricus spp. 398

Aylacini and Synergini 387

Asphondylia spp. 372

Cynipini 386–7

figs and fig wasps 401

Myopitini 375–6

Nematini 381

pistacia aphids 363

Salix spp. 384

Tephritinae 375–6

Urophora spp. 376–7

classification and cladistics 352–3

of aphids 91

cleptoparasite 82, 284, 287

climate 4, 8, 10, 14, 43, 44, 52, 86, 90, 108, 146, 169, 283, 347, 408–10, 412, 414, 420, 427, 428, 430

climate change 1, 420

cline 425

coevolution 182, 359, 388, 399–403, 406

collenchyma 126, 134, 164–5

companion cells 39, 50

conifers 6, 36, 50, 89, 93, 100, 105, 350, 360–1, 370–1, 408, 411

conserved evolution 399

convergent evolution 356, 399

coppicing 4

Costa Rica 405, 410–12

covering galls 7

crown gall 4–5, 9, 20, 24–6, 47, 50–2, 446

cultivated plants. 42, 50, 54, 57, 61, 71, 116, 204, 257, 437, 447, 454, 491

cuticle 62–3, 103, 109, 120, 124, 175, 216, 263, 271

cyanobacteria 3

cyclical parthenogenesis 94, 243, 249, 360, 387

cylinder-piston galls 151–2

cynipid galls 5, 316, 323, 439, 442

cytokinin 13, 24, 25, 56

cytoplasm 12, 22, 24, 29, 55–6, 59–60, 62, 68, 135, 215, 225, 227, 252

Darwin, Charles 234, 493, 497–9

natural selection 379, 494, 497–8

defence of gall 86–7, 368

dehiscence 150–1

Denmark 411, 420, 423

de Réaumur, René Antoine Ferchault 488

Derham, William 488, 499

desiccation 30, 67, 88, 109–10, 137, 357, 397

detritivores 284

Devon 48, 425–6

dicotyledons (dicots) 4, 6, 42, 50, 89, 240, 408

differentiation 10, 11, 20, 50, 61, 111, 113, 118, 169, 172, 206, 225, 237, 255, 258, 372

dikaryotic 29, 32–6, 62, 69, 72–3

dioecious 182, 203, 401–2

Dioscorides 476, 478

diploid 27, 32, 34, 56, 59, 62–3, 66–7, 69, 72, 80, 83, 86, 93, 98, 101, 110, 143, 232, 242–5, 249, 251, 266

Diplolepidini 228, 240–4, 249, 251, 255, 325, 388–90

dispersal 30, 44, 51, 63, 71, 73, 81, 86, 97–8, 110, 117, 126–7, 134, 139, 145, 156, 169–70, 202, 288, 368, 379, 416–7, 422, 457

distal 239, 471

distribution of galls x, 344, 418, 488

DNA (deoxyribonucleic acid) 5, 21, 24–5, 51, 240–1, 248, 354–6, 363, 369–72, 381, 388–9, 398, 400–1, 407, 422, 430, 434, 453

dormancy 31, 34, 36, 39, 44, 56, 59, 65, 72, 74, 119, 130, 132, 155, 164–6, 174, 216, 239, 250, 262, 264–5, 267, 273, 276–7, 337, 417, 458

dormant bud 164–5, 277

drugs 435, 437

ectoparasitoid 284–5, 300, 302, 304, 313–15, 318, 324, 338–9, 380, 420

eggs 7, 43, 45, 77, 79–80, 86, 90, 93–4, 97–9, 101, 105, 111–12, 114, 120, 130, 139, 144, 161, 165, 173, 176, 178, 181, 188, 190–1, 195, 202–3, 209, 211, 217, 224, 227, 231–4, 239, 243, 247, 252, 255, 258–9, 261, 264–5, 270, 278, 296–302, 306–8, 310, 312, 315–18, 321–2, 325–8, 335, 337, 339–40, 342, 346–8, 359–62, 364, 378, 382–5, 392–3, 396, 417–18, 454, 459, 482, 484–5, 488, 490, 493, 496, 499

endoparasitoid 284, 299–300, 317, 319, 326, 328, 337, 341, 380, 427

endophytic 237

endopterygote 86, 351

endosperm 13, 191, 202, 503

enemies vi, 1–2, 8–9, 14–15, 132, 237, 283, 291–4, 303, 316, 324, 329, 332, 340–8, 357, 359, 365, 367, 373, 379–80, 395–7, 400, 406, 419, 426, 428–9, 454–7, 466

England 61, 69, 72, 114, 120, 125–6, 190, 199, 257, 277, 279, 296, 319, 324, 327, 347, 420, 424–6, 432, 442, 444, 446, 450, 467, 479, 488

epidermis 7, 30, 32–4, 45–6, 58, 68–9, 71–2, 74, 78, 87, 109, 112–14, 116–19, 121, 124, 127, 129–32, 135, 146, 150, 155, 173, 215–16, 218–19, 221, 225, 232, 235, 247–28, 255, 258, 263, 270, 276, 367, 498

ergot 64–7, 450

ergotism (ergot disease) 9, 64, 437, 446–50

erinea/erineum 7–9, 76, 78, 80–2, 107–10, 116, 123, 132, 286–8

eriophyoid mite galls 286

eukaryotic 507

Europe ix, 4, 71–2, 84, 99, 101, 114, 126, 130, 140, 144, 157, 161, 167, 185–6, 189–90, 192–3, 196, 206–7, 221–2, 226, 240–1, 244, 251–2, 255–7, 261, 263, 266, 269, 272, 276, 279, 290–1, 301, 321, 323–8, 331, 337, 342–3, 345, 362, 369, 373, 376, 389–90, 397, 408–9, 413, 419–21, 423–5, 428–33, 436, 439–40, 442–4, 447, 450–4, 457–9, 461–2, 465, 471–5, 477, 494, 497

Evelyn, John 476

evolution of galling 15, 356, 358

parsimonious 353, 355

extra-intestinal digestion 105

Fabricius 488

fasciation 4, 212, 214, 503

feeding 43, 77, 102, 109, 115, 120, 180, 184, 247, 375

chewing 14, 51, 178, 181, 206, 208–9, 242, 248, 297, 315, 351

sap-sucking 93, 181, 185, 355

sucking 12, 15, 24, 78–9, 86–7, 90, 93, 106, 132, 178, 181, 185, 213, 242, 309, 355

fighting in aphids 367, 369

fig wasps 2, 26, 178–9, 181–4, 201, 203–4, 349, 357, 359, 385, 400–6

filz galls 7

Finland 234, 418, 420

florets 35, 64–6, 179, 181–3, 185–8, 191, 194, 196, 198–204, 212, 214, 305, 375, 378, 401–3, 492

flower head galls v, 178–9, 185, 200, 296, 302, 369, 379, 421

flowering plants 4, 6, 15, 84, 101, 103, 105, 183, 241, 355, 360–1, 370, 381, 408, 411

flowers 2–3, 9–10, 13, 19, 31, 35, 37–9, 44, 50, 54, 57, 59, 62, 66–7, 73–4, 84–5, 101, 120, 130, 148, 154, 156, 179, 185, 199, 201, 204, 208, 212, 218, 238, 246, 278, 300, 346, 371, 380, 390, 393, 401–2, 428, 462, 468

nectaries 371

fluctuation in populations 347, 348, 427

folds v, 6, 9, 76, 78, 82, 88–90, 99, 101, 107–8, 110–14, 116, 120, 123, 230, 238, 280, 288, 293, 316

food 1–2, 10, 12, 14, 43–4, 49, 85, 87, 94, 99–101, 103–4, 106, 108–9, 112, 114, 119, 160, 163, 179, 183, 185–7, 190–1, 193–5, 197–8, 202, 206, 217, 226, 237, 239, 283–6, 288–9, 292–3, 295–7, 302–3, 305–6, 309–10, 313–14, 317–21, 323–5, 327–32, 335–8, 341–4, 347, 357, 359–361, 374, 377, 396, 403, 414, 426–9, 438, 443–5, 463, 497, 499

food chains 283

bottom-up effects 283, 347

top-down effects 283

food webs 285–6, 296, 303, 305–6, 310, 314, 323, 328, 331, 336, 343

Andricus kollari 332

Andricus quercuscalicis 331

Asphondylia sarothamni 314

big buds of Cecidophyopsis psilaspis 288

Biorhiza pallida 330

Cynips divisa 330

Diplolepis rosae 325

Eupontania collactanea 319

Eurosta solidaginis 295

Euura lasiolepis 317

Lipara lucens 296

Massalongia betulifolia 310

Neuroterus quercusbaccarum 329

Pemphigus gall aphids 289

Phanacis hypochoeridis 324

pistacia gall aphids 289

Pontania proxima 319

Semudobia betulae 310

Taxomyia taxi 306

Tephritis bardanae 295

thrips galls 293

Urophora cardui 295

Urophora stylata 295

fossil galls and the oldest galls 349

France 45, 120, 370, 420, 430–1, 445, 447, 449–50, 452–3, 459, 462, 465, 472–3, 477

frass 188, 197, 207, 209, 232–3, 300, 302–3, 309, 351

frond 114, 350

fundatrix(-ices) 86, 87, 94–5, 109–10, 119, 129, 130, 132, 134–5, 137–9, 141–2, 144–6, 156, 162–5, 170, 288–90, 294, 361–5, 367–8, 418

fungal galls ix, 3–4, 9, 26, 31, 64, 74, 414, 446

fungi ix, 9, 18, 26–31, 37, 54, 120, 490

anamorph 218, 342

fruitbody 30

fungicides 56, 59, 64, 67

galls ix, 3–4, 9, 26, 31, 64, 74, 414, 446

virescences 40

fynbos 412–15

Galen 478, 481

Gallacoccus galls 167, 170

gall community 286, 293–4, 299, 300–1. 305, 313, 316, 318, 323, 324, 328, 343–5, 432, 488

gallicolae 165–8, 288

gall midges 3, 7, 10, 15, 46, 76–7, 101–7, 111, 115–17, 120, 123–4, 128–9, 135, 151–3, 173, 175–6, 178, 205, 217–18, 221–2, 230, 235–6, 239, 286, 294, 305, 310–12, 315, 319, 321, 344, 349, 355, 359, 369–73, 385, 407, 411–12, 415–16, 447, 455

galls 4, 178, 305, 411

leaf, bud and fruit galls 235

pouches 146

stem galls 217

gall mites 13, 43, 76–8, 90, 107–09, 111, 115, 123–4, 134, 152, 155, 288, 349, 357, 406, 411, 418, 446

gall populations 260

galls

aylacine galls 323–4

bean galls 128, 178, 231–2, 321, 326, 382

in biological control 51, 189, 193, 206, 215, 433, 435, 447, 455–6, 460

bud and rosette galls 7

classification systems for galls 6

classification based on gall shape 7

covering galls 7

crown gall 4–5, 9, 20, 24–6, 47, 50–2, 446

cylinder-piston galls 151–2

development of 12–13

distribution of x, 344, 418, 488, 538

dyes 2, 439, 467

evolution of galls 349–407

of Exobasidium 74

as food 443–5

in fruits v, 3, 9–10, 13, 19, 37, 40, 50, 54, 62, 64, 67, 70, 84, 101, 106, 108, 170, 178–82, 201, 218, 230, 246, 250, 253, 310–11, 371, 373, 390, 405, 428, 438, 445, 468, 471, 477

gall wasp galls 323

gall wasp leaf, bud and fruit galls 239

growth and development 9, 11–12, 174

hedgehog galls 7, 248, 275, 279, 393

histioid galls 6–7

huitlacoche 444

ink 2, 275, 391, 425, 439–42, 467, 471, 476, 477, 481, 487, 500

on juniper 69

knopper galls 248, 252, 275, 277–9, 331, 333, 336–7, 342, 396, 424–5, 427–9, 433, 438–9, 466

mycoplasma galls 24

nematode galls 212

parasitic plants 18, 20, 36–7, 47, 53–4

psyllid shoot tip galls 157

rose galls 244, 248, 256, 324

rosette galls v, 7, 9, 76, 82, 101, 107, 111, 153–4, 171–3, 190, 207–8, 217, 235, 462

shoot tip galls 157

spangle galls 7, 267–71, 335, 337, 348, 438, 494

spindle galls 7, 69, 261, 296

spiny galls 7, 34, 36, 69, 169, 170, 179, 248, 391, 393–4, 398–9

stem galls 13, 185–6, 200, 205, 210, 217–18, 220, 223, 228, 235, 260–1, 296, 316–17, 347, 370, 376, 379, 383, 454

sweet galls 445

tephritid galls 179, 184–5, 294, 295–301

Theophrastus 469

thistle galls 297, 300, 346, 445, 178–9, 186–7, 189–94, 196–7, 206, 214–15, 223–4, 226–7, 252, 295, 296–9, 300–1, 304–5, 374, 377, 419–20, 445, 457–61, 481

three-way galls 369

types of 6

virus galls 49

gall wasps 3–5, 10, 15, 103, 178, 184, 202, 205, 221, 228, 230, 234, 240–2, 246, 249–53, 255, 261–2, 272, 274, 319, 321, 323, 327, 329, 334–6, 344–5, 349, 355, 357, 359–60, 369, 372, 374, 382, 385–6, 388–90, 392, 394–5, 399–400, 403, 405–6, 411–12, 421, 423–5, 429, 431–2, 434, 465–6, 475, 487, 494

rose gall wasps 252, 255, 389, 400

stem galls 228

genetic bottleneck 422

genetic tumours 4–5

genotype 357

geological time scale 349, 351

Gerard, John 446, 477

Germany 250, 265, 370, 420, 424–6, 450, 454, 460, 494–6

Gironde river 430

glacial periods 431

interglacial 420–1, 429, 431

glacial refuge 421, 424, 426, 430–1

global gradients 409

Gondwana 404–5

grapevine phylloxera 88, 138–9, 141, 447, 452–3

Greece 414, 420, 433, 462, 468, 474–5

gregarious parasitoid 298, 313, 337

ground ivy gall wasp 254–5

growth hormone 50

gynandrophore 245, 266, 504

gynephore 244, 245, 246, 265

haplodiploidy 79, 86, 101, 232, 242

haploid 27, 29, 32–6, 56, 59, 61–3, 65–7, 69, 72–4, 80, 83, 86, 93–5, 98, 101, 143, 242–5, 249, 251, 266

Harvey, William 482, 484

haustoria 29–30, 32, 35–6, 54, 57, 59, 62, 74, 237

hedgehog galls 7, 248, 275, 279, 393

herbal medicine 1

herbivore/plant relationship

heteroecious/heteroecy 32, 94, 98, 163, 243, 246, 249, 263, 272, 274, 279, 360–2, 390–1, 398, 423–5, 429–32

heterogony 94, 243, 249–50, 253, 387–8

Himalayas 416, 432

Hippocrates 435, 468, 490

histioid galls 6–7

Holarctic 389, 409

holocycle 94, 142, 164, 362–3

honeydew 33, 66, 68, 88, 99, 142, 145, 167, 181, 189, 232, 242, 291, 294, 365–7, 397, 445

Hooke, Robert xi, 490–2

host plants xi, 5, 8–9, 14, 18, 20, 26, 31–3, 35–6, 49, 59, 80, 84, 88, 91, 94, 98, 105, 112, 129, 142–3, 154, 156, 163, 166, 179, 186, 196, 198, 209, 212, 214, 217–18, 237, 240, 242, 246, 257, 260, 324, 345, 357, 359–60, 363–4, 369, 373–4, 379–81, 386–9, 399, 408, 411–12, 414–15, 418–21, 444, 450–1, 469–70, 488

host species 14, 18, 35, 42, 50, 61, 92, 94–5, 112, 165, 181, 185, 192, 215–16, 224, 252, 298, 333, 358–9, 361–2, 364, 372–3, 378–80, 384, 392–3, 401, 406, 417

host switching 183

housekeeping 98, 142, 145, 365

huitlacoche 444

humidity 43–4, 108–9, 120, 292

hyperparasitoid 284, 290, 307, 315, 319, 321

hyperplasy 109, 118, 129–30

hypertrophy 108, 129

hyphae 4, 27, 29–30, 32–5, 42, 47, 55, 57, 59, 62, 65–6, 68–9, 71–2, 74, 83–5, 103, 218–19, 221, 235, 237–9, 315, 371–2

Iberia 421, 423–4, 426, 429–32

idiobiont 285, 318

Ignis Sacer 447–8

indole acetic acid (IAA) 13, 52

inflorescence 43–4, 46, 67, 72, 104, 179, 182, 201, 203, 402–3, 492

initiation 11, 135, 168–9, 369, 371

ink (gall) 2, 275, 391, 425, 439–42, 467, 471, 476–7, 481, 487

iron–gall ink 440–1

Inner Hebrides 426

inquiline 82, 156, 237, 261, 284, 293, 299, 305, 309, 313–14, 316, 321, 323, 325–9, 333–4, 336–8, 340–2, 344, 346, 359, 368–9, 371, 380, 389, 405, 428–9, 433, 466

insect mouthparts

jaws 79, 181, 184, 201, 206, 209, 246, 260, 300, 321

labium 90

mandibles 90, 97, 103, 242, 247, 307, 339, 353

maxillae 90, 97

piercing 29, 44, 77–9, 82–3, 90, 137, 178, 194, 209, 213, 215, 234, 238, 254, 258, 292, 306, 309, 312, 315, 322, 355, 383

proboscis 12, 199–200

stylets 44, 78–9, 82–3, 85–7, 90, 93, 97, 109, 126–7, 129, 132, 137–8, 142, 145, 159, 163–5, 167, 170, 188, 213, 292, 294, 366

insects, lifecycle of, see egg, larvae, nymphs, pupae, puparium, instar

insectivores 285

instar 5, 43, 46, 77, 90, 97, 101, 107, 112–13, 119–20, 126–8, 145, 149, 151, 158–60, 163, 165, 167, 169–70, 173, 175, 178, 181, 188–9, 191, 194–7, 199–200, 209, 219, 224–7, 233, 236, 239, 260, 288, 291, 297–8, 300–1, 306–8, 317, 321–2, 365–7, 373

intercellular 5, 12, 29, 498

internodes 37, 39, 42, 82, 159–60, 215, 218, 221, 261, 454

Iran 142, 263, 269, 389, 421, 430, 471, 473–4

iron–gall ink 440–1

Israel 288–92, 361–3, 414, 426

Italy 234, 241, 390, 410, 420–1, 430, 433, 451, 465–7, 471, 474, 481, 485, 488

Japan 175, 292, 373–4, 391, 410–11, 416–17, 432–3, 439, 442, 465–6

knapweeds 178, 192–7, 206–8, 215–16, 242, 295–6, 298, 301, 303, 374, 461–2, 464

knopper galls 248, 252, 275, 277–9, 331, 333, 336–7, 342, 396, 424–5, 427–9, 433, 438–9, 466

koinobiont 285, 297, 312, 317, 323, 326

Korea 379, 391, 433, 465–6

lammas leaves 271

larvae 5, 7–8, 10–13, 14, 44, 46, 77, 79–80, 83, 86–7, 90, 98–103, 105–7, 111–12, 114–15, 120, 123–4, 129–31, 146–52, 154, 160–1, 171–6, 178–9, 181, 185, 187–91, 194–200, 202, 207–11, 213–14, 217–29, 231–7, 239, 242, 247–8, 253–5, 258–9, 261, 265, 267, 270–6, 278, 280, 284–7, 289, 296–305, 307–15, 317–18, 320–2, 324, 326–8, 333–41, 343, 345, 351, 359, 371–2, 377, 380, 382–3, 394–7, 403–4, 407, 427–9, 455, 463–4, 487–8, 499

leaf fold and roll galls 110–22

leaf initials 155–6, 158, 164, 169, 174

leaflet 115, 141, 145, 151, 258, 261, 292, 363–5

leaflets 111, 115, 135, 141–2, 145, 152, 256, 258–60, 290–1, 365, 392

leaf mines 8, 336

leaves 3–5, 7–10, 13, 18–19, 32–3, 35, 38–44, 46, 49, 54, 56–7, 61–4, 67–72, 74, 76–8, 80, 82, 84–9, 92, 97, 99–101, 103–6, 108, 110–13, 115–17, 119–21, 123–4, 129–30, 132, 134–6, 138–9, 141, 144–6, 148, 150–8, 160–3, 165, 167–9, 171–5, 178, 181, 190, 206–7, 210–12, 214–15, 221–2, 224, 227–8, 230–6, 239, 244, 246–7, 250, 253–7, 261, 266, 268–72, 276–8, 284, 287–8, 290–1, 296, 298, 302–3, 308–10, 313, 316–19, 328, 338, 340–3, 357, 360–2, 365, 368, 383–4, 390, 392–3, 404–5, 414–15, 417–18, 428, 437–8, 451–4, 462, 464, 470, 473, 476, 484, 489–92, 495

Lebanon 426

lenticels 57

lerps 88, 505

lethal inquiline 325

lichens 3

lignin 109, 134, 150, 219, 222

Linnaeus 234, 445, 488–9

Lister, Martin 484, 487

Magnus 477

Maize smut disease 72–3

Malpighi, Marcello 1, 118, 234, 467, 475, 483–8, 492–3, 498–9

marble galls 1, 124, 274–7, 323–3, 332–4, 336, 342, 425–6, 429–431, 436, 438–40, 443, 470–1, 474, 479, 481

mark galls 7

Mathiolus 482

maturation 11

medicines 435–6, 476, 487

Mediterranean 114, 142, 263, 274, 360–1, 412–16, 420–1, 439, 441, 459, 471, 473

meiosis 34, 36, 56, 59, 61, 63, 66–7, 72–3, 93, 243, 251

meristem 136, 153, 155–6, 160–2, 164, 167–9, 171–6, 215, 224, 226–7, 261, 270, 274, 297, 392

mesic 413–14, 463

mesophyll 59, 62, 68, 87, 90, 93, 109, 112, 114, 116–18, 120–1, 124, 129–32, 134–5, 146, 150, 155, 172–3, 175, 232, 255

microorganisms 4, 20

microsatellites 422

middle lamella 24–5, 146, 151, 219

midrib 42, 116, 121–2, 141–2, 145, 147–8, 151, 162, 167, 175, 205, 209–10, 232, 236, 268, 273, 311, 333, 383–4, 394, 470, 474, 495

migration 94, 166, 218, 362, 421, 429

mite and insect virescences 37, 42

mitosis 505

Mivart, St George 497

modes of evolution 358

molecular taxonomy 354, 371

monocots 6, 89, 408

monocotyledons 42, 50

monoecious 95, 142, 182, 203, 401–3

monophagous 298, 373

monophyletic 240–1, 352, 356, 360, 363, 372, 375, 382–3, 386, 388, 401–3

morphs 84, 86–7, 95, 98, 137, 164, 166, 365

moths 3, 8, 12, 178, 205, 335, 349, 404–5, 433

moth galls 209

mRNA 21–2, 25

multilocular 498

multinucleate 35, 212

multivoltine 373

mutualism 2, 182–3, 203, 214, 218, 357, 401–3

mycangia 219, 238, 372

mycelium 27, 34, 36, 41, 57, 62, 66, 68–71, 74, 106, 219, 315

mycoplasma galls 24

mycorrhizas 29

Myxomycota 27–8

natural selection 379, 494, 497–8

Nearctic 241–2, 246, 369, 374, 389–90, 408

nematodes 9–10, 12, 14, 18, 22–3, 37, 51, 76, 178, 205, 212–14, 216–17, 285, 349, 408, 446, 456

nematode galls 212

neoteny 97

neotropics 105, 412

Nepal 240, 432, 465

Netherlands 311–12, 424–6

nitrogenase 53

nitrogen fixing 2–3, 52–3

nodes 46, 218, 221, 222

nodule 12–13, 25–6, 52–3

North America 4, 71, 84, 99, 117, 144, 157–8, 167, 185–6, 189, 223–4, 226, 241, 244, 251, 255, 257, 269, 323–4, 343, 345, 362, 368–9, 374, 376, 379, 382, 389–90, 392, 408, 410–11, 413–4, 419, 433, 439, 451–4, 458–60, 494–5

nutrient sink 10–12, 123, 145, 154, 163, 172, 185, 190–1, 195, 197, 212, 219, 228, 247–8, 265, 305, 343, 365, 374, 378, 383, 414, 459–60, 462, 464

nutritive cells 10–12, 87, 105, 109, 120–1, 131–2, 134, 136–7, 149–50, 155, 159, 172, 175, 202, 205, 213–16, 219, 221, 225, 227, 229, 242, 254–5, 258, 265, 270–3, 299, 326, 332

nutritive tissue 10–13, 31, 37, 42–4, 46, 78, 90, 106, 111, 115–18, 121, 124, 128–9, 131–2, 134–7, 147–50, 152–3, 155, 158, 163–5, 167–8, 172–4, 176, 184, 191–2, 195–6, 199–200, 202, 205–6, 212, 214, 217, 219, 221–3, 225, 227, 229, 231, 237, 247–8, 254, 258–9, 261, 264–5, 270–1, 273, 325–6, 377, 498

nymphs 45, 77, 79, 88–90, 94, 95, 97, 111, 113, 114, 119, 124–8, 140, 141, 154, 157–9, 163, 165–6, 168, 170, 291, 362, 365, 366, 445, 453, 490

Oak Apple Day 446

oak apple gall 252, 262–3, 514

oak apple gall wasp 252, 262–3, 514

oak cynipine galls 329

oak galls 263, 323–4, 328, 334, 342–3, 345, 389, 433, 436, 438–40, 466, 475–6, 489, 494, 498

oak pea gall wasp 266

oak spangle galls 267–8

oligophagous 99, 105, 373

omnipotent 2, 11

Oomycota 27–8, 31

opines 25, 51

organoid galls 6

Osten-Sacken, Carl R. 494

ostiole 201–3

Outer Hebrides 269, 426

ovaries 19, 35, 41, 72–3, 182, 189, 201, 371, 437

oviparous 94–5

ovipositor 84, 99, 101, 112, 130, 187–8, 194, 201–3, 209, 211, 219, 222, 224, 231–4, 238, 247, 254, 257–8, 261, 264, 266, 297, 299, 302, 304–8, 312, 315, 317–18, 320–2, 336–9, 341, 345–7, 375, 377, 382–3, 385, 396, 459, 488, 493

ovoviviparous 80, 97, 127

Palaearctic 192, 200, 241–2, 246, 255, 269, 369, 374, 389–90, 408–9, 421, 432–3

palisade 116, 118, 120–1, 124, 129, 132, 134–5, 150, 172–3, 175, 255

Panama 415

pappus 189

parasite 21, 53, 55, 62, 284, 289

parasites 3, 11, 18, 21–2, 26, 28, 29, 30, 35, 36–7, 47, 70, 212–13, 498–9

parasitic plants 18, 36

galls caused by 18, 20, 36–7, 47, 53–4

parasitoids 252, 284–5, 290, 297–8, 301–2, 304–5, 307, 309, 312, 317–18, 320, 326–7, 329, 331, 335–9, 341, 344–8, 386, 396–7, 399–400, 428–34, 457, 466

gregarious 298, 313, 337

parenchyma 5, 25, 50, 61, 74, 90, 93, 148, 151, 159, 161, 169–70, 188, 195, 206, 208–10, 216, 219, 221, 225, 227, 229, 234, 237, 247–8, 255, 258, 263, 269, 271–2, 316–17, 325–6, 351, 498

parthenogenesis 80, 83, 86, 94, 98, 101, 143, 163, 170, 229, 242–4, 249, 251, 256–7, 260, 266, 360–1, 387, 494–5, 497

cyclical parthenogenesis 94, 243, 249, 360, 387

parthenogenetic 43, 92–5, 98, 105, 126, 141, 163, 217, 232, 245, 308, 312, 360, 465, 494, 497

pathogen 25, 28–9, 31, 47, 50, 55, 59, 67, 70, 284, 288, 292, 415, 438, 447, 452, 456

patterns of galling 408–9, 412, 419, 422

pea galls 244, 249, 255–6, 266–8, 328, 334, 338, 341, 383

Pediaspidini 240–1, 244, 249, 252–3, 261–2, 387–8, 390

pedipalps 79, 83

perennials 34, 41, 70–1, 74, 160, 188–9, 460–1

Peronosporales 27–8

pests 37, 80, 82, 84, 100, 117, 157, 180, 212–14, 419, 423, 432, 435, 446–7, 454–5, 465

petals 19, 40, 43, 182

petiole 104, 142, 158, 162, 167, 169, 207, 209–11, 234, 236, 260, 350, 362, 383, 495

phenology 416, 428

phenols 121–2

phenotype 357, 364, 395, 397

pheromones 105, 188, 213, 227

phloem 22, 24, 39, 44, 50, 52–3, 87, 90, 93, 97, 113, 121, 127, 130, 136–9, 142, 163, 170, 210, 265

photosynthesis 3, 22, 29, 39, 54, 156, 173

phyllanthy/phyllody 18–19, 39–40, 43, 89

phyllodes 87

phylloxera (grapvine) 88, 138–9, 141, 447, 452–3

phylogenetic 352, 356, 370

phylum 28, 31, 213

phytomycetophagy 370–1

phytophagy 105

phytoplasma 26, 38–40, 80

pineapple galls v, 76, 95, 128, 153–4, 162–3, 169, 288, 361

pinnule 114

pit galls 7, 88–9, 90, 97, 123–5, 127

pith 7, 46, 160–1, 206, 208, 210, 219, 221–2, 225, 227, 392

plant bug pouches 137

plant bugs 22, 88, 124, 162

plant hormones, see auxins, indole acetic acid, cytokinins, tryptophan, opines, growth hormone

plasmids 5, 23–5, 50–2

Plasmodiophoromycota 27–8

plasmodium 27, 54–6

plate tectonics 358, 403

Pliny the Elder 257, 436, 439–40, 445, 473, 476–7, 481

pollen tube 66

pollination 148, 182, 204, 401–3

polymorphism 86, 95, 97–8, 401

polyphagous 105, 213, 285, 291, 302–3, 307, 309, 313, 324, 328, 334, 336, 338–9, 342, 373, 427, 430, 433

polyphyletic 89, 352–3, 356, 375, 389

polyploid 110, 134

population dynamics 344, 347, 535

Portugal 421, 429, 433

pouches v, 8–9, 76, 78, 82, 88–9, 101, 107–8, 111, 123–4, 131–2, 134–5, 137, 142, 145–6, 149–50, 152, 217, 235, 286–8, 293, 371, 402, 475

pouch gall 89, 124, 135–7, 142, 145–6, 148, 150

powdery scab of potato 56–7

predator 180, 283–6, 288, 299, 309, 315, 327, 331, 380

prepupa 86, 101, 232–3

primary host 33, 35, 69, 91–2, 94, 141–2, 144–5, 163, 166, 284, 361–2, 364, 368, 450

procecidium 382

prokaryote 24, 503

promycelium 34, 67, 72–3

propagule 22

protozoa 27–9, 31, 410

psyllid shoot tip galls 157

ptilinum 225, 228, 322

pupae 46, 83, 86, 101, 105, 107, 115, 120, 136, 146, 151, 172–3, 181, 183, 189, 198, 206, 209, 219, 223, 225, 236, 239, 250, 287, 290, 297–8, 300–1, 306–8, 312, 314–15, 317, 339–40, 455, 490

puparium 46, 181, 188, 191, 225, 297, 299–301, 322

pustules 2, 33–4, 82, 128–9, 286, 476

Rabbi Rashi 477

Ray, John 467, 483–8, 492, 526–7

Redi, Francesco 482–4

Rennie, J. 493

rhachis 350

rhizomes 74, 160, 226, 460

RNA (ribonucleic acid) 13, 21–5, 258

mRNA 21–2, 25

robin’s pincushion, see bedeguar gall rolls v, 6, 8–9, 76, 78, 80, 82, 88–90, 99, 101, 107–8, 110–11, 113–17, 119–20, 122–3, 163, 217, 235, 288, 294, 316, 367, 397

Romanes 357, 497–8

root collar 215

root crown 208

roots v, 2–3, 5, 9–10, 18–19, 22, 31, 38–9, 46, 49–52, 54, 56–7, 90, 97, 101, 108, 119, 138–9, 141–2, 144, 178, 181, 184–5, 201, 205–6, 212, 214, 228, 244, 246–7, 263, 284, 292, 349, 355, 357, 362, 364, 390, 393, 400, 404, 415, 438, 452–3, 461–3, 487

rose galls 244, 248, 256, 324

rose gall wasps 252, 255, 389, 400

rosette galls v, 7, 9, 76, 82, 101, 107, 111, 153–4, 171–3, 190, 207–8, 217, 235, 462

rostrum 189, 206

rotifer 3, 47–9, 494

rusts 3, 8, 29–32, 34–5, 42, 67–70, 72, 450

St Augustine 482–3

St Anthony 447–9

St Anthony’s fire, 76, 447–8, see also ergotism saprophagous 180

saprotrophic 35

sawflies 3–4, 7–8, 76, 98–101, 107, 111, 178, 205, 209–11, 230–4, 286, 296, 316, 326, 346–7, 349, 351, 364, 369, 381–3, 385, 392, 412, 416–18, 488, 499

and the evolution of gall structure and position 381

leaf and bud galls 231

stem galls 210

sclerenchyma 111, 121–2, 137, 149, 159, 161, 170, 200, 229, 248, 271–2, 326, 498

scleromorphic 413–15

sclerophyllous 414–15

Scotland 48, 56, 61, 70, 257, 266, 269, 277, 420, 423–7

secondary host 35, 67, 90, 94–5, 144, 163–4, 166, 361–2, 368, 450

seeds 3, 10, 13, 22, 31, 37, 39–40, 65–6, 72, 101, 104, 165, 179, 182, 185, 198, 202–3, 208, 225, 239, 250, 254, 357, 374, 387, 402–3, 447, 457–62, 464, 491

senescence 29

septum/septa 27, 29, 34, 253–4

setae 79, 238

sexual reproduction 23, 29–30, 59, 65, 68, 90, 93–5, 98, 101, 105–6, 140–2, 144, 163–6, 170, 213, 232, 237, 242–6, 248–52, 261–3, 265–6, 268–9, 271–4, 276–80, 308, 323, 329–32, 335–8, 341, 343, 345, 360–2, 364, 368–9, 387, 390–1, 393, 398, 421, 424, 426, 428–31, 453, 471, 473–4, 494–97

sexuales 94, 119, 139, 141, 165–7

sexuparae 93, 119, 139, 144–5, 166–7, 364

Shick-Shack Day 446

shoot tip galls 157

sieve tubes 22, 25, 50, 87, 90, 93

single swollen cells 48

smuts and bunts 29, 35

soldiers 86–7, 97–8, 145, 292–4, 365–8, 406

solid swellings v, 18, 36, 47, 49–50, 53–4, 214

caused by bacteria 50

caused by fungi 54

caused by parasitic plants 53

caused by viruses 49

sorus 60–1, 507

South Africa 129–30, 433, 457, 464–5

Spain 69, 142, 291, 421, 429, 433, 437

spangle galls 7, 267–71, 335, 337, 348, 438, 494

speciation 183, 358–9, 373, 389, 391–3, 406, 415

species diversity 97, 101, 180, 237, 389, 412, 415, 426

richness 14, 241, 382, 389, 408–18, 421, 426

spermatophore 43, 80, 110

spindle galls 7, 69, 261, 296

spiny galls 7, 34, 36, 69, 169, 170, 179, 248, 391, 393–4, 398–9

spongy mesophyll 116, 118, 120, 124, 129–32, 134–5, 175, 232

spontaneous generation 468, 475, 482–4, 487, 490–1

spores 4, 22, 25, 27, 29–36, 47, 56–63, 66, 68–9, 73–4, 83, 85–6, 120, 218–21, 237–9, 292, 343, 349–50, 371–2, 444, 451, 492

squatters 285–6

stamens 19, 40, 43, 59, 73, 194

starch 10, 12, 127, 147–9, 229, 247, 255, 265, 271, 445

stem boring 8, 159, 357, 406

stem galls 13, 185–6, 200, 205, 210, 217–18, 220, 223, 228, 235, 260–1, 296, 316–17, 347, 370, 376, 379, 383, 454

stems v, 2–4, 7–10, 19–20, 32, 35–6, 38, 46, 49–50, 53–4, 57, 61, 69, 72, 74, 82, 88, 99–101, 104–6, 108, 123–4, 132, 138, 146, 148, 161, 170, 178–81, 185, 198, 201, 205–6, 208–16, 220–2, 224, 227–8, 230–1, 237, 243, 246–7, 253, 255, 259, 284, 294–5, 300, 302–3, 316, 322, 324, 328, 338, 343, 349, 376, 379–80, 383, 390, 392, 404–5, 438, 451, 461–2, 464

sternal spatula 102, 219, 223, 236, 239

stigma 66, 182, 201–2, 325, 327

stomata 33–4, 59, 72, 74, 118, 129, 132, 175, 263

style 66, 182, 202

subtropical 4, 84, 97, 180–1, 400, 409

successori 159, 285–6, 332, 342

superstitions 435, 445

sycamore gall wasps 249–50, 261–2, 345

syconium 182

symbiont 104–5, 373

symbiosis 2, 371, 400

sympatric speciation 359

symplesiomorphy 353–4

synapomorphy 353

syncytium 212

Syria 425–6, 443

tannins 13, 109, 121–2, 127, 263–4, 271, 276, 334, 342, 344, 414, 438–40, 442

temperate 4, 36, 80, 83, 89–90, 92–3, 95, 98–9, 102, 169, 180, 213, 241–2, 343, 360, 382, 408–13

temperature 86, 117, 129–30, 156, 226, 257, 292

tephritid galls 179, 184–5, 294, 295–301

tephritids 7, 13, 178–9, 181, 184–7, 190–1, 193, 200, 205, 224, 228, 250, 286, 296–7, 303, 319, 346, 374, 376, 419, 421, 488

tephritid stem galls 223

thelytokous parthenogenesis 80, 86, 94, 98, 143, 163, 170, 229, 243–4, 249, 251, 257, 260, 266

thelytoky 126, 141, 243, 249–50, 252–3, 256, 308

von Hartig, Theodor 494

Theophrastus 257, 435, 438, 468–9, 471–7, 479, 481, 483

galls of 469

thistle galls 297, 300, 346, 445, 178–9, 186–7, 189–94, 196–7, 206, 214–15, 223–4, 226–7, 252, 295, 296–9, 300–1, 304–5, 374, 377, 419–20, 445, 457–61, 481

three-way galls 369

thrips 3–4, 7, 23, 76–7, 82–4, 86–7, 90, 98, 107, 111, 117, 121–4, 131–2, 135–7, 284, 286–7, 293–4, 346, 349, 357, 406

thrips galls 293

top-down effects 283

totipotent 11

tropical 4, 14, 84, 90, 92–3, 97–8, 102, 104, 162, 169, 180–1, 213, 241, 350, 362, 382, 400, 408–15, 442–3, 455, 477

rainforests 411

tryptophan 13, 52

tubers 31, 56–7, 59, 60, 61, 214

Turkey 263, 389, 390, 421, 424–6, 433, 439, 443, 473

unicellular 73, 108, 170

unilocular 473, 498

univoltine 373

van Leeuwenhoek, Anton 483

vascular tissue 24, 46, 50, 53, 59, 71, 87, 117, 120, 126, 130, 132, 137, 150, 164, 170, 175, 189, 206–08, 210, 212, 221–2, 227, 229, 270, 326

bundle 195, 199, 218

Vaucher, Jean Pierre Étienne 493–4

vectors 22–3, 25, 38, 49, 57, 80, 84, 154, 214

virescences 4, 6, 9, 17–20, 26, 36–42, 44, 46–7, 82, 212, 214

virescences and witches’ brooms 19–42

viruses 20, 22–3, 47

virus galls 49

viviparous 94–5, 97–8, 138, 141, 168

von Hartig, Theodor 494, 499

wart disease of potato 59

wax 88, 97, 142, 145, 164, 167, 290, 367

white blister of brassicas 57

White, Gilbert 488–9

white pine blister rust 70

willow galls 382–3

witches’ brooms v, ix, 1, 6, 9, 18–20, 26, 28, 30, 36–41, 78, 80

Wolbachia and loss of sex in gall wasps 251

woody tissue 150, 192, 227, 255, 270

xylem 24, 50, 52–3, 118, 130, 210, 265

Zygomycota 27–8

The New Naturalist Library

1. Butterflies—E. B. Ford

2. British Game—B. Vesey-Fitzgerald

3. London’s Natural History—R. S. R. Fitter

4. Britain’s Structure and Scenery—L. Dudley Stamp

5. Wild Flowers—J. Gilmour & M. Walters

6. The Highlands & Islands—F. Fraser Darling & J. M. Boyd

7. Mushrooms & Toadstools—J. Ramsbottom

8. Insect Natural History—A. D. Imms

9. A Country Parish—A. W. Boyd

10. British Plant Life—W. B. Turrill

11. Mountains & Moorlands—W. H. Pearsall

12. The Sea Shore—C. M. Yonge

13. Snowdonia—F. J. North, B. Campbell & R.Scott

14. The Art of Botanical Illustration—W. Blunt

15. Life in Lakes & Rivers—T. T. Macan & E. B. Worthington

16. Wild Flowers of Chalk & Limestone—J. E. Lousley

17. Birds & Men—E. M. Nicholson

18. A Natural History of Man in Britain—H. J. Fleure & M. Davies

19. Wild Orchids of Britain—V. S. Summerhayes

20. The British Amphibians & Reptiles—M. Smith

21. British Mammals—L. Harrison Matthews

22. Climate and the British Scene—G. Manley

23. An Angler’s Entomology—J. R. Harris

24. Flowers of the Coast—I. Hepburn

25. The Sea Coast—J. A. Steers

26. The Weald—S. W. Wooldridge & F. Goldring

27. Dartmoor—L. A. Harvey & D. St. Leger Gordon

28. Sea Birds—J. Fisher & R. M. Lockley

29. The World of the Honeybee—C. G. Butler

30. Moths—E. B. Ford

31. Man and the Land—L. Dudley Stamp

32. Trees, Woods and Man—H. L. Edlin

33. Mountain Flowers—J. Raven & M. Walters

34. The Open Sea: I. The World of Plankton—A. Hardy

35. The World of the Soil—E. J. Russell

36. Insect Migration—C. B. Williams

37. The Open Sea: II. Fish & Fisheries—A. Hardy

38. The World of Spiders—W. S. Bristowe

39. The Folklore of Birds—E. A. Armstrong

40. Bumblebees—J. B. Free & C. G. Butler

41. Dragonflies—P. S. Corbet, C. Longfield & N. W. Moore

42. Fossils—H. H. Swinnerton

43. Weeds & Aliens—E. Salisbury

44. The Peak District—K. C. Edwards

45. The Common Lands of England & Wales—L. Dudley Stamp & W. G. Hoskins

46. The Broads—E. A. Ellis

47. The Snowdonia National Park—W. M. Condry

48. Grass and Grasslands—I. Moore

49. Nature Conservation in Britain—L. Dudley Stamp

50. Pesticides and Pollution—K. Mellanby

51. Man & Birds—R. K. Murton

52. Woodland Birds—E. Simms

53. The Lake District—W. H. Pearsall & W. Pennington

54. The Pollination of Flowers—M. Proctor & P. Yeo

55. Finches—I. Newton

56. Pedigree: Words from Nature—S. Potter & L. Sargent

57. British Seals—H. R. Hewer

58. Hedges—E. Pollard, M. D. Hooper & N. W. Moore

59. Ants—M. V. Brian

60. British Birds of Prey—L. Brown

61. Inheritance and Natural History—R. J. Berry

62. British Tits—C. Perrins

63. British Thrushes—E. Simms

64. The Natural History of Shetland—R. J. Berry & J. L. Johnston

65. Waders—W. G. Hale

66. The Natural History of Wales—W. M. Condry

67. Farming and Wildlife—K. Mellanby

68. Mammals in the British Isles—L. Harrison Matthews

69. Reptiles and Amphibians in Britain—D. Frazer

70. The Natural History of Orkney—R. J. Berry

71. British Warblers—E. Simms

72. Heathlands—N. R. Webb

73. The New Forest—C. R. Tubbs

74. Ferns—C. N. Page

75. Freshwater Fish—P. S. Maitland & R. N. Campbell

76. The Hebrides—J. M. Boyd & I. L. Boyd

77. The Soil—B. Davis, N. Walker, D. Ball & A. Fitter

78. British Larks, Pipits & Wagtails—E. Simms

79. Caves & Cave Life—P. Chapman

80. Wild & Garden Plants—M. Walters

81. Ladybirds—M. E. N. Majerus

82. The New Naturalists—P. Marren

83. The Natural History of Pollination—M. Proctor, P. Yeo & A. Lack

84. Ireland: A Natural History—D. Cabot

85. Plant Disease—D. Ingram & N. Robertson

86. Lichens—Oliver Gilbert

87. Amphibians and Reptiles—T. Beebee & R. Griffiths

88. Loch Lomondside—J. Mitchell

89. The Broads—B. Moss

90. Moths—M. Majerus

91. Nature Conservation—P. Marren

92. Lakeland—D. Ratcliffe

93. British Bats—John Altringham

94. Seashore—Peter Hayward

95. Northumberland—Angus Lunn

96. Fungi—Brian Spooner & Peter Roberts

97. Mosses & Liverworts—Nick Hodgetts & Ron Porley

98. Bumblebees—Ted Benton

99. Gower—Jonathan Mullard

100. Woodlands—Oliver Rackham

101. Galloway and the Borders—Derek Ratcliffe

102. Garden Natural History—Stefan Buczacki

103. The Isles of Scilly—Rosemary Parslow

104. A History of Ornithology—Peter Bircham

105. Wye Valley—George Peterken

106. Dragonflies—Philip Corbet & Stephen Brooks

107. Grouse—Adam Watson & Robert Moss

108. Southern England—Peter Friend

109. Islands—R. J. Berry

110. Wildfowl—David Cabot

111. Dartmoor—Ian Mercer

112. Books and Naturalists—David E. Allen

113. Bird Migration—Ian Newton

114. Badger—Timothy J. Roper

115. Climate and Weather—John Kington

116. Plant Pests—David V. Alford

Copyright

COLLINS NEW NATURALIST LIBRARY (117)—PLANT GALLS. © Margaret Redfern, 2011. All rights reserved under International and Pan-American Copyright Conventions. By payment of the required fees, you have been granted the non-exclusive, non-transferable right to access and read the text of this e-book on-screen. No part of this text may be reproduced, transmitted, down-loaded, decompiled, reverse engineered, or stored in or introduced into any information storage and retrieval system, in any form or by any means, whether electronic or mechanical, now known or hereinafter invented, without the express written permission of HarperCollins e-books.

A CIP catalogue record for this book is available from the British Library.

EPub Edition © FEBRUARY 2011 ISBN: 978-0-00-743191-5

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