Insect Biodiversity Current Trends and Future Prospects 9781118945575, 1118945573, 9781118945605, 1118945603

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Insect Biodiversity

Insect Biodiversity Science and Society Volume II

Edited by Robert G. Foottit

Agriculture and Agri-Food Canada Ottawa Ontario Canada

Peter H. Adler

Clemson University Clemson South Carolina USA

This edition first published 2018 © 2018 John Wiley & Sons Ltd All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/go/permissions. The right of Robert G. Foottit and Peter H. Adler to be identified as the authors of the editorial material in this work has been asserted in accordance with law. Registered Office(s) John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Editorial Office 9600 Garsington Road, Oxford, OX4 2DQ, UK For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print‐on‐demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Library of Congress Cataloging‐in‐Publication Data applied for ISBN: 9781118945575

Cover Design: Wiley Cover Image: The cover art, prepared by Matthew A. Bertone (North Carolina State University, Raleigh), shows the vast morphological diversity among treehoppers based on plates by Edwin Wilson from the Biologia Centrali-Americana (sections by W. W. Fowler 1894–1896). Set in 10/12pt Warnock by SPi Global, Chennai, India Printed in the UK by Bell & Bain Ltd, Glasgow. 10 9 8 7 6 5 4 3 2 1

v

Brief Table of Contents 1

Introduction – A Brief History of Revolutions in the Study of Insect Biodiversity  1



Part I   Habitats and Regions 

2

Insect Biodiversity in the Arctic  15

3

Insect Biodiversity in Indochina: A Window into the Riches of the Oriental Region  59

4

Biodiversity of Arthropods on Islands  81

5

Beneficial Insects in Agriculture: Enhancement of Biodiversity and Ecosystem Services  105

6

Insects in Caves  123



Part II   Taxa 

7

Biodiversity of the Thysanurans (Microcoryphia and Zygentoma)  155

8

Biodiversity of Zoraptera and Their Little‐Known Biology  199

9

Biodiversity of Embiodea  219

10

Biodiversity of Orthoptera  245

11

Biodiversity of Phasmatodea  281

12

Biodiversity of Dermaptera  315

13

Biodiversity of Grylloblattodea and Mantophasmatodea  335

14

Biodiversity of Blattodea – the Cockroaches and Termites  359

15

Biodiversity of Mantodea  389

13

153

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Brief Table of Contents

16

Biodiversity of Psocoptera  417

17

Biodiversity of Ectoparasites: Lice (Phthiraptera) and Fleas (Siphonaptera)  457

18

Biodiversity of Thysanoptera  483

19

The Diversity of the True Hoppers (Hemiptera: Auchenorrhyncha)  501

20

The Biodiversity of Sternorrhyncha: Scale Insects, Aphids, Psyllids, and Whiteflies  591

21

Biodiversity of the Neuropterida (Insecta: Neuroptera, Megaloptera, and Raphidioptera)  627

22

Biodiversity of Strepsiptera  673

23

Biodiversity of Mecoptera  705



Part III  Perspectives 

24

The Fossil History of Insect Diversity  723

25

Phenotypes in Insect Biodiversity Research  789

26

Global Change and Insect Biodiversity in Agroecosystems  801

27

Digital Photography and the Democratization of Biodiversity Information  839

28

Bee (Hymenoptera: Apoidea: Anthophila) Diversity Through Time  851

29

Insect Biodiversity in Culture and Art  869



Index of Arthropod Taxa Arranged by Order and Family  899



Index of Arthropod Taxa Arranged Alphabetically  943



Index of non‐Arthropod Taxa Arranged Alphabetically  975



Subject Index  979

721

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Detailed Table of Contents

List of Contributors  xxiii Foreword  xxix Preface, Volume II  xxxiii Acknowledgments  xxxv 1

Introduction – A Brief History of Revolutions in the Study of Insect Biodiversity  1 Peter H. Adler and Robert G. Foottit

1.1 Discovery  1 1.2 Conceptual Development  5 1.3 Information Management  6 1.4 Conclusions  7 Acknowledgments  8 References  8

Part I   Habitats and Regions  13

2

Insect Biodiversity in the Arctic  15 Ian D. Hodkinson

2.1 Documenting Biodiversity – Traditional Taxonomy Versus DNA Barcoding  17 2.2 Insect Species Diversity in the Arctic  18 2.2.1 Composition of the Arctic Insect Fauna  18 2.2.2 Species Richness Trends Along Latitudinal Gradients  25 2.2.3 Geographical and Regional Variations in Species Richness  27 2.2.4 Diversity Oases Within the Arctic  28 2.3 Historical Insect Biodiversity in the Arctic – the Time Perspective  29 2.3.1 Nunataks and Glacial Refugia as Generators of Biodiversity  30 2.3.2 Endemism  31 2.4 Biodiversity on the Landscape Scale  32 2.4.1 Variation in Biodiversity on a Landscape Scale  32 2.4.2 Local Effects on Biodiversity – Predation and Natural Disturbance  34 2.5 Important Characteristics of Arctic Insect Biodiversity  35 2.5.1 Specialist Versus Generalist Species  35 2.5.2 Life‐History Adaptation  35 2.5.3 Genetic Diversity Within Species and Groups  36 2.5.4 Reproductive Variation and Parthenogenesis  36

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2.5.5 A Diversity of Adaptations for Maximizing Heat Absorption  37 2.6 Cold Tolerance – a Diversity of Adaptations  38 2.6.1 Brachyptery and Wing Polymorphism  39 2.7 Dispersal, Immigration, and Biodiversity  39 2.8 Pollinator Networks and Pollinator Biodiversity  40 2.9 A Biodiversity Paradise for Parasites?  41 2.10 Biodiversity and the Changing Arctic Climate  42 References  44 3

Insect Biodiversity in Indochina: A Window into the Riches of the Oriental Region  59 Seunghwan Lee and Ram Keshari Duwal

3.1 Physical Geography and Climate  62 3.2 Features of Insect Biodiversity in the Lower Mekong Subregion  62 3.2.1 Blattodea  70 3.2.2 Coleoptera  70 3.2.3 Dermaptera  71 3.2.4 Diptera  72 3.2.5 Embiodea  72 3.2.6 Ephemeroptera  72 3.2.7 Hemiptera  72 3.2.8 Hymenoptera  72 3.2.9 “Isoptera”  72 3.2.10 Lepidoptera  72 3.2.11 Mantodea  73 3.2.12 Mecoptera  73 3.2.13 Megaloptera  73 3.2.14 Microcoryphia and Zygentoma  73 3.2.15 Neuroptera  73 3.2.16 Notoptera (Grylloblattodea and Mantophasmatodea)  73 3.2.17 Odonata  73 3.2.18 Orthoptera  73 3.2.19 Phasmatodea  73 3.2.20 Phthiraptera  73 3.2.21 Plecoptera  74 3.2.22 Psocoptera  74 3.2.23 Raphidioptera  74 3.2.24 Siphonaptera  74 3.2.25 Strepsiptera  74 3.2.26 Thysanoptera  74 3.2.27 Trichoptera  74 3.2.28 Zoraptera  74 3.3 Insect Biodiversity and Society in Indochina  74 3.3.1 Entomophagy in the Lower Mekong Subregion  74 3.3.2 Research Initiatives  76 3.4 Conclusions  77 Acknowledgments  78 References  78

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4

Biodiversity of Arthropods on Islands  81 Rosemary G. Gillespie and Kipling Will

4.1 What is an Island?  81 4.1.1 History of the Island  82 4.1.2 Degree of Isolation  84 4.1.3 Area of the Island  84 4.1.4 Age of the Island  85 4.2 Ecological Attributes of Islands  85 4.2.1 Species Diversity on Islands  85 4.2.2 Island Colonization  86 4.2.3 Factors Facilitating Establishment  86 4.2.4 Niche Preemption  86 4.2.5 Ecological Release  87 4.2.6 Networks of Ecological Interactions  87 4.3 Evolution on Islands  87 4.3.1 Anagenesis  87 4.3.2 Cladogenesis  87 4.3.3 Adaptive Radiation  88 4.3.4 Isolation, Hybridization, and Admixture  88 4.3.5 Parallel Evolution and Convergence  89 4.4 Evolution in Other Insular Environments  89 4.4.1 Mountaintops – Sky Islands  89 4.4.2 Caves  89 4.4.3 Desert Dunes and Salt Lakes  89 4.4.4 Habitat Fragments  90 4.5 Characteristics of Island Biodiversity  90 4.5.1 Disharmony  90 4.5.2 Endemism  91 4.5.3 Loss of Dispersal Ability and Flightlessness  91 4.5.4 Innovations  91 4.5.5 Size  92 4.5.6 Reproductive Shifts  92 4.6 Conservation  92 4.6.1 Taxonomic Impediments  93 4.6.2 Restricted Ranges and Small Population Sizes  93 4.6.3 Abiotic Factors  93 4.6.4 Invasive Species  94 4.7 Conclusion  94 References  94 5

5.1

Beneficial Insects in Agriculture: Enhancement of Biodiversity and Ecosystem Services  105 Matthew S. Jones and William E. Snyder

Components of Biodiversity: Species Richness, Species Evenness, and Species Identity  106 5.2 Why Does Insect Biodiversity Matter to Agriculture?  106 5.2.1 Complementarity  107

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5.2.1.1 5.2.1.2 5.2.1.3 5.2.2 5.2.3 5.3

Temporal Complementarity  107 Spatial Complementarity  108 Behavioral Complementarity  109 Identity Effects in Pollinator, Predator, and Detritivore Communities  110 Disruptive Species Interactions in Diverse Communities  111 Degradation of Biodiversity Through Agricultural Intensification, and Its Reversal  112 5.4 Restoring Biodiversity to Agroecosystems  112 5.4.1 Restoring Key Resources  112 5.4.2 Optimizing Use of Pesticides  113 5.4.3 Diversifying Farming Landscapes at Larger Scales  113 5.5 Conclusions and Recommendations  115 5.5.1 Clarify Mechanisms Leading to Biodiversity Effects  115 5.5.2 Consider Biodiversity Effects That Span Multiple Ecosystem Services  115 5.5.3 Better Link Management Practices to Beneficial Biodiversity Effects  115 5.5.4 Rank the Relative Importance of Habitat Loss Versus Agrochemical Use  116 5.5.5 Elucidate Strategies That Facilitate Transition from Current Agricultural Production Practices to Those That Are Sustainable and Provide Improved Ecosystem Services  116 5.6 Summary  116 Acknowledgments  117 References  117

6

Insects in Caves  123 David C. Culver and Tanja Pipan

6.1 The Story of Leptodirus hochenwartii  123 6.2 The Variety of Subterranean Spaces  124 6.2.1 Overview  124 6.2.2 Caves  125 6.2.3 Soil and Interstitial Habitats  126 6.2.4 Shallow Subterranean Habitats  127 6.2.4.1 Epikarst  128 6.2.4.2 Milieu Souterrain Superficiel  128 6.2.4.3 Calcrete Aquifers  128 6.2.4.4 Unifying Features of Shallow Subterranean Habitats  130 6.3 Ecological Roles of Insects in Caves  133 6.3.1 Relative Importance of Subterranean Habitats in the Ecology of Different Insects  133 6.3.2 Trophic Roles  134 6.4 Morphological and Life‐History Adaptations of Insects to Subterranean Life  134 6.5 Probable Modes of Successful Colonization of Subterranean Space  138 6.5.1 Initial Colonization  140 6.5.2 Successful Colonization  140 6.5.3 Allopatric Versus Parapatric Speciation  141 6.5.4 Subterranean Dispersal  142 6.6 Taxonomic and Geographic Patterns of Subterranean Insect Biodiversity  142

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6.6.1 Geographic Patterns  142 6.6.2 Taxonomic Review of Troglobiotic Insects  143 6.6.2.1 Collembola  144 6.6.2.2 Diplura  146 6.6.2.3 Coleoptera  146 6.6.2.4 Fulgoromorpha  147 6.7 Human Utility and Protection of Cave Insects  147 References  147 Part II   Taxa  7

153

Biodiversity of the Thysanurans (Microcoryphia and Zygentoma)  155 Luis F. Mendes

7.1 Paleontological Data  159 7.2 Parasitism  167 7.2.1 Unicellular Parasites  167 7.2.2 Nematoda  167 7.2.3 Acarids  167 7.2.4 Strepsiptera  167 7.2.5 Fungi  167 7.3 Predation  168 7.4 Order Microcoryphia (= Archaeognatha)  168 7.4.1 Characterization  168 7.4.2 Bionomics  172 7.4.3 Taxonomy  173 7.4.4 Identification Key for Families, Subfamilies, and Paleoforms of Microcoryphia  174 7.5 Order Zygentoma (= Thysanura Sensu Stricto)  175 7.5.1 Characterization  175 7.5.2 Bionomics  179 7.5.3 Taxonomy  180 7.5.4 Identification Key for Families and Subfamilies of Zygentoma  181 7.6 Genetic Studies of Thysanurans  183 7.7 Thysanurans and Humans  184 7.8 Geographic Distribution of the Thysanurans  185 References  187 8

Biodiversity of Zoraptera and Their Little‐Known Biology  199 Jae C. Choe

8.1 Morphology  201 8.2 Life History and Ecology  204 8.3 Reproduction  208 8.4 Phylogenetic Position – “The Zoraptera Problem”  210 8.5 Conclusion  211 Acknowledgments  212 References  212

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9

Biodiversity of Embiodea  219 Janice S. Edgerly

9.1 Diversity in Habitat and Silk  223 9.2 The Promise of Silk‐Like Biomaterials and Emerging Lessons from Webspinners  228 9.3 Social Behavior  229 9.4 Families of Embiodea  231 9.4.1 Andesembiidae  231 9.4.2 Anisembiidae  232 9.4.3 Archembiidae  233 9.4.4 Australembiidae  234 9.4.5 Clothodidae  234 9.4.6 Embiidae  235 9.4.7 Embonychidae  236 9.4.8 Notoligotomidae  236 9.4.9 Oligotomidae  236 9.4.10 Paedembiidae  238 9.4.11 Ptilocerembiidae  238 9.4.12 Scelembiidae  238 9.4.13 Teratembiidae  239 9.5 Webspinners of the Fossil Record  239 9.6 Conclusion  239 References  240 10

Biodiversity of Orthoptera  245 Hojun Song

10.1 10.2 10.3 10.4 10.5 10.5.1 10.5.1.1 10.5.1.2 10.5.1.3 10.5.1.4 10.5.1.5 10.5.1.6 10.5.1.7 10.5.2 10.5.2.1 10.5.2.2 10.5.2.3 10.5.2.4 10.5.3.5 10.5.3.6 10.5.3.7 10.5.3.8

Taxonomic Classification and Phylogeny  245 Diversity and Distribution  246 Morphological and Biological Diversity  250 Societal Importance  253 Overview of Taxa  254 Suborder Ensifera  254 Superfamily Grylloidea  255 Superfamily Gryllotalpoidea  255 Superfamily Schizodactyloidea  259 Superfamily Rhaphidophoroidea  260 Superfamily Hagloidea  260 Superfamily Stenopelmatoidea  260 Superfamily Tettigonioidea  261 Suborder Caelifera  262 Superfamily Tridactyloidea  263 Superfamily Tetrigoidea  263 Superfamily Eumastacoidea  265 Superfamily Proscopioidea  266 Superfamily Tanaoceroidea  266 Superfamily Trigonopterygoidea  267 Superfamily Pneumoroidea  267 Superfamily Pyrgomorphoidea  267

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10.5.3.9 Superfamily Acridoidea  268 Acknowledgments  271 References  271 11

Biodiversity of Phasmatodea  281 Sven Bradler and Thomas R. Buckley

11.1 Phasmatodean Phylogeny  286 11.2 Overview of Taxa  288 11.2.1 Timema  289 11.2.2 Agathemera  290 11.2.3 Heteronemiinae  290 11.2.4 Aschiphasmatinae  290 11.2.5 Phylliinae – The True Leaf Insects  291 11.2.6 Heteropteryginae  292 11.2.7 Diapheromerinae  293 11.2.8 Pseudophasmatinae  294 11.2.9 Palophinae  294 11.2.10 The African Clade  295 11.2.11 Gratidiini  295 11.2.12 Clitumnini  296 11.2.13 Medaurini  296 11.2.14 Pharnaciini  296 11.2.15 Cladomorphinae  296 11.2.16 Stephanacridini  297 11.2.17 Lanceocercata – The “Marsupials” Among the Phasmatodea  297 11.2.18 Lonchodinae  299 11.2.19 Necrosciinae  300 11.3 The Phasmatodean Fossil Record  300 11.4 Phasmatodea as Research Tools  302 11.5 Importance to Human Society  304 References  304 12

Biodiversity of Dermaptera  315 Fabian Haas

12.1 Epizoic Dermaptera  315 12.2 Structure and Function  318 12.3 Locomotion  319 12.4 Distribution  319 12.5 Development and Reproduction  323 12.6 Behavior  323 12.6.1 Mating Behavior and Maternal Care  323 12.6.2 Defense  324 12.6.3 Feeding  324 12.7 Parasitism and Symbiosis  324 12.8 Fossils and Research History  324 12.9 Overview of Taxa  325 12.9.1 Lower Dermaptera  325

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12.9.2 Higher Dermaptera  326 12.10 Societal and Scientific Importance  326 12.10.1 Plant Pests, Biological Control Agents, and General Nuisances  326 12.10.2 Medical, Veterinary, and Forensic Importance  326 12.10.3 Invasive Alien Species  327 12.10.4 Pollination and Other Ecological Services  327 12.10.5 Research Tools  327 12.10.6 Conservation – Vanishing Species  328 12.10.7 Cultural Legacy  328 Acknowledgments  328 References  328 13

Biodiversity of Grylloblattodea and Mantophasmatodea  335 Monika J. B. Eberhard, Sean D. Schoville and Klaus‐Dieter Klass

13.1 Grylloblattodea  336 13.1.1 Morphology and Biology  336 13.1.2 Overview of Taxa  341 13.2 Mantophasmatodea  343 13.2.1 Morphology and Biology  343 13.2.2 Overview of Taxa  346 13.2.2.1 Tanzaniophasmatidae  349 13.2.2.2 Mantophasmatidae  349 13.2.2.3 Tyrannophasma/Praedatophasma Clade  350 13.2.2.4 Austrophasmatidae  350 13.3 Fossil Record  351 13.4 Conclusions  352 Acknowledgments  353 References  353 14

Biodiversity of Blattodea – the Cockroaches and Termites  359 Marie Djernæs

14.1 Overview of Taxa  362 14.1.1 Superfamily Corydioidea  363 14.1.1.1 Family Corydiidae  363 14.1.1.2 Family Nocticolidae  365 14.1.2 Superfamily Blaberoidea  366 14.1.2.1 Family Ectobiidae  366 14.1.2.2 Family Blaberidae  368 14.1.3 Superfamily Blattoidea  369 14.1.3.1 Family Blattidae  369 14.1.3.2 Family Lamproblattidae  370 14.1.3.3 Family Tryonicidae  371 14.1.3.4 Family Anaplectidae  371 14.1.3.5 Family Cryptocercidae  371 14.1.3.6 Termites  371 14.2 Societal Importance  373

Detailed Table of Contents

14.2.1 Cockroaches and Science  373 14.2.2 Cockroaches as Pests  374 14.2.3 Cockroaches as Food, Feed, and Medicine  375 14.2.4 Pet and Feeder Species  376 14.2.5 Ecological Importance  376 14.2.6 Conservation Status  377 References  377 15

Biodiversity of Mantodea  389 Frank Wieland and Gavin J. Svenson

15.1 Morphological and Biological Diversity  391 15.2 Phylogeny and Classification  396 15.2.1 Acanthopidae  396 15.2.2 Acontistidae  396 15.2.3 Amorphoscelidae  397 15.2.4 Angelidae  398 15.2.5 Chaeteessidae  398 15.2.6 Coptopterygidae  399 15.2.7 Empusidae  399 15.2.8 Epaphroditidae  399 15.2.9 Eremiaphilidae  400 15.2.10 Galinthiadidae  400 15.2.11 Hymenopodidae  401 15.2.12 Iridopterygidae  401 15.2.13 Liturgusidae  401 15.2.14 Mantidae  402 15.2.15 Mantoididae  402 15.2.16 Metallyticidae  403 15.2.17 Photinaidae  403 15.2.18 Stenophyllidae  404 15.2.19 Tarachodidae  404 15.2.20 Thespidae  404 15.2.21 Toxoderidae  405 15.2.22 Incertae Sedis  405 15.2.23 Suprafamilial Groups  405 15.2.23.1 Acanthopoidea 405 15.2.23.2 Artimantodea 405 15.2.23.3 Cernomantodea 406 15.2.23.4 Eumantodea 406 15.2.23.5 Mantidea 406 15.2.23.6 Mantoidea 406 15.2.23.7 Mantomorpha 406 15.2.23.8 Neomantodea 406 15.3 Morphological Convergence and Ecomorphs  406 15.4 Conclusions  407 References  407

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16

Biodiversity of Psocoptera  417 Edward L. Mockford

16.1 Classification  418 16.2 Overview of the Psocoptera  422 16.2.1 Suborder Trogiomorpha  422 16.2.1.1 Infraorder Atropetae  423 16.2.1.2 Infraorder Psocatropetae  434 16.2.1.3 Infraorder Prionoglaridetae  434 16.2.2 Suborder Troctomorpha  434 16.2.2.1 Infraorder Nanopsocetae  434 16.2.2.2 Infraorder Amphientometae  436 16.2.2.3 Superfamily Amphientomoidea  436 16.2.2.4 Superfamily Electrentomoidea  437 16.2.3 Suborder Psocomorpha  438 16.2.3.1 Infraorder Archipsocetae  438 16.2.3.2 Infraorder Caeciliusetae  438 16.2.3.3 Infraorder Homilopsocidea  441 16.2.3.4 Infraorder Philotarsetae  443 16.2.3.5 Infraorder Epipsocetae  444 16.2.3.6 Infraorder Psocetae  445 16.3 Summary of Diversity of the Psocoptera and Predictions  447 16.4 The Importance to Humans of Psocopteran Biodiversity  448 Acknowledgments  448 References  449 17

Biodiversity of Ectoparasites: Lice (Phthiraptera) and Fleas (Siphonaptera)  457 Terry D. Galloway

17.1 Phthiraptera – The Parasitic Lice  458 17.2 Siphonaptera – The Fleas  465 17.3 Medical and Veterinary Importance  474 17.3.1 Lice  474 17.3.2 Fleas  475 17.4 Community Diversity of Lice and Fleas  477 17.5 Conservation of Lice and Fleas  478 Acknowledgments  479 References  479 18

Biodiversity of Thysanoptera  483 Laurence A. Mound

18.1 18.2 18.3 18.4 18.5 18.6 18.7 18.8

What Are Thrips?  484 Family Diversity  484 The Lives of Thrips  486 Thrips Around the World  487 Thrips as Research Targets  488 Structural Diversity of Thrips  491 Thrips as Pests  493 Thrips and Human Life  494

Detailed Table of Contents

18.9 Thrips Information Sources  495 References  496 19

The Diversity of the True Hoppers (Hemiptera: Auchenorrhyncha)  501 Charles R. Bartlett, Lewis L. Deitz, Dmitry A. Dmitriev, Allen F. Sanborn, Adeline Soulier‐Perkins and Matthew S. Wallace

19.1 Overview of the Auchenorrhyncha  511 19.1.1 Cicadomorpha  511 19.1.1.1 Superfamily Cicadoidea – The Cicadas: Cicadidae and Tettigarctidae  516 19.1.1.2 Superfamily Cercopoidea – Spittlebugs or Froghoppers  518 19.1.1.3 Superfamily Membracoidea – Leafhoppers and Treehoppers  521 19.1.2 Fulgoromorpha  530 19.1.2.1 Superfamily Fulgoroidea – The Planthoppers  536 19.2 Prospectus  549 Acknowledgments  550 References  551 20

The Biodiversity of Sternorrhyncha: Scale Insects, Aphids, Psyllids, and Whiteflies  591 Nate B. Hardy

20.1 Sternorrhyncha and Society  591 20.1.1 Economic Importance  591 20.1.2 Ecological Importance  593 20.1.3 Existential Importance  593 20.2 Taxonomic Diversity of Sternorrhyncha  593 20.2.1 Phylogeny and Classification  593 20.2.1.1 Aphidoidea  594 20.2.1.2 Aleyrodoidea  594 20.2.1.3 Coccoidea  595 20.2.1.4 Psylloidea  595 20.3 Functional Diversity of Sternorrhyncha  596 20.3.1 Trophic Diversity  596 20.3.1.1 Phloem Feeding  596 20.3.1.2 Not Phloem Feeding  596 20.3.1.3 Trophic‐Breadth Variation  596 20.3.2 Trophic Evolution  597 20.3.3 Endosymbiosis  598 20.3.4 Endosymbiont Diversity  598 20.3.4.1 Endosymbiont Phylogenetic Diversity  598 20.3.4.2 Endosymbiont Functional Diversity  602 20.3.5 Endosymbiont Evolution  604 20.3.5.1 Ecological Speciation  605 20.3.5.2 Conflictual Speciation  606 20.3.6 Life‐Cycle Diversity  607 20.3.6.1 Aphid Soldiers and Eusocial Societies  608 20.3.6.2 Life‐Cycle Evolution  609 20.3.7 Genetic‐System Diversity  610 20.3.7.1 Holocentric Chromosomes  610

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20.3.7.2 20.3.7.3 20.3.7.4 20.3.8 20.3.8.1 20.3.8.2

Sex Determination and Parthenogenesis  610 Sex Ratio  611 Supernumerary Chromosomes  613 Genetic‐System Evolution  613 What Sternorrhyncha Can Tell Us About the Evolution of Sex  613 What Sternorrhyncha Can Tell Us About the Evolution of Genetic Systems  614 20.4 Conclusions  615 Acknowledgments  616 References  616 21

Biodiversity of the Neuropterida (Insecta: Neuroptera, Megaloptera, and Raphidioptera)  627 John D. Oswald and Renato J. P. Machado

21.1 Phylogeny  628 21.2 Geological Age  628 21.3 Metamorphosis and Life Stages  629 21.3.1 Adults  629 21.3.2 Eggs and Oviposition  630 21.3.3 Larvae  632 21.3.4 Pupae  633 21.4 Biology  634 21.5 Distribution  636 21.6 Overview of Orders and Families  637 21.6.1 Order Megaloptera  642 21.6.1.1 Family Corydalidae  642 21.6.1.2 Family Sialidae   642 21.6.2 Order Neuroptera  644 21.6.2.1 Family Ascalaphidae  644 21.6.2.2 Family Berothidae  645 21.6.2.3 Family Chrysopidae   645 21.6.2.4 Family Coniopterygidae  647 21.6.2.5 Family Dilaridae  647 21.6.2.6 Family Hemerobiidae  649 21.6.2.7 Family Ithonidae  649 21.6.2.8 Family Mantispidae  650 21.6.2.9 Family Myrmeleontidae  651 21.6.2.10 Family Nemopteridae  652 21.6.2.11 Family Nevrorthidae  653 21.6.2.12 Family Nymphidae  653 21.6.2.13 Family Osmylidae  655 21.6.2.14 Family Psychopsidae  656 21.6.2.15 Family Sisyridae  656 21.6.3 Order Raphidioptera  657 21.6.3.1 Family Inocelliidae  657 21.6.3.2 Family Raphidiidae  657 21.7 Societal Importance  658

Detailed Table of Contents

21.8 Scientific Importance  659 Acknowledgments  660 References  660 22

Biodiversity of Strepsiptera  673 Jeyaraney Kathirithamby

22.1 Family Bahiaxenidae  678 22.2 Suborder Mengenillidia  678 22.2.1 Family Mengenillidae  678 22.3 Suborder Stylopidia  681 22.3.1 Family Corioxenidae  685 22.4 Infraorder Stylopiformia  685 22.4.1 Family Myrmecolacidae  685 22.4.2 Family Lychnocolacidae  688 22.4.3 Family Stylopidae  688 22.4.4 Family Xenidae  689 22.4.5 Family Bohartillidae  690 22.4.6 Family Elenchidae  691 22.4.7 Family Halictophagidae  692 22.5 Conclusions  694 Acknowledgments  694 References  694 23

Biodiversity of Mecoptera  705 Wesley J. Bicha

23.1 Suborder Nannomecoptera  706 23.1.1 Family Nannochoristidae  706 23.2 Suborder Pistillifera  707 23.2.1 Infraorder Raptipedia  707 23.2.1.1 Family Bittacidae  707 23.2.2 Infraorder Opisthogonopora  709 23.2.2.1 Group Boreomorpha  710 23.2.2.2 Group Meropomorpha  711 23.2.2.3 Group Panorpomorpha  711 23.3 Societal Value of Mecoptera  715 23.4 Scientific Value of Mecoptera  716 23.5 Conclusion  716 References  716 Part III  Perspectives 

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24

The Fossil History of Insect Diversity  723 Conrad C. Labandeira

24.1 24.2 24.2.1

Importance of the Insect Fossil Record  724 Types of Insect Diversity Past and Present  725 Taxonomic and Taxic Diversity  725

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24.2.2 Ecological Diversity  730 24.2.3 Biotal Diversity  733 24.2.4 Plant–Insect Interactional Diversity  735 24.2.4.1 Short‐Term Studies  746 24.2.4.2 Intermediate‐Term Studies  746 24.2.4.3 Long‐Term Studies  747 24.2.4.4 Very Long‐Term Studies  747 24.2.5 Morphological Diversity  749 24.2.5.1 Size Disparity  753 24.2.5.2 Structural Disparity  753 24.2.5.3 Developmental Disparity  757 24.2.5.4 Key Innovations  757 24.2.6 Functional Diversity  760 24.2.6.1 Functional Feeding Groups  760 24.2.6.2 Lacustrine Ecospace Occupation  760 24.2.6.3 Parasitoids and Trophic Roles in Food Webs  761 24.2.7 Behavioral Diversity  761 24.2.7.1 Sociality  762 24.2.7.2 Mimicry and Warning Coloration  762 24.2.7.3 Pollen‐Collection Strategies  763 24.3 Biodiversity Changes Through Time  765 24.3.1 Long‐Term Environmental Change  765 24.3.1.1 Mid‐Paleozoic Beginnings of Terrestrial Ecosystems  765 24.3.1.2 Initial Taxic Radiation of Insects  765 24.3.1.3 Late Paleozoic Expansion of Herbivore Functional Feeding Groups  766 24.3.1.4 Ecological and Behavioral Changes from the Mesozoic Lacustrine Revolution  767 24.3.1.5 The Parasitoid Revolution  767 24.3.1.6 Biodiversity Ramifications of the Early Expansion of Angiosperms  768 24.3.1.7 Expansion of the Grassland Biome  769 24.3.2 Short‐Term Environmental Change  770 24.3.2.1 Permian–Triassic Global Crisis and Reductions in Biodiversity  770 24.3.2.2 Cretaceous–Paleogene Global Crisis and Reductions in Biodiversity  771 24.3.2.3 Biodiversity Realignments During the Paleocene–Eocene Thermal Maximum  772 24.3.2.4 End‐Pleistocene Extinctions and Their Meaning for the Modern World  772 24.4 Current Societal Aspects of Fossil Insect Biodiversity  773 24.4.1 Human Interests and Biases  773 24.4.2 Tools for Understanding Evolutionary and Ecological Diversification  773 24.4.3 Detection of Insect‐Borne Diseases in the Fossil Record  774 24.4.4 Insect Herbivory and Global Warming  775 24.4.5 The Current Biodiversity Crisis  775 24.5 Conclusions  776 24.5.1 The Importance of the Insect Fossil Record for Understanding Insect Diversity  776 24.5.2 The Five Fundamental Types of Diversity in the Insect Fossil Record  776 24.5.3 The Effect of Long‐Term Environmental Change on Insect Diversity  776 24.5.4 The Effect of Short‐Term Environmental Changes on Insect Diversity  776 24.5.5 How Fossil Insect Biodiversity Affects Us All  776 Acknowledgments  776 References  777

Detailed Table of Contents

25

Phenotypes in Insect Biodiversity Research  789 István Mikó and Andrew R. Deans

25.1 Phenotype Data: Past and Present  789 25.2 Phenotype Data: Present and Future  791 25.2.1 Biological Ontologies  791 25.2.2 Ontologies in Biodiversity Research  792 25.2.2.1 Referencing a Glossary  792 25.2.2.2 Generating Logically Consistent Phenotypes  793 25.2.2.3 Reasoning Across Phenotype Data  794 25.3 Challenges and Future Directions  795 25.3.1 Social Challenges to “Standardization”  795 25.3.2 Ontology Development Barriers  795 25.3.3 Ontology Implementation Barriers  796 25.3.4 Phenotype Complexity  796 25.3.5 Communicating Primarily with Semantic Phenotypes  796 25.3.6 No Clearinghouse for Phenotype Data  796 25.3.7 Reasoning Challenges  797 Acknowledgments  797 References  797 26

Global Change and Insect Biodiversity in Agroecosystems  801 David R. Gillespie, Matthew J. W. Cock, Thibaud Decaëns, Philippa J. Gerard, Sandra D. Gillespie, Juan J. Jiménez and Owen O. Olfert

26.1 Global Change  801 26.2 Insect Biodiversity in Agriculture  803 26.2.1 What Do We Mean By “Biodiversity”?  804 26.3 Effects of Global Change on Biodiversity – What Do We Know?  805 26.3.1 Crop Pests and Natural Enemies  805 26.3.1.1 Distribution  805 26.3.1.2 Community Composition  808 26.3.1.3 Other Responses to Climate Change  810 26.3.2 Soil Function and Topsoil Maintenance  812 26.3.3 Implications of Global Change for Crop Pollination  814 26.3.3.1 Evidence for Importance of Biodiversity for Pollination Service to Crops  814 26.3.3.2 Expected Effects of Global Change on Pollinator Diversity – Consequences for Society  814 26.4 Island Versus Continent Contrasts  815 26.4.1 Impacts on Biodiversity of Insects in Island Agroecosystems  816 26.5 Tropical Versus Temperate Issues  818 26.5.1 Climate Tolerances in Tropical and Temperate Species  819 26.6 Some Concluding Viewpoints  822 References  823 27

Digital Photography and the Democratization of Biodiversity Information  839 Stephen A. Marshall

27.1 27.2 27.3

The Digital Insect Collection  840 Digital Images in Interactive Keys  844 Digital Photography and Taxonomic Revisions  845

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27.4 Organization of Digital Insect Collections  848 27.5 Conclusions  849 References  849 28

Bee (Hymenoptera: Apoidea: Anthophila) Diversity Through Time  851 Sophie Cardinal

28.1 Morphological Diversity  851 28.2 Behavioral Diversity: Social, Nesting, and Floral Hosts  852 28.3 Geographical Diversity  852 28.4 Evolutionary History and Diversification  853 28.5 Conclusions  863 References  864 29

Insect Biodiversity in Culture and Art  869 Gene Kritsky and Jessee J. Smith

29.1 Prehistory  870 29.2 Insects in the Ancient World  871 29.3 The Cult of Artemis: A Case Study  874 29.4 Roman Insect Art  875 29.5 Ancient China  876 29.6 Religions of India  877 29.7 Post‐Classical Era  877 29.8 The Americas  880 29.9 Modern History  882 29.10 Japanese Art  884 29.11 Language and Literature  886 29.12 Insects in Music  889 29.13 Insects in Cinema  891 29.14 Akihabara Culture: Toys, Video Games, and Anime from Modern Japan  892 29.15 Present and Future Trends in Cultural Entomology  894 29.16 The Internet Age  895 References  896

Index of Arthropod Taxa Arranged by Order and Family  899



Index of Arthropod Taxa Arranged Alphabetically  943



Index of non‐Arthropod Taxa Arranged Alphabetically  975



Subject Index  979

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List of Contributors Peter H. Adler

Sophie Cardinal

Department of Plant and Environmental Sciences Clemson University Clemson South Carolina USA

Canadian National Collection of Insects Arachnids and Nematodes Agriculture and Agri‐Food Canada Ottawa Ontario Canada

Charles R. Bartlett

Department of Entomology and Wildlife University of Delaware Newark Delaware USA

Jae C. Choe

Wesley J. Bicha

and

Oliver Springs Tennessee USA Sven Bradler

Johann‐Friedrich‐Blumenbach Institute of Zoology and Anthropology Georg‐August‐Universität Göttingen Germany Thomas R. Buckley

New Zealand Arthropod Collection Landcare Research Auckland New Zealand

Division of EcoScience Ewha University Seoul Korea

National Institute of Ecology Seocheon‐gun Maseo‐myon Geumgang‐ro Korea Matthew J. W. Cock

Centre for Agriculture and Biosciences International Egham UK David C. Culver

Department of Environmental Science American University Washington DC USA

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List of Contributors

Andrew R. Deans

Janice S. Edgerly

Thibaud Decaëns

Robert G. Foottit

Department of Entomology Pennsylvania State University State College Pennsylvania USA Centre d’Ecologie Fonctionnelle et Evolutive Montpellier Cedex 5 France Lewis L. Dietz

Department of Entomology North Carolina State University Raleigh North Carolina USA Marie Djernæs

Department of Life Sciences Natural History Museum London UK Dmitry A. Dmitriev

Illinois Natural History Survey University of Illinois at Urbana–Champaign Champaign Illinois USA Ram Keshari Duwal

Entomological Laboratory Faculty of Agriculture Kyushu University Fukuoka Japan Monika J. B. Eberhard

Zoological Institute and Museum University of Greifswald Greifswald Germany

Department of Biology Santa Clara University Santa Clara California USA Canadian National Collection of Insects, Arachnids, and Nematodes Agriculture and Agri‐Food Canada Ottawa Ontario Canada Terry D. Galloway

Department of Entomology University of Manitoba Winnipeg Manitoba Canada Philippa J. Gerard

Biocontrol & Biosecurity AgResearch Ltd. Ruakura Research Centre Hamilton New Zealand David R. Gillespie

Agassiz Research Centre Agriculture and Agri‐Food Canada Agassiz British Columbia Canada Rosemary G. Gillespie

Department of Environmental Science University of California Berkeley California USA Sandra D. Gillespie

Biology Department University of the Fraser Valley Abbotsford British Columbia Canada

List of Contributors

Fabian Haas

Gene Kritsky

Leipzig Germany

Department of Biology Mount St. Joseph University Cincinnati Ohio USA

Nate B. Hardy

Department of Entomology and Plant Pathology Auburn University Auburn Alabama USA Ian D. Hodkinson

School of Natural Sciences and Psychology Liverpool John Moores University Liverpool UK Juan J. Jiménez

Instituto Pirenaico de Ecología (IPE) Consejo Superior de Investigaciones Científicas (CSIC) Jaca Spain Matthew S. Jones

Department of Entomology Washington State University Pullman Washington USA Jeyaraney Kathirithamby

Department of Zoology University of Oxford Oxford UK Klaus‐Dieter Klass

Senckenberg Natural History Collections Dresden Museum für Tierkunde Dresden Germany

Conrad C. Labandeira

Department of Paleobiology Smithsonian Institution National Museum of Natural History Washington DC USA and Department of Entomology University of Maryland College Park Maryland USA and College of Life Sciences Capital Normal University Beijing China Seunghwan Lee

School of Agricultural Biotechnology Seoul National University Seoul Korea Renato J. P. Machado

Department of Entomology Texas A&M University College Station Texas USA Stephen A. Marshall

School of Environmental Sciences University of Guelph Guelph Ontario Canada

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List of Contributors

Luís F. Mendes

Allen F. Sanborn

Natural History and Science Museum University of Lisbon Lisbon Portugal

Department of Biology Barry University Miami Shores Florida USA

István Mikó

Department of Entomology Pennsylvania State University State College Pennsylvania USA Edward L. Mockford

School of Biological Sciences Illinois State University Normal Illinois USA Laurence A. Mound

Australian National Insect Collection, CSIRO Canberra Australia Owen O. Olfert

Agriculture and Agri‐Food Canada Saskatoon Saskatchewan Canada John D. Oswald

Department of Entomology Texas A&M University College Station Texas USA Tanja Pipan

Karst Research Institute Postojna Slovenia

Sean D. Schoville

Department of Entomology University of Wisconsin – Madison Madison Wisconsin USA Jessee J. Smith

Department of Biology Mount St. Joseph University Cincinnati Ohio USA William E. Snyder

Department of Entomology Washington State University Pullman Washington USA Hojun Song

Department of Entomology Texas A&M University College Station Texas USA Adeline Soulier‐Perkins

Département Systématique et Evolution Muséum National d’Histoire Naturelle Paris France Gavin J. Svenson

Department of Invertebrate Zoology Cleveland Museum of Natural History Cleveland Ohio USA

List of Contributors

Matthew S. Wallace

Frank Wieland

Department of Biological Sciences East Stroudsburg University of Pennsylvania East Stroudsburg Pennsylvania USA

Palatinate Museum of Natural History POLLICHIA‐Museum Bad Dürkheim Germany

Quentin D. Wheeler

Department of Environmental Science University of California Berkeley California USA

College of Environmental Science and Forestry State University of New York Syracuse New York USA

Kipling Will

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Foreword As astronomers expand the inventory of known “Earth‐like” planets  –  worlds with conditions conducive to the possibility of life – it is fair to ask: to what end? A holy grail of astronomy has long been the discovery of extraterrestrial life. As great an event as that will be, statistically speaking, it is unsurprising. With solar systems commonplace, Earth as proof positive that life is possible, trillions of stars, and billions of years since the Big Bang, life elsewhere is a virtual certainty. The scientific end, then, will be the opportunity to study the origin and diversification of life on a remote world, and to compare it with Earth. Is DNA or a DNA‐like molecule the basis of life on both worlds? Do niches similar to those found in Earth’s ecosystems exist amongst the diverse life forms on another world? Did similar precursor conditions for life exist on both planets? But to make meaningful comparisons, we shall need detailed knowledge of life on both worlds. Recent forays into extreme Earth environments, from lakes beneath the Antarctic ice sheet to hot springs and alkaline lakes, have expanded our notion of the outer limits of life, correspondingly increasing the number and kinds of worlds on which life might be possible. And Earth’s biosphere, once conceived as a thin veil of life draped over Earth’s rocky crust, is now known to be much richer and more diverse than ever imagined, from mold spores circling  the globe at 3000  m altitude to Collembola and  round worms found thousands of feet below ground in the planet’s deepest caves. Understanding life on Earth and

its history  –  creating a lasting point of comparison for the day when ETs are found and sadly, but likely, after life on Earth has been decimated – requires that we explore and document biodiversity now. Given that more than 60% of all known species are insects, they are central to mapping the biosphere. A taxonomic inventory has been underway, of course, since the time of Linnaeus, and it continues, although with less urgency and support than it deserves. As environmental and climate changes progress and extinctions accelerate, the  relevance of a species inventory becomes more painfully evident. Without a simple baseline inventory of species, our knowledge of the biosphere is inadequate for the purposes of conservation and sustainable resource management. Without access to the adaptations of species as models for innovative materials, designs, and processes, we miss out on valuable clues to new and better ways of meeting human needs sustainably. And without discovering and learning about as many species as possible, we willingly pass up the opportunity to retell the fascinating story of evolution in detail. Even if an equal level of diversity on some other planet presents a second opportunity to study the evolution of species, we should still wish to know the biodiversity of Earth. Understanding our humanity means learning the history of transformations that account for the unique combination of features we know as Homo sapiens. Every attribute we fancy as uniquely human was modified from the genes and anatomical structures of ancestral species. And in turn, those

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Foreword

ancestral attributes were similarly modified from even more remote ancestor species. To know what it is to be human, we must trace our genealogy all the way back to the first single‐ celled ancestral species. Analogies may exist on other worlds, such as paths to intelligence or holometabolism, but we are the product of one particular evolutionary history and can only fully understand ourselves in its context. Numerically speaking, the single most conspicuous feature of life on Earth is Insecta. We flatter ourselves that this is our world, but with a million named species and a conservative estimate of at least three times that number awaiting discovery, Earth is without question the planet of the insects. Insects are a conspicuous component of nearly every terrestrial and freshwater ecosystem, the open oceans being one of few exceptions to the adaptive reach of the class. Nominally present in Antarctica, the insect fauna is species‐rich on all other continents. Great Britain is one of few places on Earth where most insect species are documented and is thus an exception, for nearly anywhere else you visit on the planet, great numbers of insects remain to be discovered. Even in Europe, several hundred new species are named annually. As a result, the biosphere will not and cannot be mapped or truly understood without intensive additional entomological study. If we wish to explore and understand the origin of biological diversity, we could scarcely do better than to begin with insects. For nearly any biological phenomenon, among the insects will be found ideal model organisms and systems. Modern genetics and evolutionary developmental biology owe much to laboratory strains of Drosophila melanogaster. Life‐history stories vary from tiny mayflies living only minutes in adult form to arid‐adapted darkling beetles living up to 20 years. Insects are the source or prime examples for our theories of most modes and tempos of speciation. Time‐to‐species, measured from initial isolation of populations, for insects ranges widely from historic time scales to those measured in millions of years. And ecologically, insects have adapted to fill just

about every defined ecological niche  –  and some we have not yet imagined. It is tragic that an accelerated rate of extinction now threatens to limit what we ultimately learn from the diversity of insect species. Much of what we do not discover and document in the next couple of centuries will simply never be known. That sounds like a lot of time in a period of sound bites and tweets, but it is the blink of an eye in evolutionary, much less geologic, time. The dire consequences of missing an opportunity to explore the origins and diversity of species cannot be overstated, yet many groups of insects have few or no living experts. And funding for fundamental exploration and inventory projects is as rare as full‐time positions for taxon specialists to do so. When Darwin convinced the scientific world of the fact of organic evolution, he set the stage for two major lines of inquiry. One would lead to the discovery of the material basis and mechanisms of inheritance and natural selection, spawning a new science of genetics focused, in part, on understanding the processes of speciation. The other opened the possibility to transform taxonomy from a speculative business of asserting relatedness among species to a precise and testable science of phylogeny reconstruction. Because a century would pass before Hennig published the finer points of cladistic theory and analysis in English in the 1960s – and for other reasons, including a misunderstanding of and bias against non‐experimental approaches in science – support for expanding and accelerating taxonomic research waned after about 1940. As we take advantage of the superior models found among insects for leading‐edge experimental work in diverse fields of the life sciences, in the face of the biodiversity crisis it is imperative that we increase, too, support for scientific natural history, taxonomy, and autecology. Although entomologists in future centuries may be limited in the number of living insects they can study, we owe it to them to conserve knowledge also of as many soon‐to‐be‐extinct species as possible. With specimens, tissues, and observations preserved in natural history museums, scientists can continue to explore

Foreword

insect phylogeny and put discoveries from a  vast array of disciplines into evolutionary perspective. This volume is a milestone in the growth of knowledge of life on Earth, and hopefully the opening shot in an overdue revolution to focus scientific resources on revealing the full diversity of insects and all they can teach us about biology, evolution, adaptation, and survival. It is an ambitious snapshot of what is and is not known. Among the chapters are vignettes of insect diversity from many perspectives: ecology, geography, fossil history, and even culture  and art. And, importantly, overviews of major taxa. Analogies between knowledge of space and of insects are not new. Howard Ensign Evans’ Life on a Little‐Known Planet put our entomological ignorance on clear display at a time in the 1960s when we were first venturing into space. A lot has transpired since. Physicists now know that all the physical mass we have seen, every planet, star, and galaxy, accounts for no more than 10% of the Universe, leaving the mysterious dark matter and dark energy yet to be explored and understood. In surprising parallel, 90% of the species of Earth are also unseen and unknown to science. Whether to prepare for an eventual comparison of the origins, evolution, and diversification of life on two or more worlds, or to assure that we have sufficient options to adapt and survive on our own changing planet, a first and necessary step is a thorough exploration of biodiversity, beginning with the most diverse clade of all: the insects. It is time to mobilize an army of experts to accelerate the exploration of insect (and all) biodiversity. A number of years ago, I brought together a group of scholars to ask whether it was feasible to complete an inventory of 10 million species in 50 years. The answer was a resounding “yes” and three paths to success were mapped out. Regrettably, the annual rate of species discovery and description is little  changed since. The overdue taxonomic “moonshot” is yet to be launched, but this can change. Practically overnight we could increase the rate of description of insects by an order of

magnitude, going from an average of 7000 or so insect species per year to 70,000. Such an inventory must include detailed descriptions of species. Every available tool must be pressed into service. DNA‐based studies can quickly identify outlier populations worthy of detailed study, and associate disparate life stages of holometabolous insects so that egg, larval, and pupal characters can be added to those of adults. But we must not shy away from the hard work of taxonomy, phylogenetics, ethology, and scientific natural history. Molecular evidence alone might tell us how many species exist (I, for one, am not yet convinced of this at the species level), but it would tell us precious little about each species. For that we need descriptive taxonomy, field natural history, and comprehensive natural history collections. The question is not how many species there are but what species there are. To know and recognize each requires that we know something of what makes each species unique in comparison with all others. First and foremost this requires a  renaissance in descriptive taxonomy, but accompanied by a revival of organismal‐level biology generally and most especially a returned emphasis on scientific natural history. A completed taxonomic inventory embellished by natural history observations creates a foundation of knowledge upon which a bright future may be built: deeper understanding of the functions of ecosystems down to the species level; a vast library of inspiration for engineers and entrepreneurs to create sustainable ways to meet human needs; and early detection of invasive species and evidence of climate and environmental change, to name only a few. I commend the editors and contributors to this work for pointing us in the right direction and reminding us just how spectacularly diverse are the insects. Quentin Wheeler President College of Environmental Science and Forestry State University of New York

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Preface, Volume II This compilation, which comprises Volume II of Insect Biodiversity: Science and Society, continues the goals and strategies of Volume I. We have brought together contributors who are experts in the study of insect biodiversity and  have considered topics that complement those  of Volume I. Chapters are grouped along  the same lines as those in Volume  I, namely, “Habitats and Regions,” “Taxa,” and “Perspectives”. Part I provides further coverage of insect biodiversity in regional settings (the Arctic, Asia, and islands) and in particular habitats (agricultural crops and caves). The

treatments of taxa in Part II, combined with those of Volume I, provide a complete overview of diversity in all insect orders. Part  III includes historical, cultural, technical, and climatic perspectives on insect biodiversity. Once again, it has been particularly difficult to locate and to convince biodiversity experts to contribute to this project and to provide reviews of contributions. Simply put, insect taxonomists and biodiversity specialists are a diminished resource, and those who remain are extremely busy. We are grateful for the enthusiasm and efforts of those who were able to contribute.

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Acknowledgments We are grateful to the following scientists who graciously reviewed one or more chapters for this volume: A. G. Appel, H. Aspöck, C. S. Bazelet, E.  P. Benson, P. D. Brock, T. R. Buckley, M.  S.  Caterino, R. H. Cherry, N. Cliquennois, J. L. Cook, M. E. Dakin, R. Dallai, B. N. Danforth, L. Deharveng, L. A. Durden, L. Espinasa, O.  S.  Flint, N. M. Franz, A. N. García Aldrete, M.  Gottardo, H. H. Hobbs III, M. S. Hoddle, M. H. Hrabar, J. S. Johnston, Y. Kamimura, J. Kits, J. Kukalova‐Peck, D. W. Langor, P. G. Mason, Y. Mashimo, H. E. L. Maw, K. B. Miller, J. C. Morse,

C. A. Nalepa, L. Packer, P. Poolprasert, W.  K.  Reeves, J. M. Rivera, J. A. Rosenheim, A. M. C. Santos, M. V. Scherrer, B. M. Shepard, D.  S. Sikes, G. B. Smith, D. S. Sriviastava, R.  Thornhill, T. Uchifune, A. G. Wheeler, Jr., L. Williams III, J. J. Wilson, S. W. Wilson, and S. L. Winterton. We thank Eric Maw for his tremendous efforts in producing the indices and for correcting, formatting, and standardizing the ­figures and captions. We gratefully acknowledge the encouragement and support of the staff at Wiley, especially Sonali Melwani.

1

1 Introduction – A Brief History of Revolutions in the Study of Insect Biodiversity Peter H. Adler1 and Robert G. Foottit2 1 2

Department of Plant and Environmental Sciences, Clemson University, Clemson, South Carolina, USA Canadian National Collection of Insects, Arachnids, and Nematodes, Agriculture and Agri‐Food Canada, Ottawa, Ontario, Canada

John Platt (1964), in his iconic paper “Strong Inference,” asked “Why should there be such rapid advances in some fields and not in others?” The answer, he suggested, was that “Certain systematic methods of scientific thinking may produce much more rapid pro­ gress than others.” As a corollary to Platt’s (1964) query, we ask “Why, within a field, should there be such rapid advances at some times and not at others?” The answer, we sug­ gest, is “revolutions” – revolutions in thinking and technology. In the study of life’s diversity, what were the revolutions that brought us to a 21st‐century understanding of its largest component  –  the insects? Some revolutions were taxon specific, such as the linkage of diseases to vectors (e.g., mosquitoes), which necessitated the need to discover and understand species. Others included all insect taxa, such as the develop­ ment of light microscopy. Some were small, such as the invention of the Malaise trap. Some were mighty, such as the molecular revolution. As discovery revealed an ever‐increasing wealth of biodiversity, patterns began to emerge. The organization and explanation of these pat­ terns received quantum boosts from Carolus Linnaeus’s systems of classification and nomen­ clature, Charles Darwin’s natural explanation for species and their relationships, and Willi

Hennig’s procedural framework for inferring relationships. The revolutions of significance in  understanding biodiversity (Fig. 1.1) have, therefore, largely been those that enabled and enhanced (i) the discovery process, (ii) the con­ ceptual framework, and (iii) the management of information.

1.1 ­Discovery Perhaps the most revolutionary of all the devel­ opments that enabled the discovery of insect biodiversity was the light microscope, invented in the 16th century. The first microscopically viewed images of insects, a bee and a weevil, were published in 1630 (Stelluti 1630). Other excellent early examples of microscope‐enabled illustrations of insects, such as ants, fleas, flies, and even a fold‐out centerfold of a louse, were featured in Robert Hooke’s 1665 publication, Micrographia (Neri 2011). Improvements in magnification and resolution over the next two centuries ensured that the microscope would continue as the primary enabler of insect biodi­ versity research. By the time light microscopy had achieved its theoretical limit of resolution in the late 1800s, the study of insects and their diversity had become a well‐established enter­ prise, although still largely descriptive in nature.

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

2

Insect Biodiversity: Science and Society 1590 1600

1630 1635

Light microscope is invented

First microscopically viewed insects are illustrated France's Muséum National d'Historie Naturelle becomes first modern museum

1660

1689

First dichotomous identification key is produced

1700 1720 1734

Réné Réaumur publishes natural history monographs on insect species

1758

Linnaeus's Systema Naturae 10th edition establishes beginning of modern zoological nomenclature

1780 1800

Thomas Say publishes first comprehensive taxonomic treatment of insects in the New World

1824

Insect genitalia are introduced as taxonomic characters

1840

Chromosomes are discovered

1859

Charles Darwin's On the Origin of Species is published

1880

Taxonomic value of non-morphological characters (e.g., sound production) recognized Glass microscope slides with Canada balsam and glass coverslips are used for insects

Figure 1.1  Selected highlights in the insect biodiversity time line.

Additional developments in microscopy, includ­ ing those used routinely by researchers, such as phase‐contrast microscopy (invented in the early 1930s) and scanning electron microscopy (first commercially available in the 1960s), improved the ability to interpret, although rarely to discover, structural characters. The light microscope, however, remains the most fundamental tool in insect biodiversity research. The microscope enabled an explosion of dis­ coveries of new species and new characters that permitted refinements in classification and identification. The study of insect genitalia, for instance, would not have been possible before the microscope. The scientific value of insect genitalia was well understood by the early

1840s: “At the end of the abdomen are placed the anal appendages, an examination of which is imperative for the correct discrimination of species. Already, in 1842, Rambur had become fully alive  to the importance of these charac­ ters…” (McLachlan 1874, p. 6). As genitalia were analyzed for each group, the number of species increased. For example, the number of spe­ cies of black flies (Simuliidae) described from Linnaeus’s “backyard” (Fennoscandia) doubled in 1911, the year genitalia were introduced as taxonomic characters for the family (Lundström 1911). And following the introduction of geni­ talic characters for leafhoppers (Cicadellidae) in 1922 (DeLong 1922), the discovery of new spe­ cies surged. With the microscope came the development of new preparation and preservation techniques (Bracegirdle 1998). Glass microscope slides, ini­ tially with coverslips of mica, became dominant in the 1800s, and by the 20th century, coverslips of glass with standardized thickness became the arrangement routinely used today. The early US federal entomologist Theodore Pergande was using microscope slides with Canada balsam to preserve and study aphids as early as the 1870s (Miller and Foottit 2017). The early choice of Canada balsam, a natural product from the bal­ sam fir (Abies balsamea), as a mounting medium has ensured that slides prepared more than 100 years ago are still interpretable today. When human interests collide with insects, science progresses. Threats to food, fiber, health, and shelter have led to dramatic leaps in discovering and understanding insect biodiver­ sity. In the early 1800s, Rafinesque described 36 species of aphids, prompted by his recognition that these tiny insects are often deleterious to their host plants (Miller and Foottit 2017). Thaddeus Harris’s splendid 1841 book and sub­ sequent expanded editions provided the vade mecum for dealing with the scourges of agri­ culturally important insects and a foundation for future biodiversity exploration. In Harris’s (1841) words, “Some knowledge of the classifi­ cation of insects … seems to be necessary to the farmer, to allow him to distinguish his friends

1 Introduction

from his enemies of the insect race.” The ­agricultural ravages of Lygus, for example, even­ tually demanded deeper understanding of the pests and helped to launch the career of noted mirid specialist Harry Knight (1917), who went on to describe 1345 species of plant bugs (Schuh 1995). The year 1897 brought about a revolutionary improvement to human health and ensured that mosquitoes would become one of the taxonom­ ically best‐known groups of insects on the planet. That was the year Ronald Ross (1897) found malarial parasites in the gut of a “dappled‐ winged mosquito” (Anopheles sp.). As the focus on vectors intensified, taxonomists bore down on the question of species and their differential vectorial competency. Complexes of cryptic species eventually were revealed (Coluzzi et al. 2002). The genera with the most notorious vec­ tors, Aedes and Anopheles, became some of the taxonomically best‐known mosquitoes. At least 75 species of Anopheles are now known to trans­ mit malarial agents to humans (Foster and Walker 2009). At a finer scale, the Anopheles gambiae complex includes the most efficient malarial vectors. From genes to organisms, this species complex ranks among the most taxo­ nomically well‐studied groups of insects. The inevitable conclusion is that the degree of taxo­ nomic activity and sophistication is correlated with the severity and prevalence of disease. Society has always had its adventurous souls. Premiere among them have been the naturalists who pushed into Earth’s remote frontiers, exploring new continents, new biomes, and new habitats to collect insects (Conniff 2011). Tropical prospecting, in particular, yielded a torrent of new biodiversity, epitomized by Terry Erwin’s (1982) fogging of tropical canopy and revolutionary suggestion that the tropical rain­ forests hold tens of millions of undiscovered species. The tropics still have vast numbers of undiscovered species, but the new frontier of biodiversity exploration and discovery is in the genome. Among the chief drivers of the discovery of insect biodiversity have been the taxonomic spe­

1881 1883

Édouard-Gérard Balbiani discovers polytene chromosomes Charles V. Riley suggests the existence of what became known as cryptic species

1893

Concept of type specimens is formalized

1897 1900

Ronald Ross links malaria to mosquitoes

1905

Thomas H. Morgan initiates insect genetic studies with Drosophila

1911

Berlese funnel is invented

Charles P. Alexander describes the first of more than 10,000 crane fly species

1920

1930

Frits Zernike invents phase-contrast microscopy

1934

Malaise trap is invented

1942

Ernst Mayr introduces the biological species concept

1950

1961 1966 1970 1977 1982 1983

First edition of the International Code of Zoological Nomenclature is published Scanning electron microscopes become commercially available English-language version of Willi Hennig’s Phylogenetic Systematics is published Electronic databases, keys and computerbased analyses introduced Electrophoresis-based taxonomy of insects becomes widely used Fred Sanger introduces the chain-termination method for DNA sequencing Terry Erwin’s fogging of tropical canopy suggests 30 million insect species Polymerase chain reaction (PCR) developed by biochemist Kary Mullis

2000 2003

Genome sequenced for Drosophila melanogaster Paul Hebert and colleagues introduce the DNA barcode (mt COI gene) Massively parallel sequencing becomes readily available

2017

Figure 1.1  (Continued)

cialists who devoted their lives to their c­ entral passion – a particular group of insects. Pre‐emi­ nent was Charles P. Alexander (1889–1981), who, over a period of nearly 70 years beginning in 1911, described more than 10,000 species of crane flies, a feat that, in his own words, “seems certain it never will be done again” (Wheeler 1985). Examples of other productive specialists include the following:

3

4

Insect Biodiversity: Science and Society ●●

●●

●●

●●

●●

●●

Adolphe Hustache and Eduard Voss: more than 9000 and more than 5000 species, respectively, of Curculionidae (C. H. C. Lyal, personal communication); Max Bernhauer and Malcolm Cameron: 5251 and 4136 species, respectively, of Staphylinidae (Herman and Smetana 2001); Alexandre A. Girault: 4843 species of Chalcidoidea (Noyes 2001); Dwight DeLong: 2712 species of Cicadellidae (Dmitriev 2017); Hans Malicky: 2453 species of Trichoptera (Morse 2017); José C. M. Carvalho: 2078 species of Miridae (Schuh 1995).

Trade‐offs, however, often accompanied these herculean efforts. At times, synthesis and analysis were sacrificed, and a dearth of identi­ fication keys, illustrations, or monographs hob­ bled future workers faced with accessing the mountain of taxonomic information needed for basic identification and subsequent taxonomic work. From the ubiquitous aerial net to the more specialized variants, the development of col­ lecting tools and techniques has helped to reveal the Lilliputian world of insects. Aspirators, beating sheets, kick nets, pan traps, pitfall traps, ultraviolet lights, and the like have become a part of the entomologist’s standard field accou­ trements. Many bear the names of their inven­ tors: the Berlese funnel, invented in 1905, and its 1918 modification, the Tullgren funnel; the popular bulk‐collecting Malaise trap, invented in 1934 by the eccentric René Malaise (Sjöberg 2014); and more taxon‐specific, if not obscure, tools such as the McPhail trap, dating by name from 1933 but with roots in the 1890s (Steyskal 1977). Even in the current technological age, with hand‐held devices enabling everything from geolocation of collecting sites to instanta­ neous transmittal of information and images, the simplest tools remain the sine qua non of the field entomologist. The simplicity of these tools evokes the words of Thomas Huxley, who upon grasping the central message in Darwin’s On the

Origin of Species remarked: “How extremely stupid not to have thought of that!” (Huxley 1901). When entomologists realized that the domi­ nant human sense  –  vision  –  did not always rule in the insect world, the opportunity was set to discover a new realm of biodiversity: cryptic species. A pioneer in appreciating the existence of cryptic species was C. V. Riley, who as early as 1883 recognized that galls induced on different parts of the hackberry tree repre­ sented different species of psyllids (Wheeler et al. 2010). The limits of biodiversity discovery imposed by structural homogeneity were lifted as new character sources were explored. The value of non‐visual communication signals  – acoustical, luminescent, and chemical – in spe­ cies discovery started to become apparent in the 1860s. But it was not until the beginning of the 20th century that one of the first taxonomic decisions was made on the basis of such sig­ nals: in this case, a firefly’s flashing pattern (Lloyd 1990). Many new species, often isomor­ phic, have since been revealed on the basis of songs and drumming patterns in groups as diverse as crickets, green lacewings, and stone­ flies (Alexander 1962, Stewart and Zeigler 1984, Henry et al. 2013). Likewise, the flashing patterns of fireflies (Lloyd 1990) and chemical signals such as the sex pheromones of many insects have signaled the presence of new spe­ cies (König et al. 2015). The discovery of chromosomes in the early 1840s (Sedgwick and Tyler 1939), and particu­ larly of dipteran giant polytene chromosomes by Édouard‐Gérard Balbiani in 1881 (Zhimulev et al. 2004), provided an entirely new source of characters that could reveal biodiversity hidden beneath uniform morphology. Beginning in the “Fly Room” of Thomas H. Morgan at Columbia University in 1911, studies of Drosophila chro­ mosomes have contributed epic insights into the patterns and processes of insect biodi­versity. They played a central role in the evolutionary synthesis (Patterson and Stone 1952), provided understanding of the spectacular radiation of Hawaiian Drosophila (Carson and Kaneshiro

1 Introduction

1976), became the primary model of  modern genetics, and ushered in the genomics era (Markow and O’Grady 2007). Chromosomal studies of other insects, especially black flies, chironomids, and mosquitoes, have uncov­ ered an abundance of biodiversity. About one‐­ quarter of the described Nearctic species of Simuliidae, for instance, were discovered through studies of their polytene chromosomes (Adler et al. 2004). Molecular biology – the term can be traced to 1938 (Tabery et  al. 2016)  –  revolutionized the fields of taxonomy and systematics, and contin­ ues to fuel much of biodiversity research. Early electrophoretic approaches (e.g., Avise 1974) reached their zenith of popularity in the 1970s and 1980s before being eclipsed by more sophis­ ticated techniques. The development of the polymerase chain reaction (PCR) technique by Nobel laureate Kary Mullis in 1983, combined with the introduction of chain‐termination DNA sequencing in 1977, further enabled the progress of the molecular revolution. The sequencing of the entire genome of Drosophila melanogaster in 2000 (Adams et  al. 2000) offered a glimpse of future possibilities. Among the most promoted, and controver­ sial, revolutions in the study of biodiversity is DNA barcoding, a natural incarnation of the universal product code that was first put into commercial use for a pack of chewing gum in 1974 (Fox 2011). In essence, a short genetic sequence – a DNA barcode – typically the mito­ chondrial cytochrome c oxidase I (COI) gene, is used to discover and identify species. The COI gene was introduced in 2003 by Paul D. N. Hebert, the “father of barcoding” (Marshall 2005), and his colleagues as “the core of a global bioidentification system for animals” that would solve the species identification problem and provide insights into biodiversification (Hebert et  al. 2003). Major enterprises are now built around it, such as the Barcode of Life Data System, which has more than 5 million sequences in its database (BOLD 2014). DNA barcoding has become a routine part of taxon­ omy, but like any tool, its value is in the manner

in which it is used, the ability of the user to ask perceptive questions and extract the insights, and the recognition of its limitations. The capa­ bility of bringing ever‐more powerful, less expensive, more accessible molecular tools to bear on problems of species and their relation­ ships is advancing at unprecedented speed. Eventually, all taxonomists and systematists might have a desktop or hand‐held laboratory for the job, a trajectory reflected in the history of computers. Other novel approaches to discovering and identifying species include near‐infrared spec­ troscopy, which has its insect‐taxonomy roots in the early 1950s (Rodríguez‐Fernández et al. 2011), and cuticular hydrocarbon analysis, which began to be applied to taxonomy in the 1970s (Kather and Martin 2012). Although often profitable, these and other techniques were not routinely adopted. General access to  the necessary equipment, technology, and expertise might limit the potential of such tech­ niques and preclude them from generating the same level of species discovery and informa­ tional capacity enabled by simpler techniques.

1.2 ­Conceptual Development Modern biological classification and nomen­ clature began officially with the tenth edition of Systema Naturae (Linnaeus 1758). The ini­ tial simplicity of the Linnaean classification system for insects, based on a single source of characters  –  wings  –  was broadly appealing (Sorensen 1995). This new system provided the leap to a new paradigm, a starting point from which gradual improvements could be made. The taxonomic categories established by Linnaeus offered opportunities for investiga­ tors to specialize in particular groups of insects  –  especially, at first, Coleoptera and Lepidoptera – and generated interest in explor­ ing the globe in search of insects to classify (Sorensen 1995). Perhaps it was the diversity of insects and the variety of examples they afforded that gave

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Insect Biodiversity: Science and Society

entomologists the insight and basis to accept, promote, and contribute to Darwin’s evolution­ ary theory, following the 1859 publication of On  the Origin of Species. Early entomologists, notably Henry Bates, Charles V. Riley, Alfred Russel Wallace, and Benjamin Walsh, were among the most ardent supporters of evolution by natural selection, regularly corresponding with Darwin, who considered himself an ento­ mologist (Sorensen 1995). Darwin’s scientific revolution also elevated the prestige of ento­ mologists. Long considered little more than the  eccentric fringe of society, entomologists became some of the most qualified individuals to render judgment on the big scientific issues of the day (Sorensen 1995). The development of modern species con­ cepts went hand in hand with the discovery of new forms of insect diversity. Aristotle’s immu­ table “eidos” (form) eventually gave way to more realistic interpretations of species, most notably the biological species concept empha­ sizing reproductive isolation (Mayr 1942). Continued debate and discussion have brought the various concepts of species into sharper focus (Wheeler and Meier 2000). Yet, despite the insights that have come from these debates, much of the insect‐biodiversity community continues to describe species without articulat­ ing the concept being used. Willi Hennig, himself an entomologist (a dip­ terist), established the methodology of phyloge­ netic systematics (cladistics), finally providing a rigorous and testable framework for discovering the genealogical relationships of all organisms, which Darwin’s intellectual revolution showed must exist. Hennig’s Grundzüge einer Theorie der Phylogenetischen Systematik was written while he was a German sanitation officer in Italy during the Second World War and was pub­ lished in 1950 (Schmitt 2013). His tour de force, Phylogenetic Systematics  –  the 1966 English translation of his German book  –  became the paradigm for biological classification. The many software programs now available for phyloge­ netic analysis (Felsenstein 2017) are based on the fundamental principles presented by Hennig.

1.3 ­Information Management The need to know has always demanded effi­ cient organization, management, and retrieval of information. These demands have produced some of the standard hallmarks of biodiversity research, such as identification keys, mono­ graphs, museums, and rules and guidelines. Biologists have long appreciated the utility of an identification key. The first dichotomous key – for plants of Britain – or at least the first proof of concept, is attributed to Richard Waller in 1689 (Griffing 2011). Computer‐based interac­ tive multi‐access keys represent popular modern tools that address identification needs (Dallwitz 2000). The lack of comprehensive treatments of taxa or of regional faunas often stymied progress in understanding insect biodiversity. Thaddeus Harris’s introductory letter to America’s father of entomology, Thomas Say, dated 7 July 1823, lamented this problem: “An ardent love of Natural Science has induced me … to devote some of my leisure moments to the study of Botany & Entomology; but the want of books … has not permitted me to make any great profi­ ciency” (Weiss and Ziegler 1931). Thomas Say subsequently provided a comprehensive treat­ ment, consolidating and making accessible the current knowledge about insect species in North America. Earlier, Réné Réaumur in his six volumes (1734–1742) of insect natural history and William Kirby and William Spence in their four‐volume set of books (1815–1826) had done the same for Europe (Sorensen 1995). Say, who described more than 1000 new species, pro­ duced three volumes (1824–1828) on North American insects (Sorensen 1995). Museums, too, consolidate and organize information and make it accessible, enabling the  discovery of species within their cabinets. Museum collections are typically the sources for revisions and monographs. The first natural his­ tory museum of the modern world might have been France’s Muséum National d’Historie Naturelle, established in 1635 (Nishida 2009). In America, the distinction falls to the American

1 Introduction

Philosophical Society in Philadelphia, with roots traceable to 1770; its collection later was transferred to the Academy of Natural Sciences of Philadelphia (Simpson 1942). Today, muse­ ums worldwide hold roughly a billion insects (Nishida 2009). Prospecting among these hold­ ings is now more likely to reveal new biodiver­ sity than would collecting in many parts of the world. For generations, the Linnaean nomenclatural and hierarchical classification systems have been the heart and soul of the storage and retrieval system for communication about bio­ diversity. They have been strengthened by a set of international rules and guidelines, first for­ malized and published as the International Code of Zoological Nomenclature in 1961, but their origins can be traced back to the early 1840s. Although alternatives to the Linnaean system have been proposed, such as the PhyloCode (Rieppel 2006), the Linnaean system remains the premiere information system for the organization of life forms. The idea of type specimens, the touchstones of taxonomy, has a long history covering several centuries, although the early years were marred with loose understanding and variable use  – eventually more than 230 uses (Farber 1976). The type concept, as understood today, was refined with a proposal for standardization of terms and definitions by Oldfield Thomas (1893), including clarification of the term “type”, later given the name “holotype” by Schuchert (1897), and the introduction of the term “paratype”. Revolutions that led to enhanced discovery of biodiversity were so successful, and the current pace of discovery has increased so dramatically in recent decades, that the major impediment we now face – the so‐called “taxonomic impedi­ ment” (Taylor 1983) – is the ability to deal with the large amount of information, particularly given the ever‐diminishing personnel devoted to the task. Computerized methods and tech­ niques of the bioinformatics revolution that began around the start of the new millennium provide a mechanism to deal with the ever‐

growing mass of biodiversity data, especially sequence data (Bloom 2001, Attwood et  al. 2011), but some aspects of information man­ agement remain challenging. Premiere among these challenges is the description of new spe­ cies. The task of describing just the remaining species of the putatively well‐known North American dipteran fauna will require eight full‐ time “Alexanders,” equivalent to 560 scientific years (Thompson 1990). Yet, given the revela­ tions of DNA barcoding, which suggest an even richer North American insect fauna than previ­ ously appreciated, the estimate of eight Alexanders might be too conservative (Hebert et al. 2016). To speed the processing of new spe­ cies, proposals have been made to replace descriptions with diagnoses, particularly DNA‐ based diagnoses (Renner 2016), and even to establish new species based on photographs rather than physical specimens (Marshall and Evenhuis 2015). Although expeditious, these practices will need to be weighed against the sacrifice of extracting future information from detailed descriptions and actual physical specimens. The universalization of the world’s scientific community, beginning in the 1980s, although still often at the mercy of political winds, was enabled by advancements of the Internet and digital age, including a personal computer on the desk of nearly every entomologist and insect enthusiast. Professionals and amateurs now take for granted near‐instantaneous accessibil­ ity to colleagues, digital specimens, literature, and online language translators. Excuses for duplication of effort, such as descriptions of the same species (e.g., in the Nearctic and Palearctic regions during the Cold War), have been ren­ dered empty.

1.4 ­Conclusions The results of revolutions in the science of insect biodiversity are expressed in the follow­ ing pages by those who have benefited from, participated in, and helped drive the study of

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Insect Biodiversity: Science and Society

insect biodiversity. We emphasize that the progress of biodiversity science is a cumula­ tive process  –  well stated by Courtney and Weigmann (2016) – not a replacement process of one technology for another. Future workers are well advised to view biodiversity holistically, from molecules to organisms, drawing on all available options to discover and interpret the natural world.

­Acknowledgments We thank J. C. Morse and A. G. Wheeler, Jr, for thought‐provoking discussions and relevant literature; A. G. Wheeler, Jr, and Q. D. Wheeler for their insights on a draft of the manuscript; M. S. Caterino, C. Dietrich, and J. C. Morse for pointing us to the Staphylinidae, Auche­ norrhyncha, and Trichoptera databases, res­ pectively; and C. H. C. Lyal for providing the numbers of Curculionidae described by selected taxonomists.

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1 Introduction

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McLachlan, R. 1874. A Monographic Revision and Synopsis of the Trichoptera of the European Fauna. Part I. Napier Printers, London, UK. 46 pp. + 5 plates. Miller, G. L. and R. G. Foottit. 2017. The taxonomy of crop pests: the aphids. Pp. 627–639. In R. G. Foottit and P. H. Adler (eds). Insect Biodiversity: Science and Society. Volume 1, Second edition. John Wiley and Sons, Chichester, UK. Morse, J. C. (ed). 2017. Trichoptera World Checklist. http://entweb.clemson.edu/ database/trichopt/index.htm [Accessed 28 February 2017]. Neri, J. 2011. The Insect and the Image: Visualizing Nature in Early Modern Europe, 1500–1700. University of Minnesota Press, Minneapolis, Minnesota. 272 pp. Nishida, G. M. 2009. Museums and display collections. Pp. 680–684. In V. H. Resh and R. T. Cardé (eds). Encyclopedia of Insects. 2nd edition. Academic Press, Burlington, Massachusetts. Noyes, J. S. 2001. Interactive Catalog of World Chalcidoidea. Taxapad [CD‐ROM]. Vancouver Centre, Vancouver, Canada. Patterson, J. T. and W. S. Stone. 1952. Evolution in the Genus Drosophila. Macmillan Co., New York, New York. 610 pp. Platt, J. R. 1964. Strong inference. Science 146: 347–353. Renner, S. S. 2016. A return to Linnaeus’s focus on diagnosis, not description: the use of DNA characters in the formal naming of species. Systematic Biology 65: 1085–1095. Rieppel, O. 2006. The PhyloCode: a critical discussion of its theoretical foundation. Cladistics 22: 186–197. Rodríguez‐Fernández, J. I., C. J. B. de Carvalho, C. Pasquini, K. M. G. de Lima, M. O. Moura and G. G. C. Arízaga. 2011. Barcoding without DNA? Species identification using near infrared spectroscopy. Zootaxa 2933: 46–54. Ross, R. 1897. On some peculiar pigmented cells found in two mosquitos fed on malarial blood. British Medical Journal 2: 1786–1788. Schmitt, M. 2013. From Taxonomy to Phylogenetics—Life and Work of Willi Hennig.

Koninklijke Brill NV, Leiden, The Netherlands. xiv + 208 pp. Schuchert, C. 1897. What is a type in natural history? Science 5: 636–640. Schuh, R. T. 1995. Plant Bugs of the World (Insecta: Heteroptera: Miridae): Systematic Catalog, Distributions, Host List, and Bibliography. New York Entomological Society, New York, New York. 1329 pp. Sedgwick, W. T. and H. W. Tyler. 1939. A Short History of Science. Revised edition by H. W. Tyler and R. P. Bigelow. MacMillan Co., New York, New York. 512 pp. Simpson, G. G. 1942. The first natural history museum in America. Science 90: 261–263. Sjöberg, F. 2014. The Fly Trap. Vintage Books, New York, New York. 278 pp. Sorensen, W. C. 1995. Brethren of the Net: American Entomology 1840–1880. University of Alabama Press, Tuscaloosa, Alabama. xiv + 357 pp. Stelluti, F. 1630. Persio Tradotto in Verso Sciolto e Dichiarato da Francesco Steluti. Appresso Giacomo Mascardi, Rome, Italy. 218 pp. + index. Available online at Linda Hall Library Digital Collections. http://lhldigital.lindahall. org/cdm/ref/collection/nat_hist/id/44661 [Accessed 12 January 2017]. Stewart, K. W. and D. D. Zeigler. 1984. The use of larval morphology and drumming in Plecoptera systematics, and further studies of drumming behavior. Annales de Limnologie 20: 105–114. Steyskal, G. C. 1977. History and use of the McPhail trap. Florida Entomologist 60: 11–16. Tabery, J., M. Piotrowska and L. Darden. 2016. Molecular biology. In E. N. Zalta (ed). The Stanford Encyclopedia of Philosophy. https:// plato.stanford.edu/archives/spr2016/entries/ molecular‐biology/ [Accessed 20 December 2016]. Taylor, R. W. 1983. Descriptive taxonomy: past, present, and future. Pp. 93–134. In E. Highley and R. W. Taylor (eds). Australian Systematic Entomology: a Bicentenary Perspective. CSIRO, Melbourne, Australia. Thomas, O. 1893. Suggestions for the more definite use of the word “Type” and its

1 Introduction

compounds, as denoting specimens of a greater or less degree of authenticity. Proceedings of the Zoological Society of London 1893: 241–242. Thompson, F. C. 1990. Biosystematic information: dipterist’s ride the third wave. Pp. 179–201. In M. Kosztarab and C. W. Schaefer (eds). Systematics of the North American insects and arachnids: status and needs. Virginia Agricultural Experiment Station Information Series 90–1, Blacksburg, Virginia. Weiss, H. B. and G. M. Ziegler. 1931. Thomas Say: Early American Naturalist. Charles C. Thomas Publisher, Springfield, Illinois. xiv + 260 pp. Wheeler, A. G., Jr. 1985. Charles P. Alexander: a tribute, with emphasis on his boyhood in Fulton County, New York, and his studies at

Cornell University. Journal of the New York Entomological Society 93: 1141–1164. Wheeler, A. G., Jr, E. R. Hoebeke and E. H. Smith. 2010. Charles Valentine Riley: taxonomic contributions of an eminent agricultural entomologist. American Entomologist 56: 14–30. Wheeler, Q. D. and R. Meier. 2000. Species Concepts and Phylogenetic Theory: a Debate. Columbia University Press, New York, New York. 256 pp. Zhimulev, I. F., E. S. Belyaeva, V. F. Semeshin, D. E. Koryakov, S. A. Demakov, O. V. Demakova, G. V. Pokholkova and E. N. Andreyeva. 2004. Polytene chromosomes: 70 years of genetic research. International Review of Cytology 241: 203–275.

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Part I Habitats and Regions

15

2 Insect Biodiversity in the Arctic Ian D. Hodkinson School of Natural Sciences and Psychology, Liverpool John Moores University, Liverpool, UK

Arctic climatic regimes are among the most extreme on Earth and present formidable adap­ tive challenges for the resident insect fauna. Consensus climate‐change models predict that polar regions will be subject to greater and more rapid changes in mean temperature, particularly during the winter, than areas nearer the equator (ACIA 2005). The typically cold‐adapted Arctic insect fauna is thus increasingly facing added climatic stress, as well as competition from invading species originating in warmer regions to the south. The Arctic insect biota, however, is no stranger to change. Perhaps more than any other insect fauna on Earth, that of the Arctic has been subject to disruption, extinction, and recolonization wrought by repeated cycles of glacial advance and retreat throughout the Quaternary and Holocene geological periods. Glacial events in the Arctic have also strongly influenced faunal interchange between the Palearctic and Nearctic Regions and the devel­ opment of their respective insect faunas. The Bering Strait area has repeatedly served as a dis­ persal corridor for animals and plants between the Old and New Worlds. Consequently, a clear understanding of present‐day Arctic insect bio­ diversity is predicated on knowledge of histori­ cal events that have influenced speciation and distribution, and on the multiplicity of adap­ tations that have evolved to allow insects to survive in the unforgiving Arctic environment.

The land area north of the Arctic Circle (66°33′45.6″N) is vast, with a low human‐­ population density. Its southern treeless reaches, often referred to as the Low Arctic, comprise large portions of the continental land masses of northern Canada and Alaska, middle Greenland, northern Scandinavia, and most of northern Russia Fig. 2.1). The High Arctic comprises most of the Canadian Arctic Islands, northern Greenland, Svalbard, the Russian Arctic island archipelagos, and the northern edge of Russia around the Taimyr Peninsula. The continental areas south of the treeline, yet within the Arctic Circle, are frequently referred to as the subarctic and often support relatively open scrub/forest birch or conifer forest, also known as boreal forest or taiga. Some areas south of the Arctic Circle, notably Iceland, southern Greenland, and areas around Hudson Bay, Canada, are often included within the sub‐ or Low Arctic on the basis of their climate and vegetation. Our knowledge of the insect faunas of these regions is relatively incomplete, and the exact distribu­ tions of most species are only partially known and are often based on intensive collecting at relatively few well‐studied sites. Furthermore, there are problems of taxonomy and nomencla­ ture, with unrecognized synonymies existing among widely distributed species, and with many species yet to be formally described. The advent of DNA barcoding of species is beginning to

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

16

Insect Biodiversity: Science and Society

High Arctic Low Arctic Subarctic

Beringia Chukotka Pen.

ka as Al

on Yuk Canada

ia Siber

Wrangel Is

Alaskan N Slope McKenzie R Delta

New Siberian Is

Russia

Lena R Delta Somerset Is Churchill Cornwallis Is

Hudson Bay Lowlands Southampton Is

Devon Is

Ungava Pen.

Baffin Is Iqaluit

Ellesmere Franz Josef Land Yamal Is Pen. Moffen

Uummannaq

Kangerlussuaq Greenland

Taimyr Severnaya Zemlya

Novaya Zemlya

Svalbard Kolguev Is Kanin Pen. Zackenberg Fennoscandia Kilpisjärvi

Iceland

Figure 2.1  Map showing the major regions of the Arctic and some of the important sites named in the text. Image created by author.

address taxonomic challenges but is, at the same time, starting to reveal many unsuspected sib­ ling species. The Arctic environment imposes severe con­ straints on insect life histories, and the species present must be well adapted to the harsh con­ ditions (Bale et  al. 1997). Biodiversity is, one can argue, expressed through the range of adap­ tations displayed. Arctic winters are severe and prolonged, with unpredictable extreme events,

and are thus potentially lethal to insects. They can also cause sublethal damaging effects to insect development and reproduction. Arctic summers are short and cool, providing a lim­ ited thermal budget for insect growth and development, thereby restricting rates of popu­ lation increase (Bale et  al. 1997, Hodkinson 2005). Microclimates can be highly variable in both space and time. During summer, extreme daily temperature fluctuations can demand an

2  Insect Biodiversity in the Arctic

exceptionally wide range of thermal tolerance. The seasonal photoperiodic regime changes from one of complete winter darkness to 24‐ hour summer daylight, denying insects the photoperiod cues they may require to synchro­ nize their seasonal or diurnal cycles (Hodkinson 2005). The Arctic is an area of low precipita­ tion, and summer desiccation can be a risk, par­ ticularly for species with soil‐dwelling larvae. Low plant diversity and poor plant‐food quality provide restricted opportunities for specialist herbivorous insects, which face the additional problem of synchronizing their life histories with the growing and flowering period of their host (Danks 1981). To survive and flourish in these harsh and unpredictable Arctic environments, insects require high adaptive flexibility across a range of charac­ teristics, most notably their morphology, behavior, ecophysiology, and use of food resources. Their reproductive strategy and dispersal characteristics need to be sufficiently robust to recover from potentially high mortality and to recolonize areas following localized extinction (Downes 1965; Hodkinson and Wookey 1999; Danks 2004, 2006; Hodkinson 2005). Although the Arctic may be less species‐rich than most major world biomes, some individual insect species attain huge population densities. During the early summer growing season, mos­ quito and black fly populations reach plague densities in some areas of the Arctic and cause severe torment to humans, mammals, and birds by removing large quantities of blood (e.g., Witter et  al. 2012). Outbreaks of Lepidoptera larvae, causing widespread defoliation and tree death, have also become a common phenome­ non in northern Scandinavia (e.g., Jepsen et al. 2008). This chapter explores Arctic insect biodiver­ sity in its broadest sense, building on the infor­ mation presented in the Arctic Biodiversity Assessment of 2013 (Hodkinson et al. 2013). It begins by considering the increasing impor­ tance of DNA barcoding studies to our under­ standing of the diversity of insects that inhabit the Far North. It then summarizes, where

­ ossible, the number of recognizable species in p the various regions of the Arctic, and identifies the dominant insect groups present. The broad geographical distribution patterns of the spe­ cies are then examined, and the relationship between latitude and species diversity in the Arctic is explored. The importance of historical events, associated with glacial maxima and minima, in determining these present‐day geo­ graphical patterns of biodiversity is discussed. At the local habitat and landscape scales, the role of spatial heterogeneity in physical and biotic factors as determinants of local patterns of insect biodiversity is surveyed, together with the diversity of adaptive features that match the species to the Arctic environment template. The chapter concludes by linking Arctic insect biodiversity with integral ecosystem character­ istics, such as food‐web complexity and polli­ nation networks, and their likely response to a changing climate.

2.1 ­Documenting Biodiversity – Traditional Taxonomy Versus DNA Barcoding Most of our knowledge of the species diversity and distribution of Arctic insects is based on the morphological species concept and traditional methods of taxonomy. With the decline in the number of specialist taxonomists, progress toward a comprehensive taxonomy of Arctic species has been slow and fraught with difficul­ ties, particularly with respect to the validity of old records and unrecognized synonymies, not to mention the many undescribed species in diverse and poorly studied families such as the Chironomidae, Ceratopogonidae, Cecidomyi­ idae, and Mycetophilidae (Ekrem et  al. 2012, Stur and Borkent 2014). Over the past 15 years, the search for a simple and repeatable method of characterizing taxonomic units has led to the development of genetic profiling to produce simple yet characteristic DNA barcodes for many individual insect species. The relatively

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Insect Biodiversity: Science and Society

low species richness of the Arctic fauna lends itself to this approach, which can be used to compare species integrity and distribution across the wide circumpolar region. Molecular data from the barcode region can also be used to examine the relationships among taxa and to test the degree of agreement between tradi­ tional and molecular taxonomies. In the following discussions, I have stretched the limits of the Arctic a little southward to include the subarctic area around Churchill on Hudson Bay, Canada, as many of the extensive barcoding studies from there include taxa whose ranges extend well into the Arctic. A similar recent and extensive Norwegian study of the insects of freshwater and humid habitats in the Finnmark region of subarctic Scandinavia also included DNA barcoding of many species. Groups for which published DNA barcode data are now available include the Diptera families Muscidae (Churchill: Renaud et  al. 2012a), Chironomidae (Svalbard: Stur and Ekrem 2011), Mycetophilidae (Finnmark: Søli and Rindal 2012), and Ceratopogonidae (Finnmark: Stur and Borkent 2014), as well as Coleoptera (Churchill: Woodcock et  al. 2013), Plecoptera (Churchill: Zhou et  al. 2009; Finnmark: Boumans and Brittain 2012), Trichoptera (Churchill: Zhou et  al. 2009, 2010), Ephemeroptera (Churchill: Zhou et al. 2009; Finnmark: Kjaerstad et al. 2012), parasitoid Hymenoptera (Churchill: Fernandez‐ Triana et  al. 2011, Stahlhut et  al. 2013), and Lepidoptera (northern Fennoscandia: Mutanen et al. 2012). DNA barcoding has already, within a short time, contributed a wealth of significant infor­ mation toward our understanding of Arctic bio­ diversity. It has allowed us to associate larval and adult forms in important groups such as the Chironomidae (Diptera) and Trichoptera (Stur and Ekrem 2011, Ruiter et al. 2013) and to asso­ ciate males and females in difficult groups of the Chironomidae (Ekrem et al. 2010). Independent support has been provided for the Linnean morphology‐based species concept in groups such as the Ceratopogonidae and Muscidae (Diptera) and Coleoptera in the Arctic, while at

the same time revealing a number of additional sibling or cryptic species not recognized by traditional methods (Renaud et  al. 2012a, Woodcock et al. 2013, Stur and Borkent 2014). Barcoding has also aided the recognition of syn­ onymies in groups such as Lepidoptera and the circumpolar distributions of several species of Plecoptera and Ephemeroptera (Boumans and Brittain 2012, Kjaerstad et  al. 2012, Mutanen et al. 2012).

2.2 ­Insect Species Diversity in the Arctic 2.2.1  Composition of the Arctic Insect Fauna

There is no comprehensive database of Arctic insects, but a wealth of information, of varying reliability, is available across a widely scattered literature and online databases, such as the checklist of non‐marine arthropods of Alaska (Alaska Entomological Society 2012). Often this literature focuses on particular groups of insects in a restricted geographical area and is poorly cross‐referenced to other areas of the Arctic and elsewhere. Diversity trends, how­ ever, are best illustrated by examples from among the most geographically broad and inclusive studies. Despite its drawbacks, Danks’s (1981) monumental catalog of the Arctic arth­ ropods of North America (Table 2.1) still remains by far the most wide‐ranging and com­ prehensive summary of any Arctic insect fauna on a continental scale. This catalog is, however, now dated with respect to both nomenclature and species completeness, as Danks and Smith (2009) readily acknowledge, but nevertheless it serves usefully to illustrate general diversity trends. Since the compilation of the Arctic Biodiversity Assessment (Hodkinson et al. 2013), which made an initial attempt to summarize Arctic invertebrate diversity, further good sum­ maries of regional faunas have been published for Greenland (Böcher and Kristensen 2015), the High Arctic islands of the Barents Sea (Table  2.2) (Coulson et  al. 2014), and for

2  Insect Biodiversity in the Arctic

Table 2.1  Comparison of insect species diversity expressed as the number of genera and species of insects in each family in the High and Low Arctic Regions of North America. Order

Arctic families

Ephemeroptera

Metretopodidae

Odonata

Plecoptera

Orthoptera Phthiraptera

Hemiptera

Arctic genera

Low Arctic species

High Arctic species

1

1

0

Baetidae

1

7

0

Heptageniidae

1

1

0

Leptophlebiidae

1

1

0

Ephemerellidae

1

1

0

Aeshnidae

1

4

0

Coenagrionidae

1

1

0

Corduliidae

1

1

0

Pteronarcidae

1

1

0

Chloroperlidae

3

3

0

Perlodidae

5

5

0

Perlidae

2

2

0

Capniidae

1

6

0

Nemouridae

3

5

0

Acrididae

3

4

0

Philopteridae

21

37

23

Trichodectidae

1

1

0

Menoponidae

7

10

5

Ricinidae

1

2

2

Echinophthiriidae

2

2

2

Linognathidae

1

1

0

Pediculidae

1

1

0

Hoplopleuridae

2

2

1

Polyplacidae

1

2

0

Lygaeidae

1

1

0

Miridae

4

8

0

Anthocoridae

1

1

0

Saldidae

4

9

1

Corixidae

2

3

0

Cicadellidae

7

9

0

Delphacidae

1

1

0

Psyllidae

2

9

0

Aphididae

17

20

3

Coccidae

1

1

0

Ortheziidae

1

1

0

Pseudococcidae

3

2

1 (Continued)

19

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Insect Biodiversity: Science and Society

Table 2.1  (Continued) Order

Arctic families

Thysanoptera

not stated

Neuroptera Coleoptera

Diptera

Arctic genera

Low Arctic species

High Arctic species

3

2

1

Chrysopidae

1

1

0

Hemerobiidae

1

2

0

Carabidae

16

85

1

Halipliidae

1

2

1

Dytiscidae

7

24

2

Hydrophilidae

2

6

0

Silphidae

3

3

0

Staphylinidae

17

23

4

Byrrhidae

3

5

0

Bupestridae

1

1

0

Elateridae

2

7

0

Cantharidae

2

2

0

Dermestidae

1

1

0

Cucujidae

1

1

0

Coccinellidae

5

6

0

Latridiidae

2

2

1

Cerambycidae

5

5

0

Chrysomelidae

6

13

0

Curculionidae

9

14

1

Trichoceridae

1

5

2

Tipulidae

13

52

9

Dixidae

1

1

0

Chaoboridae

2

2

0

Culicidae

2

17

3

Simuliidae

6

28

0

Ceratopogonidae

4

4

3

Chironomidae

62

159

93

Bibionidae

1

1

0

Scatopsidae

2

3

0

Mycetophilidae

9

17

9

Sciaridae

4

3

5

Cecidomyiidae

2

2

2

Rhagionidae

2

2

0

Tabanidae

1

4

0

Empididae

4

20

7

Dolichopodidae

7

31

2

2  Insect Biodiversity in the Arctic

Table 2.1  (Continued) Order

Siphonaptera

Lepidoptera

Arctic families

Arctic genera

Low Arctic species

High Arctic species

Platypezidae

1

1

0

Phoridae

2

4

1

Syrphidae

13

21

6

Pipunculidae

1

1

0

Micropezidae

2

3

0

Piophilidae

4

7

5

Acartophthalidae

1

1

1

Agromyzidae

7

18

5

Milichiidae

1

2

1

Sciomyzidae

4

6

0

Heleomyzidae

5

9

1

Sphaeroceridae

2

3

0

Drosophilidae

2

2

0

Ephydridae

5

10

2

Chloropidae

2

2

0

Scathophagidae

9

28

5

Anthomyiidae

19

138

7

Muscidae

25

166

21

Calliphoridae

12

12

4

Oestridae

2

3

0

Sarcophagidae

1

1

0

Tachinidae

8

8

6

Pulicidae

1

1

1

Leptopsyllidae

2

2

0

Ceratophyllidae

5

9

3

Incurvariidae

1

1

0

Gelechiidae

1

1

0

Plutellidae

1

1

0

Tortricidae

10

19

3

Hesperiidae

2

2

0

Papilionidae

2

3

0

Pieridae

4

13

2

Lycaenidae

5

5

2

Satyridae

3

17

0

Nymphalidae

7

12

3

Pterophoridae

3

3

1 (Continued)

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Insect Biodiversity: Science and Society

Table 2.1  (Continued) Order

Trichoptera

Arctic families

Arctic genera

Low Arctic species

High Arctic species

Pyralidae

7

7

0

Geometridae

16

24

2

Sphingidae

1

1

1

Lymantriidae

1

2

2

Arctiidae

4

5

0

Noctuidae

15

28

5

Rhyacophilidae

1

1

0

Glossosomatidae

1

1

0

Hydroptilidae

1

1

0

Phryganeidae

2

2

0

Brachycentridae

1

2

0

Limnephilidae

9

15

1

Leptoceridae

1

1

0

Symphyta

Tenthredinidae

9

39

8

Siricidae

2

2

0

Parasitica

Braconidae

10

14

3

Ichneumonidae

78

131

35

Mymaridae

1

1

0

Hymenoptera

Aculeata

Eulophidae

1

1

1

Encyrtidae

3

2

2

Pteromalidae

4

2

2

Chalcididae

1

0

1

Figitidae

1

1

0

Alloxystidae

3

4

0

Cynipidae

1

1

0

Proctotrupidae

2

2

0

Diapriidae

1

1

0

Scelionidae

1

1

0

Platygastridae

1

1

0

Ceraphronidae

1

1

0

Formicidae

1

1

0

Vespidae

1

2

0

Megachilidae

1

1

0

Apidae Total

1

12

3

677

1,567

330

Data are from Danks (1981) and should be viewed with the caveats noted in the text.

2  Insect Biodiversity in the Arctic

Table 2.2  Recorded insect biodiversity of the High‐Arctic Islands of the Barents Sea. Insect group

Svalbard including Bjørnøya

Franz Josef Land

Novaya Zemlya

Phithraptera

37

0

7

Ephemeroptera

1?

0

1

Plecoptera

0

0

3

Trichoptera

1

0

1

Hemiptera (Aphidoidea)

3

0

1

Coleoptera

19

0

28

Diptera

122

8

147+

‐ Chironomidae

66+

7

73

Siphonaptera

2

0

1

Lepidoptera

3

0

14

Hymenoptera

39

0

40

‐ Ichneumonidae

22

0

20

‐ Braconidae

5

0

4

‐ Tenthredinidae

7

0

?

‐ Apidae Total

0

0

3

227

8

243

Based on Coulson et al. (2014), with the chironomids of Franz Josef Land updated from Krasheninnikov and Gavrilo (2014). Dubious records and vagrants are excluded.

s­ ubarctic Fennoscandia (Table 2.3) (Ekrem et al. 2012). Taken together, the latter two studies, which provide strongly contrasting examples of High Arctic islands and subarctic mainland faunas, confirm and reinforce the general trends seen in the Biodiversity Assessment, while emphasizing the importance of more‐compre­ hensive faunal surveys. The Fennoscandia data in particular illustrate how intensive collecting at 107 sites in just one regional area added 505 species to the known fauna and led to the discovery of 68 undescribed species, with a further 79 identified by barcoding as perhaps warranting specific status. The Arctic insect fauna, comprising probably no more than 3000–4000 species, is relatively simple compared with that of areas closer to the equator (Chernov 1995, 2002). Just over half of the known insect orders are found in the low and High Arctic. Within large and important world­

wide insect groups such as the Coleoptera and Diptera, family representation in the Arctic is just 10% and 30%, respectively (Chernov 2002). The Carabidae are among the most diverse groups of Coleoptera, but generic representa­ tion in the Arctic is around 0.5%. For the North American Arctic, Danks (1981) lists 144 insect families, 677 genera, and more than 1500 spe­ cies. Of the recorded Arctic species, only around 20% can be considered true Arctic insects, as the range of the remaining species often extends well beyond the Arctic, including many spe­ cies  that display an Arctic–alpine distribution (Chernov 2002). The proportion of true Arctic species, however, increases significantly in the polar desert region closer to the North Pole, with several species occurring beyond 80°N, such as the staphylinid beetle Micralymma brev­ ilingua at 81°09’N on Komsomolets Island, Severnaya Zemlya (Makarova et al. 2007). Fewer

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Insect Biodiversity: Science and Society

Table 2.3  Recorded insect species diversity of aquatic and humid habitats in subarctic Finnmark, Scandinavia. Insect group

Diptera

Total species

New record for Finnmark

New species

969

453

68 (79)

‐ Bolitophilidae

6

6

0

‐ Chironomidae*

385

196

33 (60)

1

1

0

‐ Diadocidiidae ‐ Mycetophilidae*

277

182

20 (19)

‐ Psychodidae*

18

11

1

‐ Ceratopogonidae*

54

54

14

‐ Chaoboridae

3

3

0

‐ Empidoidea

225

?

0

Ephemeroptera

39

2

0

Heteroptera

49

16

0

Megaloptera

4

0

0

Neuroptera

22

7

0

Plecoptera

32

0

0

Trichoptera Total

126

27

0

1,241

505

68 (79)

Based on a summary table from Ekrem et al. (2012), with the addition of data on Ceratopogonidae, Empidoidea, Psychodidae, and Chaoboridae from Stur and Borkent (2014), Jonassen et al. (2013), Kvifte and Andersen (2012), and Andersen and Kvifte (2012). Some total figures have been revised using data from Andersen and Hagenlund (2012), Søli and Rindal (2012), Greve and Andersen (2012), Roth and Coulianos (2014), and Boumans and Brittain (2012). The new species figure in parentheses is an estimate of the additional possible new species suggested by DNA barcode analysis. * Includes barcode studies.

than 40% of the insect families in the Low Arctic occur in the High Arctic (Danks 1981). Some workers have thus argued that the composition of the Arctic insect fauna represents a diverse collection of evolutionary lines that show serial impoverishment with increasing latitude. Chernov (2002) argued, much more interestingly from a biodiversity perspective, that the fauna demonstrates the ability of many distinct evolu­ tionary lines to adapt independently to the constraints of life in the Arctic. He contends that the Arctic fauna should be considered distinc­ tive,  with its own characteristic composition related to the adaptive success of the constituent species. A conspicuous feature of Arctic insect fau­ nas,  which reflects this adaptational trend, is

the  proportional reduction in the number of herbivorous and other terrestrial species with increasing latitude, and a corresponding in­­ crease in the proportion of species associated with aquatic and wet habitats Fig. 2.2) (Danks 1981, 1992b). This feature might relate to the thermal buffering provided by aquatic habitats or to the potential problems of summer desicca­ tion among terrestrial insects in the relatively dry polar desert of the High Arctic, where despite low annual precipitation, streams, lakes, and ponds still persist. Throughout the Low and High Arctic, permafrost impedes drainage, leading to the creation of abundant wet or aquatic habitats. The most species‐rich and dominant groups in the insect fauna of the low and High Arctic

2  Insect Biodiversity in the Arctic

Figure 2.2  The changing relative percentages of herbivorous, aquatic, and other terrestrial insect species groups, with respect to increasing climate severity in the Arctic regions of North America (based on Danks 1992b; redrawn from Hodkinson et al. 2013). At high latitudes, aquatic species are predominantly Diptera with aquatic stages and water beetles.

100 Herbivorous

Insect fauna (%)

80

60 Other terrestrial 40

20

Aquatic

0 Temperate

Low Arctic

High Arctic

are biting and sucking lice (Pthiraptera); aphids (Aphidoidea); the beetle families Carabidae, Staphylinidae, Dytiscidae, Chrysomelidae, and Curculionidae; and several dipteran groups, most notably the Chironomidae, Tipuloidea, Mycetophilidae, Simuliidae, Empididae, Doli­ chopodidae, Syrphidae, Scathophagidae, An­­ thomyiidae, and Muscidae. Among Lepidoptera, the dominant families are the Noctuidae and Geometridae, and within the Hymenoptera, the sawflies belonging to the Tenthredinidae and parasitoids of the family Ichneumonidae are the most species rich (Danks 1981, Brodo 1990, Chernov 1996, Coulson 2000, Konst­ antinov et  al. 2009, Hodkinson et  al. 2013, Chernov et  al. 2014, Böcher and Kristensen 2015). These trends tend to recur repeatedly across the different sectors of the Arctic. Not all these groups, however, are represented in the High Arctic, where the fauna is further impoverished. Here, beetles of the families Chrysomelidae, Staphylinidae, and Latridiidae (minute scavengers), although species‐poor, form an increasing proportion of the coleop­

Extreme Arctic

teran fauna, and the Diptera are increasingly represented by the Chironomidae (Chernov et al. 2000, 2001, 2014; Olsvik et al. 2001; Chernov 2002; Chernov and Makarova 2008; Coulson et al. 2014). Ichneumonid wasps are often sur­ prisingly speciose in the High Arctic. 2.2.2  Species Richness Trends Along Latitudinal Gradients

There is a broad general trend for insect diver­ sity in the Arctic to decrease progressively with increasing latitude as decreasing temperature acts to reduce the number of species able to complete their life cycle. Such trends have been widely observed in several well‐represented insect groups. Among butterflies, for example, in both Middle Siberia and Beringia, the num­ ber of species declines from 30–40 at southern sites with a mean July temperature of 10–15 °C to fewer than five species at northern sites where mean July temperature falls below 5 °C (Chernov 1995). The carabid beetles of the Taimyr Peninsula show a similar trend along

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a  latitudinal transect spanning a mean July temperature gradient of 12.5–4.0 °C, with the number of species declining from 59 to three (Chernov 1995, Chernov and Makarova 2008, Chernov et  al. 2014). This declining pattern is repeated among ground beetles in Arctic Alaska, Canada, and Norway (Nelson 2001, Olsvik et  al. 2001, Ernst and Buddle 2015). Arctic chyrsomelid beetles number around 40 species, but this number declines to around four truly Arctic species at the most northerly sites (Chernov et al. 1994, Mededev 1996, Makarova et  al. 2007). Bumblebee species in European Russia decline from around 30 to just four Arctic species (Potapov et  al. 2014). Physically smaller beetle taxa, such as the Chrysomelidae and Staphylinidae, seem better able to offset the effects of lower temperatures, and consequently their diversity tends to fall off more gradually with increasing latitude (Sokolov 2003, Chernov et  al. 2014). Latitudinal temperature anoma­ lies  occasionally produce deviation from the expected typical south‐to‐north pattern. Lake‐ inhabiting chironomid‐community compo­ sition in the Canadian Arctic Islands, for example, tracks the existing temperature anom­ aly between the relatively warmer northern and  southern islands and the cooler central islands, such as Cornwallis and Devon (Gajewski et al. 2005). Not all insect groups respond to temperature gradients in this way. Sawflies (Hymenoptera) are a notable exception, with species richness increasing in the Low Arctic before eventually decreasing in the High Arctic (Kouki et al. 1994, Kouki 1999). This trend is related to the diver­ sity and abundance of their host plants, pre­ dominantly willow (Salix species), a genus that exhibits peak diversity at high northern lati­ tudes. The Arctic sawflies, however, are largely restricted to the family Tenthredinidae, with the High Arctic species confined within the sub­ family Nematinae (Roininen 2002, Hjältén et al. 2003). The composition of many specialist insect‐ herbivore communities along latitudinal gra­ dients in the Arctic is determined by the

availability of suitable host plants and the ability of species to synchronize their life histories with the correct phenological state of their hosts (Høye and Forchhammer 2008). Many species of psyllid, for example, are associated with woody shrubs, notably species of Alnus, Betula, Ledum, Salix, and Vaccinium, over large areas of the Low Arctic, including Alaska, Greenland, Russia, and Scandinavia. In many cases, the host plants extend farther north, and sometimes farther south, than their associated psyllid (Hodkinson and MacLean 1980, MacLean and Hodkinson 1980). The absence of the insects reflects a breakdown in developmental syn­ chrony between the larval stages and the avail­ ability of rapidly growing catkins or shoots (Hodkinson et al. 1979). The host‐plant range of the psyllid Cacopsylla groenlandica, which develops on four species of Salix in southern Greenland, becomes progressively more res­ tricted farther north as some hosts reach their northern limit, and synchrony with other hosts breaks down (Hodkinson 1997). For many Arctic aphids and sawflies, precise timing of egg hatch is similarly vital to ensure synchronous and thus successful development (Gillespie et al. 2007, Barstad and Nilssen 2012). Some more southerly species, such as the but­ terfly Parnassius mnemosyne feeding on Corydalis solida in Russia, for comparable rea­ sons, reach their northern limit at or just within the Arctic Circle (Bolotov 2013). Among insect groups that feed on nectar or pollen as adults, such as the butterflies Boloria chariclea and Colias hecla in northeastern Greenland, syn­ chrony of flight activity with flower availability is essential (Olesen et  al. 2008, Høye et  al. 2014). Phenological mismatches along distri­ bution gradients can also potentially limit the extent to which different insect parasitoids exploit their hosts. The parasitoid guild of the winter moth Operophtera brumata in subarctic birch forest in North Norway, for instance, displays a well‐ordered successional pattern of oviposition by different species on specific larval instars of the host. Their populations also decline along an elevational gradient of

2  Insect Biodiversity in the Arctic

increasing climate severity (Vindstad et  al. 2011). It may be argued, however, that a diverse succession of several parasitoid species mini­ mizes the probability of a major phenological mismatch between the host and its total parasi­ toid burden throughout the season (Vindstad et al. 2011). Latitudinal biodiversity gradients have also been observed in stream and river insects in the Arctic on both local and regional scales. Within the Mackenzie River system in Canada, generic richness on a local scale tends to decrease with  increasing latitude in the Plecoptera and Ephemeroptera but to increase in the Chirono­ midae. A general decline in species richness was most apparent on the regional scale, but trends generally followed those observed at the local scale (Scott et al. 2011). 2.2.3  Geographical and Regional Variations in Species Richness

There is a general trend in insect groups for the proportion of species displaying a Holarctic distribution to increase as one moves from the subarctic, and regions to the south, to the High Arctic (Danks 1981). Broadly distributed spe­ cies with Holarctic, circumboreal, northern ­circumpolar, and Arctic cosmopolitan distribu­ tions comprise a significantly large proportion of the Arctic insect fauna. Among moths of the family Noctuidae, for example, the fauna of the High Arctic comprises 100% Holarctic species but falls to around 42% in Iceland and the Yukon (Mikkola et al. 1991). The Lepidoptera fauna of the area around Amderma on the Kara Sea, comprising 27 species, is dominated by Hol­arctic species, most of which are con­ fined to tundra habitats (Kullberg et al. 2013). Similar high proportions of Holarctic spe­ cies  (> 68%) occur among several Diptera families,  including Culicidae, Trichoceridae, ­ Calliphoridae, and Anthomyiidae, and in Arctic Hemiptera of the families Lygaeidae and Saldidae (Danks 1981, 1990; Makarova and Makarov 2006). A lower proportion of Holarctic species is found in some aquatic

orders such as the Plecoptera and Trichoptera, probably because they tend to be restricted more to the southern parts of the Arctic (Randolph and McCafferty 2005, Maka­ rova and Böcher 2009, Teslenko 2009). Despite the relatively high proportion of Holarctic species in the Far North, there are well‐documented differences in the species composition and biodiversity of insect groups in different geographical regions of the Arctic. The number of butterfly species (132 in total), for instance, varies across regions, with the highest regional total (74 species) in the East European Arctic (Chernov and Tatarinov 2006). The fauna is dominated by four genera, Colias (the yellows or sulfurs), Boloria (the fritillaries), Oenis (the graylings), and Erebia (mountain ringlets), which make up more than 50% of the species generally, but which become increas­ ingly dominant the farther north one goes. Similar variation in faunas across regions is par­ ticularly well documented among Arctic fami­ lies of the Coleoptera and Plecoptera (Chernov et  al. 1994, Chernov and Makarova 2008, Teslenko 2009, Chernov et  al. 2014, Ernst and Buddle 2015). These regional differences imply that at least part of the fauna in each Arctic region is derived separately from more south­ ern elements. The Bering Strait region, for reasons explained later, has provided a major evolutionary stage for the diversification of the Arctic biota. The strait itself has, however, proven to be less of a geographical barrier between faunas than other less obvious physiographic obstacles. Among many insect groups, there is a strong faunal dis­ junction, for example, between the eastern and western sectors of the North American Arctic (Danks 1993). Several groups show a progres­ sive decline in biodiversity from west of the Mackenzie River through the region between the Mackenzie and Hudson Bay to the area east of Hudson Bay (Danks 1981, Danks and Smith 2009). The reason is unclear but is probably linked to glacial history. Groups following this trend include beetles of the families Carabidae and Chrysomelidae; flies belonging to the

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Tipulidae, Anthomyiidae, and Muscidae; and butterflies of the families Pieridae, Satyridae, and Nymphalidae. Hymenoptera groups with strongly contrasting biology such as the her­ bivorous sawflies (Tenthredinidae), parasitoid Ichneu­monidae, and social bees (Apidae) exhibit similar trends (Danks 1981). For many of the Arctic islands, the diversity of the insect fauna is lower than in larger adjacent land areas and often reflects the capacity of the species to cross the intervening ocean barriers. Svalbard, in the High Arctic, is relatively spe­ cies‐poor, reflecting its isolated position, but nonetheless it is much richer than the even more isolated Franz Josef Land (Coulson et al. 2014). Greenland, earlier regarded as part of the Nearctic Region, is particularly interesting as its insect fauna comprises a mixture of Holarctic, Nearctic, and Palearctic elements, suggesting colonization from north, west, and east, but the proportions differ among species groups (Maka­ rova and Böcher 2009). The Trichoptera (eight species) show predo­ minantly Nearctic affini­ ties, the Coleoptera (37 species), almost exclu­ sively a mixture of Holarctic and Palearctic species, and the Lepidoptera (42 species) a mix of all three zoogeographical elements. The dip­ teran family Muscidae (37 species), by contrast, shows Holarctic and Nearctic affinities but lacks a Palearctic element. 2.2.4  Diversity Oases Within the Arctic

Within the Arctic regions, several localities sup­ port a richer insect fauna than might be expected from their northerly latitudinal posi­ tion. These thermal oases might result from particularly favorable microclimatic conditions or from the presence of warm or isothermal springs (Elvebakk 2005). They often represent the northernmost limit of species that are gen­ erally absent from the adjacent areas. Such oases often occur at sheltered south‐ or west‐facing sites, often with a reflective body of water in front and a cliff behind (Mikkola 1992). Con­ sequently, such sites occur most frequently at

the sheltered heads of fjords or adjacent to sea coasts where climate is ameliorated by a warmer ocean current. Good examples of such ento­ mological oases include Ossian Sarsfjell on Kongsfjord (Svalbard), Kilpisjärvi (Finnish Lapland), Lake Hazen and Alexandra Fjord on Ellesmere Island, and Truelove Lowland on Devon Island (all within the Canadian Arctic), and Zackenberg (northeastern Greenland) (Oliver 1963, Bliss 1987, Mikkola 1992, France 1993, Svoboda et al. 1994, Ring 2001, Høye and Forchammer 2008, Timms et  al. 2013). Ossian Sarsfjell, for example, supports the only, or most northerly, Svalbard populations of the moths Pyla fusca and Apamea zeta and the carabid beetle Amara quenseli (Coulson et  al. 2003c). The predatory dytiscid beetles Hydro­porus morio and Hydroporus polaris occur commonly in shallow ponds at Alexandra Fjord (de Bryun and Ring 1999). Similarly, Zackenberg repre­ sents the northern limit for several insect spe­ cies in northeastern Greenland. Isolated warm or isothermal spring‐fed streams and their surrounding area likewise provide locally favorable habitats for insects in a colder landscape (Bolotov et  al. 2012). On the north slope of Alaska along the northern foothills of the Brooks Range, springs with a winter temper­ ature of 4–11 °C, compared with a January aver­ age air temperature of around −29 °C, support a diverse aquatic insect fauna comprising 24 spe­ cies of Plecoptera and 33 species of Trichoptera (Kendrick and Huryn 2014). Similarly, the ther­ mophilic staphylinid beetle Omalium caesum on Svalbard is known only from the vicinity of the hot springs on Bockfjord in northwest Spitsbergen. Springs, with their surrounding vegetation and potential to support atypically diverse insect communities, are found at several other places in the Arctic, such as the Chukotka Peninsula of Russia and several sites on Greenland, including Disko Island, and along the Arctic East Coast (Katenin 2001, Roeselers et al. 2007). Nutrient‐rich areas around bird cliffs, result­ ing from accumulations of decaying organic

2  Insect Biodiversity in the Arctic

matter and bird faeces at their base, provide high spots of invertebrate activity and dense populations of prey items, such as Collembola, for predatory beetles (Zmudczynska et al. 2012). On Svalbard, the staphylinid beetles Atheta graminicola and Boreophila subplana are typi­ cally found immediately below bird cliffs (Strand 1942, Hågvar 1971). On the Brogger Peninsula on Kongsfjord, the freeze‐intolerant A. gramini­ cola is found only below cliffs on the warmer west‐facing side of the peninsula but is absent from the cliffs on the colder east‐facing side. Nutrient‐enriched soils around human settle­ ments also support an enhanced fauna. Atheta graminicola occurs in soils on slopes below cattle sheds at the Russian settlement of Barents­ burg, together with two species of the chiro­ nomid Smittia and the sciarid Lyoriella sp. (Coulson et al. 2013). Atheta graminicola is also found on the barren island of Moffen beyond 80°N (Fjellberg 1983). This occurrence might relate to a slightly warmer microclimate created by the large walrus colony. Collectively, the above sites have, because of their greater rela­ tive diversity, provided the focus for insect bio­ diversity studies in the Arctic, possibly to the neglect of other, more remote sites.

2.3 ­Historical Insect Biodiversity in the Arctic – the Time Perspective The diversity of some groups of insects, includ­ ing predatory and scavenging beetles of the families Carabidae, Dytiscidae, and Staphy­ linidae, and lake‐ or pond‐dwelling Chirono­ midae larvae, has proved particularly useful in reconstructing the past climate in several regions of the Arctic. Their subfossil remains have frequently been well preserved in perma­ frost layers, tephra, or lake sediments, and many species show little morphological change over long time periods, with some beetle species dat­ ing back virtually unchanged to the Miocene

(Elias et  al. 2006). By analyzing the thermal characteristics of the sites where these species occur today, it is possible to predict what the cli­ mate might have been like at particular times in the past, and to understand how the Arctic cli­ mate has since changed (Elias 2001). A good example of the application of such proxy data is the use of information on the thermal tolerance range for 147 beetle species to reconstruct the changing terrestrial climate and the corre­ sponding beetle diversity in Alaska and the Yukon Territory during the Pleistocene (Elias 2000a, 2000b, 2001; Elias and Mathews 2002). Deposits of subfossil chironomid head capsules have similarly proved valuable in reconstructing the historical temperature and nutrient pro­ files  of lakes at several sites in Arctic Canada, Svalbard, Russia, and Greenland, especially within the past 10,000 years. These studies gen­ erally show the gradual faunal shift from depau­ perate communities of cold‐adapted species living under oligotrophic conditions following the last ice age to more‐diverse recent commu­ nities typically dominated by species adapted to warmer, nutrient‐rich conditions. Often the speed of community change seems to have accelerated in the past 150–200 years (Brooks and Birks 2004, Wooller et  al. 2004, Thomas et al. 2008, Porinchu 2009). Not all chironomid communities, however, demonstrate such con­ tinuous steady climate warming. Those on Southampton Island in the eastern Canadian Arctic, contrary to the general trend, provide evidence for a Medieval Warm Period and for recent cooling (Rolland et al. 2008, 2009). Lakes on the Ungava Peninsula show a marked shift in chironomid communities occurring around 6000 years bp (Saulnier‐Talbot and Pienitz 2010). Subfossil insect communities, nevertheless, indicate that some areas of the Arctic have been much warmer in the past than they are nowa­ days. For example, chironomid data suggest that Nikolay Lake in the Lena Delta, Russia, was 2–3 °C warmer around 10,000 years bp than it is today (Alfimov et al. 2003, Andreev et al. 2004).

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Even further back in time, during the last inter­ glacial period, the beetle fauna of Jameson Land of northeastern Greenland was different from that of the present. Of 20 species present dur­ ing  the interglacial period, just six occur on Greenland today; the remainder are now pre­ dominantly Palearctic species (Bennike and Böcher 1994). The temperature at that time was around 5 °C warmer than currently. 2.3.1  Nunataks and Glacial Refugia as Generators of Biodiversity

There is debate as to whether fragmented popu­ lations of Arctic insects and other invertebrates might have survived the last glaciation in situ on nunataks: small, isolated, ice‐free areas protrud­ ing from the continental ice sheets (Böcher 2012). There is scant evidence that this was a frequent occurrence in the Arctic, although recent molecular evidence for two Arctic plants and for members of the Trechus pertyi species group of small wingless carabid beetles in the high Alps suggests this possibility should not be  immediately dismissed (Lohse et  al. 2011, Schneeweiss and Schönswetter 2011). What is not disputed, however, is that large, ice‐free ref­ ugia were present at high latitudes at the time of the last glaciation and played an important role in diversification of the arctic fauna and flora. At the times of glacial maxima, a large area of northwestern Alaska, parts of the western Yukon, and eastern Siberia remained free of ice. These areas, which have served as major glacial refuges for plants and animals, have come to be known as Beringia (Elias 2000a, 2000b; Elias et al. 2000). Furthermore, at times of glaciation, lower sea level is thought to have exposed the shallow sea bed of the Bering Strait to produce a continuous land bridge between the Palearctic (West Beringia refuge) and Nearctic (East Beringia refuge), which has served as a major dispersal corridor between the two regions (Elias and Crocker 2008). Climatic conditions in Beringia, however, have played a major role in filtering the movements of species. Recent evo­ lutionary analyses among Holarctic butterflies

of the genera Boloria and Polyommatus sug­ gest that multiple independent dispersal events between the regions, numbering seven and five, respectively, are required to explain the distribution and diversity of existing species (Simonsen et al. 2010, Vila et al. 2015). Insights into the history and diversity of the  Beringia insect fauna have been obtained from  deposits of subfossil insects (Coleoptera, Hemiptera, and Hymenoptera) well preserved in frozen organic matter or volcanic tephra in permafrost areas of Alaska, Canada, and Russia (Alfimov et al. 2003, Elias et al. 2006, Kuzmina et  al. 2008, Wooller et  al. 2011, Kuzmina and Mathews 2012). The earliest deposits date back to the late Miocene, more than 5 million years bp, and contain many species that are little changed up until today. The species composi­ tion of this ancient beetle fauna, comprising several hundred recorded species in 46 families has, however, changed significantly over time, indicating major shifts in climate (Elias 2000a, 2000b; Alfimov and Berman 2001; Kuzmina and  Mathews 2012). Miocene beetle samples suggest a much warmer environment in the Beringia region, with coniferous forest reaching to the shores of the Arctic Ocean. Associations of Arctic tundra species, indicative of marked climatic cooling, appeared in parts of Beringia around 2 million years bp, leading to the onset of the Quaternary and a noticeable decline in  insect diversity at any one locality. More recently, Holocene warming, following the end of the last glacial period, has been documented through temperature‐induced changes in the composition of subfossil chironomid midge communities of lakes in northern Alaska and western Yukon (Kurek et  al. 2009, Irvine et  al. 2012). There is also evidence that the moist condi­ tions on the Bering Land Bridge during the last glaciation differed significantly from the drier conditions in East and West Beringia, creating a filter or barrier to insect dispersal. The species of insects typically adapted for living on dry tundra steppe differed markedly on the two sides of the land bridge despite the opportunity

2  Insect Biodiversity in the Arctic

to cross. Weevils of the genus Stephanocleonus and the pill beetle Morychus viridis (Byrrhidae) were typical of West Beringia, together with the leaf beetles Chrysolina arctica, Chrysolina brunnicornis bermani, and Galerucella inter­ rupta circumdata. By contrast, the weevils Lepidophorus lineaticollis and Vitavitus thulius were typical of East Beringia. Some leaf beetles, such as Phaedon amoraciae, however, bridged the divide (Elias and Crocker 2008, Berman et al. 2011). Repeated cycles of glaciation, coupled with isolation of species in Arctic refuges, followed by range expansion as conditions warmed, has generated considerable diversity in insect fau­ nas in the Arctic and beyond. The Arctic insect fauna has a long history of instability and change. The Beringian refugia still influence insect biodiversity today, with many species dis­ playing amphi‐Beringian, East Beringian, or West Beringian distributions, and these regions represent diversity hotspots for several Arctic insect groups, including chrysomelid beetles, weevils, crane flies, and noctuid moths (Danks 1981; Kononenko et  al. 1990; Chernov and Makarova 2008; Elias 2010a, 2010b). 2.3.2 Endemism

No insect orders or families are restricted to the Arctic. Species endemic to the Arctic tend to be widely distributed across orders and families, but collectively make up just a small propor­ tion of the Arctic insect fauna. Endemics occur most  frequently in the Hemiptera, Coleoptera, Diptera, Lepidoptera, and Hymenoptera (e.g., Dubatolov and Phillip 2013, Hodkinson et  al. 2013). They are infrequent or absent among smaller aquatic orders such as the Ephemeroptera (Harper and Harper 1997; Randolph and McCafferty 2001, 2005). Incomplete taxonomic resolution, however, frequently prevents species with widely disparate distributions from being declared endemic. For example, the Arctic stonefly Amphinemura palmeni (Plecoptera) is known only from northern Finnmark, Norway, the northern Yukon, Canada and, more improb­

ably, from wooded sites in Wisconsin, USA (Boumans and Brittain 2012). At the other extreme, among known Arctic Anthomyiidae (Diptera), 68% are Holarctic, with the remainder Nearctic endemics. This high endemicity merely represents a lack of distributional data, particu­ larly from northern Russia. Other large pockets of apparent Arctic endemism, reflecting lack of information, exist for Nearctic aphids and aleo­ charine Staphylinidae (Coleoptera) (Richards 1963, Lohse et al. 1990, Hodkinson et al. 2013). DNA barcoding coupled with wider collecting should, given time, eventually resolve many of these problems (Hodkinson et al. 2013). Arctic endemic species seem to fall into two groups: those resulting from post‐glacial recol­ onization of formerly glaciated area and those derived from a fauna that survived the glacia­ tions in situ in Arctic refugia. In the former group are those species found on islands such as Svalbard and Greenland. Svalbard, for exam­ ple, has two endemic aphids with a highly local­ ized distribution: Acyrthosiphon svalbardicum feeding on Dryas octopetala and Sitobion cal­ vulus on Salix polaris (Strathdee et  al. 1993, Gillespie et al. 2007). Their host plants, by con­ trast, have much broader Arctic distributions. These aphids are, in turn, host to two endemic hymenopterous parasitoids, Diaeretellus sval­ bardicum and Aphidius leclanti (Braconidae) (Chaubet et  al. 2013). A small endemic moth, Plutella polaris, has a similar highly restricted distribution on Svalbard and is poorly known (Baraniak 2007). On Greenland, endemic Hemi­ptera include nine Aphidoidea species, two Psylloidea, and five Coccoidea; endemic Diptera include 13 Chironomidae and two Mycetophilidae; endemic Hymenoptera include eight Braconidae and 28 Ichneumonidae; and endemic Coleoptera number just a single spe­ cies of Carabidae (Böcher and Kristensen 2015). The amphi‐Beringian region, representing the old Beringia glacial refugium, especially the tundra steppe region of northeastern Siberia, is particularly rich in endemic species, notably among the Lepidoptera, Diptera (Tipuloidea), and Coleoptera (Curculionidae, Carabidae,

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Chryso­melidae, and Staphylinidae) (Kononenko et al. 1990, Mikkola et al. 1991, Ryabukhin 1999, Chernov and Makarova 2008, Konstantinov et  al. 2009, Khruleva and Korotyaev 2012, Hodkinson et al. 2013).

2.4 ­Biodiversity on the Landscape Scale 2.4.1  Variation in Biodiversity on a Landscape Scale

Arctic regions contain a diversity of terrestrial and freshwater habitats, from the birch forests of the subarctic to the polar deserts of the High Arctic, and from large permanent lakes to inter­ mittent streams. On a local scale, the nature of the ground surface can change markedly over a short distance, creating a mosaic of contrasting habi­ tats. Most Arctic insects have highly specific eco­ logical requirements that restrict their distribution across these habitats on both local and landscape scales. Some species are more broadly distributed across multiple habitat types. Individual species may differ in the range of habitats they occupy under different environmental regimes. Taken together, this ensures that dissimilar habitats tend to support insect communities of differing

s­ pecies composition, an important consideration for characterizing total biodiversity. On Svalbard, for example, the species composition and popula­ tion density of terrestrial chironomids differs among dry, mesic, and wet habitats (Sendstad et al. 1976) (Table 2.4). A similar response to habi­ tat moisture is seen in Canadian Arctic carabid beetles and Finnish “nematoceran” Diptera (Salmela 2011, Ernst and Buddle 2013). The species of Diptera that are associated with different types of decaying organic matter (Table 2.5) also tend to be habitat specific, often resulting in localized distributions. Tundra ground‐dwelling bugs (Hemiptera) of Dolgii Island in the Barents Sea show habitat selection with respect to humidity and salinity (Table 2.6), but habitat affinity changes in different parts of their broader range (Makarova and Makarov 2006). At the highest latitudes, insects may be  restricted to topographical features in the landscape that create slightly more favorable microclimatic conditions, such as well‐drained, south‐facing slopes, raised areas, and river ter­ races (Chernov et al. 2014). Areas that are snow‐free for the longest are  more favorable for the development of aphids and beetles of the families Staphylinidae, Chrysomelidae, and Latridiidae, compared with adjacent areas exhibiting late snow cover

Table 2.4  Vegetation types near Ny‐Ålesund, Svalbard, with their dominant terrestrial Chironomidae species. Density (m−2)

Dominant vegetation type

Wet or dry

Dominant terrestrial chironomid species

Dryas octopetala

Dry

Smittia extrema

26

Saxifraga oppositifolia, Cetraria delisei

Dry

S. extrema, Metriocnemus ursinus

7

Moss tundra

Moderately wet

M. ursinus, Limnophyes brachytomus, Linophyes pumilio, S. extrema

269

Deschampsia alpine

Wet

Paraphaenocladius impensus, M. ursinus, Linophyes eltoni, L. pumilio, Chaetocladius perennis

411

Carex ursina

Wet

Pseudosmittia sp., Smittia brevipennis

11

Based on Sendstad et al. (1976), with names updated from Lindegaard (1997) and Fauna Europea website (https:// fauna‐eu.org/).

2  Insect Biodiversity in the Arctic

Table 2.5  Diptera families and species involved in the decomposition of organic matter on Svalbard. Organic substrate

Important fly families and species

Dung

Sphaeroceridae (2 spp.), Scathophaga furcata (Scathophagidae)

Seaweed

Coelopidae (2 spp.), Scathophaga litorea

Soil organic matter and associated microorganisms

Terrestrial Chironomidae, Mycetophilidae (8 spp.), Scatopsidae (1 sp.), Sciaridae (14 spp.), Trichoceridae (Trichocera, 7 spp.)

Carrion

Calliphoridae including Protophormia terraenovae, Boreellus atriceps, and Cynomya mortuorum

Mixed bird cliff detritus

Heleomyzidae, especially Heleomyza borealis and Neoleria prominens

Table 2.6  Habitat preferences of Hemiptera species found on Dolgii Island (Barents Sea) in various parts of their broader distribution. Dolgii Island

Finnish Lapland

Novaya Zemlya

Chiloxanthus stellatus

MN

MN

DN

Chiloxanthus arcticus

MS, DN, DS

Calacanthia trybomi

DN, DS

Salda littoralis

MS

Nysius groenlandicus

DN

DN

Chlamydatus acanthioides

MN

MN

Species

Yakutia

Wrangel Island

Northern Alaska

MN

DN

Northern Yukon

Greenland

MN DN

DN

MN, MS

DN

DN MN DN

From Makarova and Makarov 2006. D, well‐drained; M, moist; N, non‐saline; S, saline near sea coast.

(­ Maka­ rova et  al. 2007, Avila‐ Jiminez and Coulson 2011). Similarly, on Severnaya Zemlya and Ellef Ringnes Island, the beetles Chrysomela septentrionalis (Chrysomelidae), Dienerella ele­ gans (Latridiidae), and Micralymma brevilingua (Staphylinidae) are confined to, or reach their highest density on, lemming mounds (Makarova et  al. 2007, Chernov and Makarova 2008). In tundra landscapes, such as on the north slope of Alaska, taxonomic richness and diversity in the Hemiptera, Diptera, and Hymenoptera tend to be higher in areas with shrub cover compared with open tundra (Rich et al. 2013).

In parasitoid species guilds associated with particular widespread insect‐herbivore species, such as Symphyta or Lepidoptera larvae, there can be considerable variation in species compo­ sition in both space and time (Roininen et  al. 2002, Vindstad et  al. 2010). The hymenopter­ ous  parasitoid guild associated with outbreak populations of the winter moth O. brumata on mountain birch in northern Fennoscandia, for example, comprises three Ichneumonidae, four Braconidae, and one Eulophidae. The species composition at any one point seems to be gov­ erned by the dynamic processes of large‐scale

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extinction and recolonization (Vindstad et  al. 2010). The insect communities of Arctic lakes, ponds, and streams similarly differ widely, depending on variations in habitat conditions. For lake‐ dwelling chironomids in the Canadian Arctic islands, Greenland, and Finnish Lapland, spe­ cies composition varies with respect to lake parameters such as size and depth, morpho­ metry, degree of exposure, pH, nutrient levels, and mean temperature (Nilsson and Soderberg 1996, Brodersen and Andersen 2002, Gajewski et al. 2005, Nyman et al. 2005). Even within sin­ gle lakes, there may be differences in the insect communities of the littoral and profundal zones and among different areas of the littoral zone with and without aquatic macrophytes (Beaty et al. 2006, Reuss et al. 2014). Insect communi­ ties in small seasonal pools can also vary mark­ edly. Among the 10 mosquito species associated with snow‐melt pools in Swedish Lapland, true Arctic species, such as Ochlerotatus nigripes, are confined to pools more than 400 m above sea level. The richest habitats, containing 8–10 spe­ cies, are pools without vegetation or with detri­ tus on the bottom and surrounded by Empetrum nigrum and Betula nana (Dahl et al. 2004). Arctic streams and rivers are equally variable in their nature, ranging from small spring‐fed tundra streams to major Arctic river systems. Insect communities in Arctic Alaska and Low Arctic habitats in Labrador and Quebec, for example, differ in species and functional compo­ sition among streams of different types, includ­ ing glacier streams, tundra streams, mountain streams, and spring‐fed streams, whether on lowland tundra or in the mountains (Huryn et al. 2005, Lento et  al. 2013). The main drivers of diversity are water temperature, nutrient com­ position, distance from source, channel and substrate stability, sediment load, and suscepti­ bility to freezing. Comparison of the chirono­ mid fauna of a glacier‐fed stream (Bayelva) on Svalbard with that of an adjacent stream (Londonelva) fed by rainfall and snow melt illus­ trates how these factors interact to influence

diversity. Bayelva had a higher discharge rate, a greater sediment load, and a lower mean tem­ perature. Londonelva was warmer, with a more stable substrate, higher chlorophyll‐a content, and higher conductivity (ion concentration). Chironomids in Bayelva were absent from the first 300 m downstream of the glacier snout, where Diamesia spp. first appeared, but their population densities then increased with dis­ tance. Populations in Londonelva, a mixture of Diamesia and Orthocladiinae species, remained relatively constant along its length, as did species richness. By contrast, Bayelva was species‐poor closest to the glacier but increased in diver­ sity  with distance from the glacier, matching Londonelva in species richness (24 versus 23 species) at the farthest downstream site. The species of Diamesia also differed between the streams, with Diamesia aberrata and Diamesia bohemani dominant in Bayelva and Diamesia arctica and Diamesia bertrami the main species in Londonelva (Lods‐Crozet et al. 2007). Pollution effects on the complexity of aquatic insect communities can be observed close to human habitation, even in the Arctic. Eutro­ phication and addition of metals to streams at Iqaluit, Nunavut, Canada, led to a loss of diver­ sity and a dramatic shift from a diverse benthic community containing Ephemeroptera, Ple­ ­ coptera, and Trichoptera to one dominated by chironomids of the subfamily Orthocladiinae (Medeiros et al. 2011). 2.4.2  Local Effects on Biodiversity – Predation and Natural Disturbance

In many Arctic lakes, the presence or absence of predators can determine the diversity of insect communities, although there are conflicting data as to whether this effect is positive or nega­ tive. In the Toolik Lake region of northern Alaska, lakes with lake trout or burbot, in addi­ tion to slimy sculpin, sustained a more diverse community and higher population density of chironomid species than those in which these fish were absent or in which sculpin alone were

2  Insect Biodiversity in the Arctic

present. The percentage of predaceous chirono­ mids was highest in lakes without fish (Goyke and Hershey 1992). However, in a later study in the same area, large insect predators such as Dytiscidae (Coleoptera), Corixidae (Hemiptera), and Chaoboridae (Diptera) were consistently absent from lakes with fish (Tate and Hershey 2003). In contrast with Alaska, chironomid spe­ cies richness and diversity at Senja Island, northern Norway, was highest in fishless lakes, particularly in the profundal zone (Mousavi et al. 2002). Habitat disturbance by vertebrates also can alter insect biodiversity. For example, grazing by large populations of lesser snow goose caused severe habitat degradation to the vegetation of the supratidal Hudson’s Plain salt marsh in Canada. This resulted in a shift in the composi­ tion of the chironomid communities living in the shallow pools and a loss of carabid beetle species in the surrounding areas (Milakovic et al. 2001, Milakovic and Jefferies 2003). A move toward increased species richness of chironomid com­ munities, resulting from eutro­phication of ponds by seabirds, has been noted at Cape Vera on Devon Island in Canada (Michelutti et al. 2011).

life cycles, especially at the highest latitudes, such as larvae of the moths Xestia aequaeva (Noctuidae) and Psychophora cinderella (Geo­ metridae) on Severnaya Zemlya (Makarova et  al. 2012). Often there is a distinct order of food preference, as illustrated by the four com­ mon moth species of the families Noctuidae, Arctiidae, and Lymantriidae at Meade River, Alaska (MacLean and Jensen 1985). There, deciduous shrubs are preferred, with subse­ quent choices varying among semi‐deciduous shrubs, forbs, evergreen shrubs, and graminoid species. Similar serial food preferences are recorded for the plant‐feeding bugs Nysius groen­ landicus and Chlamydatus pullus in Greenland (Böcher 1971, 1975). Even when food choice is confined to species within a plant genus, there is often a similar hierarchy of preferred hosts, such as among the Alaskan Salix‐feeding saw­ flies (MacLean and Jensen 1985). The soil‐dwell­ ing crane fly Tipula carinifrons frequently shows wide flexibility in its choice of dead or living food items, and at the northern limit of its range it feeds exclusively on the surface mat of blue‐ green bacteria (Striganova 1982). 2.5.2  Life‐History Adaptation

2.5 ­Important Characteristics of Arctic Insect Biodiversity 2.5.1  Specialist Versus Generalist Species

Arctic insects display wide diversity in the way they exploit their limited resources. Some spe­ cies, such as aphids, are highly specialized, feed­ ing on a single host plant and reliably completing their life cycles on high‐quality growing plant tissues within a short summer interval. Most Arctic species, however, face a greater level of unreliability and unpredictability in their food supply, which is often of poor nutritional qual­ ity. Survival and development thus depend on their ability to spread risk across a wide range of food resources: that is, to be a generalist. This is well‐illustrated among herbivores with extended

All Arctic insects are faced with the problem of  completing their life cycles given a highly restricted growing season and a low annual ther­ mal budget for development (Danks 1992a, 1999). Even within a single species, individuals at the northern range limit often perform less well than individuals living toward the southern range limit. Adults of the circumpolar whirligig beetle Gyrinus opacus, for example, accumulate significantly less fat and are sometimes smaller in Greenland than in Sweden (Svensson 2005). Among specialist herbivores, there is a need to synchronize life his­ tories with the time interval when their host‐plant tissues are favorable to support growth and devel­ opment (Danks 1986, Berg et al. 2008). Some spe­ cies, particularly among the Diptera, must similarly match adult feeding with flower availa­ bility. Arctic insects have overcome these prob­

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lems through a diversity of strategies that involve overwintering in one or more of the egg, larval, pupal, or adult stages (Hodkinson 2005, Danks 2007a). At one extreme, as in the aphids A. sval­ bardicum and S.  calvulus on Svalbard, develop­ ment can be rapid, with two, or exceptionally three, gene­ rations occurring per annum (Strathdee et al. 1993, Gillespie et al. 2007, Hullé et al. 2008). Many Arctic species have annual life histories synchronized with the growing period of the host plant, such as in all the host‐plant‐specific species of jumping plant lice or Psylloidea (Hemiptera), many sawflies (Hymenoptera), including Pristophora spp., and the bugs N. groenlandicus and C. pullus in Greenland (Böcher 1971, 1975; Hjältén et al. 2003; Hodkinson 2005, 2009; Barstad and Nilssen 2012). Annual life cycles are also found in some baetid mayflies and some car­ abid beetles (Filippov 2007, Giberson 2007, Andersen 2013). Life cycles longer than one year tend to be free‐running and less well‐syn­ chronized with the seasons, often involving repeated diapause; the insects involved develop where and when conditions permit and can overwinter several times. Extended life histo­ ries range from 1–2 years for some carabid bee­ tles to longer durations for many other species (Filippov 2007, Andersen 2013). Examples of  the latter include Psychophora spp. (Lepidoptera) and Chrysolina spp. (Coleoptera) on Severnaya Zemlya and Wrangel Island (2–6 years); the tipulid flies T. carinifrons (6–8 years) and Pedicia hannai (4–5 years) (Diptera) in Russia and Alaska, respectively; terrestrial chi­ ronomids of the genera Smittia, Metriocnemus, and Limnophyes (2–3 years) (Diptera) on Svalbard; and the moth Gynaephora groen­ landica (Lepidoptera) in Canada (7 years) (MacLean 1973; Sendstad et al. 1976; Lantsov 1982; Khruliova 1994; Morewood and Ring 1998; Makarova et al. 2007, 2012). Among truly aquatic species, some larger chironomids in shallow Alaskan tundra pools take a full seven years to complete development (Butler 1982). The percentage of Finnish butterfly species overwintering in the extendable larval stage

increases at the highest latitude (Virtanen and Neuvonen 1999). 2.5.3  Genetic Diversity Within Species and Groups

Although DNA barcoding is widely used to char­ acterize species, relatively few studies have exam­ ined the detailed variation and evolutionary history of insect groups in the Arctic. Arctic spe­ cies are often included as part of broader studies of widely distributed taxa. A good example is the butterfly genus Boloria (Nymphalidae), in which repeated switching of host plants away from the ancestral Violaceae emerges as an important fac­ tor leading to diversification in Arctic and alpine environments (Simonsen et al. 2010). It has been frequently presumed that species show reduced genetic diversity toward the northern part of their ranges, resulting from recent limited colonization following glacial retreat, a simplistic supposition that is now being more widely challenged (Hewitt 2004). Haplotypes of the carabid beetle Amara alpina from the Beringia refugium, for example, proved more diverse than those from the Hudson Bay region and several areas south of the Arctic Circle, reflecting the longer continuous occupa­ tion of Beringia (Reiss et  al. 1999). Similarly, Arctic sulfur butterflies (Colias spp.) exhibit high heterozygosity among the genes that code for the metabolic enzymes phospho­ glucose isomerase, phosphoglucomutase, and glucose‐6‐phosphate dehydrogenase, which is maintained even at the northern limit of their distributions (Wheat et al. 2005). Arctic members of the aquatic Ilybus angustior complex (Coleoptera: Dytiscidae), how­ ever, show relatively little variation in mitochon­ drial DNA, despite their widespread distributions. This pattern suggests a recent spread from a  restricted glacial refugium (Nilsson and Ribera 2007). 2.5.4  Reproductive Variation and Parthenogenesis

Several Arctic insects display variation in their mode of reproduction. Parthenogenesis is found

2  Insect Biodiversity in the Arctic

with greater frequency in Arctic insects than in  those from elsewhere (Danks 2004). It is widespread across Arctic insect groups, occur­ ring in Ephemeroptera (e.g., Baetis bundyae); Chironomidae (e.g., Smittia and other species) and Limoniidae (e.g., Symplecta scotica) in the Diptera; Trichoptera (e.g., Apatania zonella); Hemiptera (e.g., N. groenlandicus and several aphid, psyllid, and coccid species); Hymenoptera (both Parasitica and Symphyta); and Coleoptera (Curculionidae) (Richards 1963, Oliver and Danks 1972, Bengtson et al. 1974, Andersen and Øystein 1987, Giberson et  al. 2007, Staŕy and Brodo 2008, Böcher and Nachman 2011, Nokkala et al. 2013). Parthenogenesis, it has been argued, reduces the need for mating in harsh environments or cli­ matically unfavorable seasons while preventing the rapid selection of poorly adapted genotypes during particularly favorable years (Danks 2004). In several of the previous examples, males are absent or the ratio of males to females differs among sites. For N. groenlandicus in northern Greenland, this variation has been interpreted as asexual reproduction taking place in all‐female populations at isolated inland sites and sexual reproduction (50 : 50 sex ratio) occurring at more extensive favorable sites near the coast. However, in populations of the Vaccinium‐feeding psyllid Cacopsylla myrtilli in northern Scandinavia, which shows variation in sex ratio across its range, the situation is more complicated. The diploid males at the northern range limit produce normal haploid gametes by meiosis, but the accompany­ ing females are triploid and almost certainly par­ thenogenetic. Farther south, males produce diploid gametes, owing to asynapsis at meiosis, and are thus similarly non‐functional. Thus, although males are present in varying propor­ tions, reproduction in C. myrtilli seems to be entirely parthenogenetic (Nokkala et al. 2013). 2.5.5  A Diversity of Adaptations for Maximizing Heat Absorption

Arctic insect species often display unique ther­ mal biological adaptations. Many are small, with

a concomitant large surface area to volume ratio that facilitates the rapid absorption of available heat. Often this is accompanied by dark colora­ tion that maximizes heat gain at low ambient temperatures. Melanism is common in many Arctic groups, including Diptera, bumblebees, parasitoid Hymenoptera, Trichoptera, aphids, and butterflies such as Boloria and Colias spe­ cies. Several widely distributed species show intraspecific clinal variation in size and mela­ nism across their latitudinal distribution (Danks 1981, 2004). Dark coloration is often augmented by a higher‐than‐average degree of hairiness, a mechanism for trapping and retaining absorbed heat (Danks 1981). Numerous species display characteristic behavioral traits that maximize thermal gain, often in both larval and adult stages. Behavioral adaptations include sun‐basking by butterflies, such as Boloria and Colias species, on dark backgrounds, and by smaller insects, such as Diptera and parasitoid Hymenoptera, within sheltering flowers (Kevan 1972, Danks 1981). One of the best documented examples is the larva of the moth G. groenlandica, which dis­ plays both sun‐basking and avoidance behavior when cool winds blow. The pupae are also ori­ ented on the tundra surface to maximize heat absorption (Kevan et  al. 1982; Bennet et  al. 2000, 2003). Typical flight activity of most insect species in the High Arctic tends to be concen­ trated in a boundary layer of warmer air within a meter of the soil surface, especially when a wind is blowing (Coulson et al. 2003a). The short, cool Arctic summer, with 24‐hour daylight, necessitates that insects maximize their heat absorption on a daily basis. Most spe­ cies remain active throughout the diurnal cycle, when weather conditions permit, thereby maxi­ mizing their heat gain. It is difficult, however, to disentangle the effects of diurnal variation in temperature and light intensity, as the two are  highly correlated. Some species, including the  carabid beetles Patrobus assimilis and Notiophilus aquaticus, the terrestrial chirono­ mid Smittia extrema, the blow fly Protophormia terraenovae, and the mosquito O. nigripes, have

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weak peaks of higher activity, often around the noon solar maximum (Sendstad et  al. 1976, Danks 1981, Erikstad 1989). Other species, such as the winter crane fly Trichocera borealis, show no diurnal pattern (Syrjämäki 1968, Dahl 1970). Among Svalbard stream insects, especially chironomid larvae, peak dispersal by down­ ­ stream drift tends to occur at around midday as the stream temperature is rising (Marziali et al. 2009).

2.6 ­Cold Tolerance – a Diversity of Adaptations The ability to withstand extreme low winter temperatures and to operate effectively at low summer ambient temperatures is a particular feature of Arctic insects and a prominent ele­ ment of their evolved biodiversity. The posses­ sion of many of these adaptations, found at the physiological, biochemical, and behavioral lev­ els, sets Arctic insects apart from their equiva­ lents in temperate and tropical regions (Danks 2004). Physiological adaptations to winter tempera­ tures include the ability to tolerate freezing, the capacity to avoid freezing by supercooling the hemolymph fluids to low, sub‐zero tempera­ tures, and the capability to lose freezable body water by desiccation under freezing conditions. Many Arctic insects are freeze tolerant, often in more than one life stage. Such tolerance is well‐ known in Arctic sawflies and their hymenop­ terous endoparasitoids belonging to the families  Braconidae, Ichneumonidae, and Pteromalidae (Humble 2006). It also occurs, among others, in  the dipteran Heleomyza bore­ alis (Heleomyzidae), many Arctic Chironomidae and aquatic Empi­ didae, and the stonefly Nemoura arctica (Danks 1971, 2007b; Irons et  al. 1993; Worland et  al. 2000; Walters et  al. 2009). Heleomyza borealis, for example, freezes at around −7 °C but can survive down to −60 °C. Many Arctic insects, by comparison, are freeze‐ intolerant but are freeze‐resistant down to their supercooling point (SCP), usually between

−20  to −40 °C, before their tissues freeze and rupture, although death can occur at tempera­ tures above the SCP (Neved 1998, Sømme 1999). An extreme example is the Alaskan flat bark beetle Cucujus clavipes puniceus (Cucujidae) in which some individuals appear to survive temperatures as low as −100 °C with­ out freezing. Supercooled individuals, at such an extreme low temperature, avoid freezing by assuming a glass‐like state (vitrification) (Sformo et al. 2004). Cold survival through des­ iccation, involving the loss of most of the body water, occurs in several soil‐dwelling inverte­ brates with thin water‐permeable cuticles, including many Collembola (Ring and Danks 1994, Holmstrup et  al. 2002, Sorensen and Holmstrup 2011). Although not formally recorded, it is equally likely to occur among Arctic insects; for example, several overwinter­ ing Arctic Chironomidae appear capable of surviving desiccation as larvae in this manner (Danks 1971). Similarly, the ability of Arctic insects, as opposed to Collembola, to survive anoxia while encased in winter ice is poorly documented (Hodkinson and Bird 2004). Other winter survival strategies of arctic insects require synthesis of a wide range of protective chemicals that serve several purposes (e.g., Sømme 1999, Duman et  al. 2004). Cryoprotectants, primarily polyhydric alcohols, such as glycerol, sorbitol, ribitol, erythritol, and threitol, and C12 sugars including trehalose and sucrose lower the supercooling point and pro­ tect partially frozen tissue from ice nucleation. Thermal hysteresis or antifreeze proteins stabi­ lize the supercooled state and retard the inocu­ lation of ice crystals (Duman et  al. 2003). Ice nucleation proteins, by contrast, initiate con­ trolled and non‐damaging ice formation in freeze‐tolerant species. Several, but not all, Arctic insects display spe­ cific adaptation of their metabolism to cold summer conditions (Hodkinson 2003). Often they have significantly lower threshold temper­ atures for activity than their temperate coun­ terparts and their metabolic response rate to rising temperature, measured as the Respiratory

2  Insect Biodiversity in the Arctic

Quotient (Q10), reacts at low temperatures more rapidly than expected (Bertram 1935, Danks 1981). The rate of a chemical reaction normally doubles for every 10 °C rise in temperature (i.e., Q10 = 2). Values significantly above the expected value of two indicate an enhanced metabolic response to temperature. Such a response is found across several orders of Arctic insects. Examples include the beetles Atheta gramini­ cola (Q10 = 2.8), Isochnus flagellum (3.2), Simplocaria metallica (3.1) on Svalbard, the crane fly P. hannai (2.6) in Alaska, and the wide­ spread lepidopteran G. groenlandica (4.2–7.7) (MacLean 1973, Proctor 1977, Strømme et  al. 1986, Bennett et  al. 1999). Furthermore, some species show adaptation of their metabolic rate to minimize energy expenditure at nonpro­ ductive times for growth and development. Gynaephora groenlandica, for instance, displays markedly different metabolic rates, at the same temperature, early in the year when higher qual­ ity food is available, as compared with later in the year when food quality declines, or immedi­ ately following food ingestion as opposed to when not feeding (Bennett et  al. 1999). The cold‐adapted larvae of G. groenlandica also dis­ play reduced amounts of mitochondrial DNA in the fat‐body cells during autumn and winter, suggesting lower metabolic activity, compared with summer‐collected specimens (Levin et al. 2003). 2.6.1  Brachyptery and Wing Polymorphism

Several Arctic species are apterous or display varying degrees of brachyptery, usually seen as a mechanism for conserving energy in a harsh environment in which flight is rarely possible. In general, the proportion of brachypterous species, or individuals within a species, tends to increase with latitude (Danks 1981, Roff 1990). For exam­ ple, four beetle species living on the Severnaya Zemlya Archipelago in the Russian High Arctic, namely M. brevilingua, Chrysolina subsulcata, C. septentrionalis, and Dienerella filum, lack func­ tional wings (Makarova et  al. 2007). Arctic

Carabidae, including the genus Pterostichus, are also often flightless (Reiss et al. 1999, Chernov and Makarova 2008). Among the Tipulidae of the North American Arctic, about 25% of species show wing reduction and reduced flight capabil­ ity (Byers 1969). Some species such as Limonia lindrothi have reduced wings in both sexes, but in most species, the female is brachypterous. For Arctic beetles, there appears to be a much greater preponderance of winged species on recently colonized islands such as Svalbard and Green­ land than at sites, such as northwest Taimyr, Severnaya Zemlya, and Wrangel Island where beetles have persisted for much longer (Chernov and Makarova 2008, Chernov et al. 2014). This pattern might not  hold across all orders. The polymorphic endemic aphid S. calvulus, for example, is apterous, yet shows a highly restricted distribution on Svalbard (Gillespie et  al. 2007). Some Svalbard parasitoid species also display conspicuous brachyptery in both sexes. Diaeretellus svalbardicum (Hymenoptera: Braconidae) parasitizes A. svalbardicum, another species endemic to Svalbard, and Stenomacrus gro­enlandicus (Hymenoptera: Ichneumonidae) develops in species of Diptera. (Chaubet et al. 2013).

2.7 ­Dispersal, Immigration, and Biodiversity The biodiversity of Arctic insects strongly reflects post‐glacial recolonization, and most commu­ nities probably have yet to reach their potential equilibrium in terms of species composition. Dispersal is thus important on both local and geographical scales. Even today, glaciers are retreating in many parts of the Arctic, continu­ ally exposing fresh land surfaces on which new insect communities begin to develop, such as on the forelands of Midtre Lovenbre in Svalbard and Almajallojekna in Arctic Sweden (Hodkinson et  al. 2004, Franzen and Dieker 2014). Elsewhere, severe climate, coupled with the instability of the soil surface resulting from  disruptive frost heave, leads to the local

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e­ xtirpation of insect communities. Community composition in these areas echoes, at least in part, the relative dispersal ability of the constit­ uent species and their speed of recolonization. Wind‐blown dispersal plays a significant role in ensuring a ready supply of colonizing insects. On a local scale, even barren areas such as gla­ cier surfaces or forelands frequently receive a constant rain of wind‐dispersed insects from the surrounding area, most of which are poorly adapted for survival in situ during the early stages of community assembly, but which pro­ vide food for predatory early colonizers such as beetles and spiders (Edwards 1972, Hodkinson et al. 2002, Coulson et al. 2003a). Associations between insects with wind‐dispersed plant seeds or wind‐blown detritus can serve a similar purpose. The bug Nysius groenlandicus in west­ ern Greenland, for instance, lays eggs on the ripe achenes of Dryas integrifolia (Böcher 1975). When meteorological conditions conspire, the Arctic receives sporadic but often large influxes of wind‐blown insects from areas far to the south (Coulson et al. 2002). Many of these are vagrant species with little chance of survival but add to the recorded biodiversity of the Arctic. Records include a wide range of live sightings, which include common European butterflies such as the painted lady (Vanessa cardui), green‐veined white (Pieris napi), and Camberwell beauty (Nymphalis antiopa), and the Siberian aphid Cinara abieticola on Svalbard, several hun­ dred kilometers north of their host‐plant range (Coulson and Refseth 2004). The most notable regular adventive is the diamondback moth (Plutella xylostella) from mainland Russia, which frequently finds its way  to Svalbard in large numbers and is also known from polar desert on Severnaya Zemlya (Coulson et  al. 2002, Maka­ rova et al. 2012). It is unable to establish popu­ lations in the High Arctic, but its presence indicates meteorological conditions that might correspond with large influxes of smaller, less obvious, potentially colonizing insects into remote corners of the region. There is some evidence that migration of indigenous insects in the Arctic occasionally

boosts biodiversity in an adjacent geographical area. The sparse butterfly fauna of the Kanin Peninsula and Kolguev Island in the Russian Arctic, for example, is regularly supplemented by immigrant species from farther south (Bolotov 2013). Such migratory tendencies probably explain the live butterfly observed on the pack ice off the Siberian coast by the Jeannette Expedition in 1881 (De Long 1884).

2.8 ­Pollinator Networks and Pollinator Biodiversity In subarctic environments, pollinator networks are similar to those of the temperate and boreal zones, with bees, butterflies, noctuid moths, and hover flies prominent. The main potential pollinators of lingonberry (Vaccinium vitis‐ idaea) at sites along the Tanana River, Alaska, for example, include introduced and indigenous bees of the genera Apis, Bombus, Andrena, and Dialectus; wasps of the genus (Dolichovespula, 2 species); and hover flies including Melangyna spp. and Syrphus spp. (Davis et  al. 2003). However, farther north, the species richness and abundance of bees and butterflies declines and nectar or pollen‐feeding flies (Diptera) become increasingly important as the main pol­ linators (Kevan 1972, Pont 1993, Elberling and Olsen 1999, Larsson et  al. 2001, Kolosova and Popatov 2011, Potapov et al. 2014). The flowers of several Arctic plant species, such as Dryas octopetala, Saxifraga oppositifolia, and Ranun­ culus acris, provide sheltered basking sites for small insects but not all these plants are insect pollinated and not all the associated insect species successfully carry pollen (Kevan 1972, Totland 1996). Care is needed in interpreting associations; parasitoid wasps, for instance, are frequently found in High Arctic flowers but their role, if any, in pollination is uncertain (Klein et al. 2008). High Arctic pollination networks are often surprisingly complex. Comparative data, based on observational studies for Ellesmere Island, Canada (82°N); Zackenberg, Greenland (75°N);

2  Insect Biodiversity in the Arctic

Kangerlussuaq, Greenland (66°N); and Umman­ naq, Greenland (71°N) indicate the same general picture (Oleson and Jordano 2002, Lundgren and Olesen 2005). The number of potential pol­ linator species ranged from 26 to 91, the number of plant species visited varied from 15 to 31, and the number of insect–flower species interac­ tions were between 63 and 286. Recent analysis of pollen types collected from the bodies of insects at Zackenberg suggests that the numbers of interactions among insects and plants is sig­ nificantly higher than the aforementioned observational studies suggest (Olesen et  al. 2015). Smaller, isolated Arctic islands such as Svalbard, however, differ from some of the above sites in the absence of bees and the scarcity of Lepidoptera, and consequently probably have smaller networks. Nevertheless, many species of anthophilous insect species are present but their smaller size and lack of specialist adaptation means that their effectiveness in pollen transfer is probably lower. This situation might, in part, be compensated for by their higher abundance. Most frequently recorded among the pollen‐ carrying anthophilous Diptera species are the muscid flies, especially of the genus Spilogona (Muscidae), anthomyid flies (Anthomyiidae), hover flies (Syrphidae), fungus gnats (Myceto­ philidae and Sciaridae), mosquitoes (Culicidae), terrestrial and freshwater midges (Chirono­midae, such as Smittia), and predatory species of Empididae such as Ramphomyia spp. (Kevan 1972, Pont 1993, Elberling and Olesen 1999, Larson et  al. 2001, Olesen et  al. 2008). Less fre­ quently recorded dipteran visitors to flowers include the Calliphoridae, Tachinidae, Coelo­pidae, and Piophilidae. Flower‐feeding bugs, such as N. groenlandicus and C. pullus, might also transfer pollen (Olesen et al. 2008, Hod­kinson et al. 2013).

2.9 ­A Biodiversity Paradise for Parasites? Given the relatively low numbers of insect spe­ cies inhabiting the Arctic, a surprisingly dispro­ portionate number are parasites, as ectoparasites

of birds and mammals and ecto‐ or endopara­ sites of other invertebrates. Warm‐blooded ver­ tebrates appear to create a favorable thermal environment in which ectoparasites flourish. The flea fauna of Arctic Norway, for instance, comprises 14 species belonging to five families living on eight species of small mammals (Hastriter et  al. 2004). Two hosts, the shrew Sorex araneus and the northern red‐backed vole Clethrionomys rutilis, are each host to nine spe­ cies, suggesting that fleas spread their risks across several hosts. At higher latitudes, fleas appear less diverse with just two promiscuous species, Ceratophyllus vagabundus and Mioc­ tenopylla arctica, found in the nests of Svalbard bird species, including kittiwakes, gulls, geese, and ducks (Coulson and Refseth 2004, Coulson et al. 2009). Chewing lice (Mallophaga) are par­ ticularly diverse in the Arctic, with 12 genera and 37 species recorded from 21 bird species on Svalbard alone. Lice are often specific to par­ ticular regions of the body, and Arctic birds such as the purple sandpiper host up to four species in four separate genera (Coulson and Refseth 2004). At many sites in the Arctic, such as Ellesmere Island and Churchill in Canada, Zackenberg in Greenland, and Svalbard, parasitoid Hymeno­ ptera species, particularly the Ichneumonidae, form a disproportionately large element of the insect community, given the apparent low diver­ sity of potential host species available (Coulson and Refseth 2004, Fernandez‐Triana et al. 2011, Stalhut et  al. 2013, Várkonyi and Roslin 2013). On Svalbard, well‐represented families and their main hosts include the Braconidae (ovi­ positing in sawfly and Lepidoptera larvae, and aphids), Ceraphronidae (aphids), and Ichneumo­ nidae (spiders and flies). Among the Ichneumo­ nidae, individual species frequently specialize on selected prey. Aclastus borealis and Gelis glacialis, for instance, are spider specialists, whereas Atractodes pusillus and Stenomacrus groenlandicus breed in different species of Diptera (Coulson et al. 2003b). The depredations of ectoparasitic blood‐feed­ ing flies, primarily the many species of Culicidae,

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Simuliidae, and Ceratopogonidae that breed in the ponds, streams, and swampy areas of the tundra, have been widely chronicled by nearly every expedition that has set foot in the Arctic (Hocking 1960). The effect of insect harassment on native mammals, such as caribou, reindeer, and breeding birds, has been extensively docu­ mented and needs little elaboration here, although the relative importance of the endo­ parasitic warble flies (Oestridae), two species of which breed in caribou, might have been under­ estimated (Witter et al. 2012).

2.10 ­Biodiversity and the Changing Arctic Climate Insect biodiversity in the Arctic is responding rapidly to climate amelioration, a phenome­ non  apparent to indigenous people as well as scientists (Hodkinson et  al. 2013). One might expect that insect communities will change significantly as cold‐adapted species move northward or to higher elevations and other species move into the Arctic for the first time. Several studies, spread across numerous insect groups, document recent range changes and extension into previously unoccupied areas. Examples include butterfly species on Herschel Island (Canada), the ladybird beetle Coccinella trasversoguttata at Zackenberg in northeast­ ern Greenland, and significant shifts in the community of microgastrine parasitoid wasps (Hymenoptera: Braconidae) (about 70 spp.) over the past 50–70 years in Churchill, Mani­ toba. (Böcher 2009, Fernandez‐Triana 2011, Leung and Reid 2013). Similarly, chironomid communities in ponds on Ellesmere Island have increased significantly in species richness over the past 200 years as temperatures have risen (Quinlan et al. 2005). In the mountains of Padjelanta National Park, northern Sweden, the composition of the but­ terfly and moth community has changed since 1944, with 17 species shifting elevation down­ wards, 12 upwards, and the remaining 17 spe­ cies distributions remaining unaltered (Franzen

and Ockinger 2012). By contrast, the ichneu­ mon parasitoid community on Ellesmere Island has remained relatively stable over the past 55 years, despite a raised mean temperature and increased precipitation (Timms et  al. 2013). Similarly, the muscid fly community in Churchill changed relatively little over a recent 52‐year time span, except for aquatic and semi‐aquatic Spilogona spp. (Renaud et  al. 2012b). Surprise changes are observed even in individual species. On Svalbard, the aphid A. svalbardicum has revealed a hitherto unknown winged morph in the past 20 years, and has extended its distribu­ tion to sites previously too cold to support development (Hodkinson et  al. 2002, Simon et al. 2008, Avila‐Jimenez and Coulson 2013). Changing climate is likely to disrupt pheno­ logical synchrony in Arctic insect food webs, with consequences for overall insect biodiver­ sity at any one locality. Relationships affected will include those between insect herbivores and host plants, flowering plants and their visit­ ing and pollinating insects, and parasitoids and predators and their insect hosts. Not all species will be affected equally. Among butterflies at Zackenberg, for instance, recent warming has produced significantly earlier flight activity in the Arctic fritillary, B. chariclea, but not in the northern clouded yellow, C. hecla (Høye et  al. 2014). Disruption to parasitoid–host synchrony, resulting from differential responses to temper­ ature, might lead to population outbreaks of the host, as mentioned earlier for Scandinavian birch forests (Hance et al. 2007). Responses may be subtle. Beneficial endosymbiont populations in insect parasitoids might be more susceptible to raised temperature than their parasitoid host; in tritrophic interactions the release of volatile chemicals from plants, which attract parasitoid species to their insect‐herbivore host, is tem­ perature‐dependent, and thus their effective­ ness is likely to change (Hance et  al. 2007). Availability of suitable insect pollinators might restrict the rate of spread of plants with special­ ized floral anatomy, such as the louseworts (Pedicularis spp.), whereas widespread insect‐ pollinated Arctic plants, such as Dryas spp. and

2  Insect Biodiversity in the Arctic

Cassiope tetragona, might attract additional pollinators among the invading species (Kevan 1972, Klein et al. 2008). Arctic tundra, during summer, provides prime breeding sites for many migratory species of wading and passerine birds, such as snow buntings, purple sandpipers, dunlins, knots, phalaropes, and turnstones, which depend on a continuing supply of insects, especially flying Diptera such as Chironomidae and Tipulidae, to feed their developing young (Ridley 1980, Meltofte and Lahrmann 2006). Arctic insect species usually have short adult flight periods, and a seasonal succession of many species is usually required to meet the birds’ require­ ments. The reliability of the food supply thus depends on the availability of an appropriate diversity and abundance of emerging insect species throughout the nesting season, which in turn is related to temperature (Bolduc et al. 2013). In areas such as Svalbard and Bylot Island in the Canadian Arctic, warmer sum­ mers can lead to a telescoping effect in which total insect emergence is shifted or compressed into a narrower time interval, leading to pheno­ logical asynchrony with the demands of the feeding birds (Hodkinson et al. 1996, McKinnon 2012). However, at Taimyr, Russia, peak insect abundance seems to shift toward the opti­ mum  breeding date for the birds (Tulp and Schekkerman 2008). Climate change has important conservation implications for the biodiversity of cold‐adapted insect species in the Arctic. As climate warms, their habitat disappears and they must retreat to colder areas farther north or at higher eleva­ tions, otherwise extinction looms (Hope et  al. 2013). A good example is the disappearance of  cold‐adapted chironomids of the genera Oliveridia/Hydrobaenus and Pseudodiamesa from the fauna of Lake CF8 on Baffin Island over the past 75 years (Thomas et  al. 2008, Axford et al. 2009). Even assuming that species are able to translocate to appropriate habitats to escape extirpation, growing evidence suggests that areas previously regarded as potential cold refuges, such as the Hudson Bay Lowlands, are

now beginning to warm (Ruehland et al. 2013). Increasing shrub cover over a wider area is lead­ ing to the loss of open habitat and is supporting the establishment of incoming insect species through the creation of new and different habi­ tat (Klein et al. 2008, Rich et al. 2013). Changes to climate have implications for insect harassment of birds and mammals through­out the Arctic. The activity of mosqui­ toes (Culicidae), black flies (Simuliidae), and warble flies (Oestridae) is highly temperature‐ dependent. Increased adult mortality and reduced reproduction in Brunnich’s guillemot at Cape Pembroke in northern Hudson Bay, for example, occurs in warm years when mosqui­ toes are particularly abundant (Gaston and Elliot 2013). Models of biting fly intensity, using historical weather data for the summer range of the Bathurst caribou herd, suggest that mos­ quito attack has decreased between 1957 and 2008 but that of black flies and warble flies has increased (Witter et al. 2012). Arctic biting flies, particularly mosquitos, are known vectors of encephalitis viruses, such as Jamestown Canyon virus, snowshoe hare virus, and Northway virus, which infect a range of Arctic mammals includ­ ing Dall sheep, bison, Arctic fox, caribou, and snowshoe hare (Bradley et al. 2005). The former two viruses can also cause encephalitis in  humans. Changing climate, coupled with the spread of domestic livestock into the Arctic and range expansion in the exiting fauna, might increase the incidence of such insect‐borne diseases. This brief review shows how Arctic insects are capable of responding to a changing climate in various different ways, at both population and community levels. The inherent diversity of  their adaptations allows for a variety of responses, and we should not assume that all Arctic species will respond similarly. Some will be subject to the negative effects of increasing climatic stress, whereas others will respond positively to a warming climate. What is certain is that the Arctic insect community in a hun­ dred years time is likely to look different from that of today.

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2  Insect Biodiversity in the Arctic

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3 Insect Biodiversity in Indochina: A Window into the Riches of the Oriental Region Seunghwan Lee1 and Ram Keshari Duwal2 1 2

School of Agricultural Biotechnology, Seoul National University, Seoul, Korea Entomological Laboratory, Faculty of Agriculture, Kyushu University, Fukuoka, Japan

The Mekong River is a trans‐boundary river of Southeast Asia (Fig. 3.1). It is the seventh long­ est river in Asia and the twelfth longest in the world, estimated at 4350 km (Liu et  al. 2009). The main source of the river is melting snow on the Tibetan plateau to the north. The river flows southwest through Yunnan Province (China) and Indochina (Myanmar, Laos, Thailand, Cam­ bodia, and Vietnam), and finally meets the China Sea. The region of Indochina comprising these countries is hereafter referred to as the Lower Mekong Subregion. A small tributary near the junction of the three countries of Myanmar, Laos, and Thailand separates the northern upper Mekong River and southern lower Mekong River (Fig. 3.1). The rich biodiversity of the territory is poorly explored, although information on some insect orders of the Lower Mekong Subregion is avail­ able in a few data sets and catalogs for Thailand (Charernsom and Suasa‐ard 2000), the Oriental Region (Delfinado and Hardy 1975, 1977), and the world (Steinmann 1989, Morse et al. 2011, Blackman and Eastop 2012, Anonymous 2014). These catalogs barely cover the diversity of the Indochina peninsula and provide only fragmen­ tary information on certain groups. Several web‐based data sets are also available (Myers Enterprises 2009−2014, Ascher and Pickering 2016). Summary biodiversity information in

Table 3.1 and Table 3.2 supplements the existing catalogs and data sets by providing additional data derived from searches of more than 1300 literature references. During the literature search, we confirmed that knowledge of insect biodiversity for these countries is not well docu­ mented, except for a few groups. Therefore, using the work of Charernsom and Suasa‐ard (2000), we have prepared a checklist, by coun­ try, of insects in the Lower Mekong Subregion, except for the Coleoptera, for which catalogs and comprehensive revisions are available. Many taxa remain undocumented, although our data suggest that the insect fauna of Thailand and Vietnam is somewhat better known than that of Cambodia, Laos, and Myanmar. The purpose of this chapter is to compile and synthesize the available knowledge of insect biodiversity of the Lower Mekong Subregion as a window into the biodiversity of the Oriental Region and a basis for further research. It is difficult to find reliable data on biodiversity studies from the Lower Mekong Subregion because of unstable politics, weak economies, and civil wars that took place in Cambodia, Laos, and Vietnam; these conditions have con­ tributed to a lack of local expertise in biodiver­ sity studies. However, international researchers have contributed to the understanding of a few groups. Most existing specimen collections are

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

60

Insect Biodiversity: Science and Society 95°

90°

105°

100°

110°

30°

30°

er M Upp

China

ng eko

India 25°

25°

Myanmar

Vietnam

20°

20°

os

on

ek

La

rM we Lo g

Thailand

South China Sea

15°

15° Cambodia

Andaman Sea 10°

10°

Gulf of Thailand

90°

95°

100°

105°

10°

Figure 3.1  Map of Indochina showing the location of the Mekong River. Dashed line and arrows indicate the separation point of the upper and lower Mekong. Author originated. Base map exported from R package maps (Original S code by Richard A. Becker, Allan R. Wilks. R version by Ray Brownrigg. Enhancements by Thomas P Minka and Alex Deckmyn. (2016). maps: R package version 3.1.0.)

scattered across the foreign institutions where collection teams were based. However, several institutions in Thailand and Vietnam now undertake the preservation of museum speci­ mens. In general, the citizens of these coun­ tries are not aware of economically and environmentally important insects that directly affect society. Moreover, rapid deforestation and severe destruction of habitat in tropical Asia is resulting in the disappearance of biodi­ versity, including insect species and unique populations.

Several international organizations are actively working on the conservation of biodiversity in  this region, such as BirdLife, the CEPF (Critical Ecosystem Partnership Fund, a joint initiative of the European Union, Conservation International, the Global Environmental Facility, the government of Japan, the MacArthur Foundation, and the World Bank), the UNDP (United Nations Development Programme, which promotes biodiversity conservation as part of its remit to support under‐developed coun­ tries), the WCS (Wildlife Conservation Society),

62

Insect Biodiversity: Science and Society

and the WWF (World Wide Fund for Nature). However, these programs focus mainly on endangered birds, mammals, and reptiles, which are easily seen and counted. We summa­ rize the first initiative by the Korean govern­ ment toward the sustainable development and management of biodiversity in countries in the  Lower Mekong Subregion, including the insect fauna.

3.1 ­Physical Geography and Climate The Lower Mekong Subregion extends geo­ graphically from 8° to 29°N and from 92° to 110°E. It is bordered by southwestern China and eastern India, and on the south and east, it is flanked by the South China Sea (Fig. 3.1). It occupies an area of nearly 1.9 million km2, which is 4.35% of the total land mass of Asia (McCune 1947, Tordoff et al. 2012). The land­ scape is dominated by several mountain peaks  below 3048 m. However, the ice‐capped Hkakabo Razi and Gamlang Razi mountain ranges of Myanmar, bordering China and India, rise to 5881 m and 5870 m, respectively. This geographical variation is associated with a range of tropical, subtropical, temperate, and alpine ecoregions. The central and southeastern ecore­ gions consist of plateaus and low river basins in Cambodia, Laos, Thailand, and Vietnam, which include 22,230 km2 of conserved forest areas, and are covered by dry evergreen forest that often occurs with deciduous dipterocarp or mixed deciduous forest (WWF 2014a). To the north, about 15,948 km2 are preserved and occupied by highlands and valleys of Laos, Myanmar, and Vietnam with subtropical forest (WWF 2014b). The lower elevations, up to 600−800 m, are subtropical, representing a transition form of wet evergreen lowland forest and montane forest, typically composed of mixed vegetation of southern and northern ecoregions. At elevations of 800−2000 m, mon­ tane evergreen forest dominates. Elevations

above 2000 m consist of a moderate diversity of broadleaf trees and mixed vegetation similar to that in the Himalayas, followed by subalpine vegetation on the highest peaks. The diversity of flora and fauna in countries along the Lower Mekong Subregion is influ­ enced by climate and other geographic con­ ditions. The area generally exhibits tropical monsoons with rainy, hot, humid summers and mild, dry winters that are affected by wind from the Pacific Ocean. The southern part of the sub­ region experiences high summer temperatures of about 38 °C, with high humidity and pre­ cipitation, including thunderstorms (McCune 1947), whereas winters are drier and warm, with an average temperature of about 21 °C. The northern part has considerable seasonal tem­ perature variation, with averages of 28 °C in the summer and 16 °C in the winter (McCune 1947). Freezing temperatures are rare in the region, except for the extreme northern parts of Viet­ nam and Myanmar where the elevations are above 3000 m.

3.2 ­Features of Insect Biodiversity in the Lower Mekong Subregion The published data on the biodiversity of the Indochina peninsula show it to be a species‐rich area with large numbers of endemic plant and animal species, but the status of insects is rarely mentioned (Tordoff et  al. 2012). According to  available references, approximately 19,697 insect species have been reported from coun­ tries in the Lower Mekong Subregion (Table 3.1, Table 3.2) – 1.96% of the total number of insects in the world. This number is small compared with the known Palearctic insect fauna, from which we can infer that the Lower Mekong Subregion is not well explored. In compiling Table 3.1 and Table 3.2, we made an effort to clarify synonymies of species; however, some errors might appear in the taxonomy of some groups owing to a lack of published information

3  Insect Biodiversity in Indochina

Table 3.2  Number of insect species and genera, excluding Coleoptera, in the Lower Mekong Subregion. Number of species

Number of genera

Blaberidae

20

10

Blattellidae

23

14

Blattidae

20

11

Ceratopogonidae

Cryptocercidae

1

1

Ectobiidae

1

1

Panesthiidae

2

1

Polyphagidae

1

1

Raphidiomimidae

1

1

14

9

Apachyidae

1

1

Carcinophoridae

1

1

Chelisochidae

13

9

Diplatyidae

18

5

Forficulidae

47

16

Labiduridae

11

6

Labiidae

20

8

Pygidicranidae

23

7

Spongiphoridae

4

2

Family

Blattodea*

Dermaptera Anisolabididae

3

3

Agromyzidae

77

12

Anisopodidae

1

1

Anthomyiidae

10

7

Asilidae

81

35

Asteiidae

1

1

Athericidae

2

1

Aulacigastridae

2

2

Bibionidae

5

3

Blephariceridae

1

1

35

23

1

1

66

22

5

4

Bombyliidae Braulidae Calliphoridae Canaceidae

Number of genera

Cecidomyiidae

12

12

Celyphidae

24

3

125

27

Chamaemyiidae

2

2

Chaoboridae

2

1

Chironomidae

13

8

Chloropidae

26

20

Clusiidae

18

7

Conopidae

13

3

Corethrellidae

1

1

Ctenostylidae

2

2

Culicidae

595

46

Curtonotidae

5

2

Cypselosomatidae

6

3

Diadocidiidae

2

2

Diastatidae

1

1

Diopsidae

8

3

Dixidae

Diptera Acroceridae

Number of species

Family

 

3

2

Dolichopodidae

98

34

Drosophilidae

96

15

Empididae

59

6

Ephydridae

36

25

Fanniidae

2

1

Heleomyzidae

1

1

Heteromyzidae

1

1

Hippoboscidae

63

16

Hybotidae

92

10

Keroplatidae

8

7

Lauxaniidae

21

7

Limoniidae

52

21

Liphistiidae

3

1

Lonchopteridae

9

1

Megamerinidae

2

1

Micropezidae

17

4

Milichiidae

12

5 (Continued)

63

64

Insect Biodiversity: Science and Society

Table 3.2  (Continued) Number of species

Number of genera

222

38

50

4

Tanypezidae

Mydidae

2

1

Tephritidae

Mythicomyiidae

1

1

Thaumaleidae

Nemestrinidae

5

1

Therevidae

Neriidae

8

6

Tipulidae

Neurochaetidae

1

1

Ulidiidae

3

2

Nothybidae

2

1

Vermileonidae

1

1

Odiniidae

2

1

Xylomyidae

14

2

Oestridae

2

2

Xylophagidae

6

1

1

1

Family

Muscidae Mycetophilidae

Pediciidae

Family

Number of species

Number of genera

172

92

15

1

327

82

1

1

2

1

27

9

Tachinidae

1

1

13

4

103

34

Embonychidae

1

1

1

1

Oligotomidae

19

5

Pipunculidae

74

13

Teratembiidae

1

1

Platypezidae

3

1

Ephemeroptera

Platystomatidae

38

18

Baetidae

4

3

Psilidae

17

4

Behningiidae

2

2

Psychodidae

45

8

Caenidae

1

1

2

2

Ephemerellidae

22

14

Periscelididae Phoridae Piophilidae

Ptychopteridae Pyrgotidae

Embiidina Embiidae

7

5

Ephemeridae

18

4

Rhagionidae

25

4

Heptageniidae

39

15

Rhiniidae

43

8

Isonychiidae

1

1

Sarcophagidae

50

10

Leptophlebiidae

12

6

Scathophagidae

1

1

Neoephemeridae

6

3

Scatopsidae

2

1

Palingeniidae

1

1

Scenopinidae

5

4

Polymitarcyidae

3

2

Sciaridae

22

8

Potamanthidae

10

4

Sciomyzidae

10

4

Prosopistomatidae

2

1

Sepsidae

58

9

Teloganodidae

1

1

Simuliidae

33

1

Tricorythidae

3

1

Sphaeroceridae

71

33

Vietnamellidae

2

2

Stratiomyidae

Acanaloniidae

1

1

Achilidae

1

1

Aphrophoridae

1

1

84

31

Streblidae

8

5

Syrphidae

103

44

Tabanidae

238

7

Hemiptera (Auchenorrhyncha)

 

 

3  Insect Biodiversity in Indochina

Family

Number of species

Number of genera

Number of species

Number of genera

Hebridae

17

5

Helotrephidae

46

9

Family

Caliscelidae

2

1

Cercopidae

13

10

Cicadellidae

247

116

Cicadidae

250

57

Hydrometridae

Cixiidae

1

1

Hypsipterygidae

Coelidiidae

1

1

Largidae

Delphacidae

Henicocephalidae

1 1

1

1

15

4

28

19

2

2

Derbidae

2

2

Lygaeidae

50

25

Dictyopharidae

6

3

Malcidae

3

2

Flatidae

5

5

Mesoveliidae

2

1

Fulgoridae

Leptopodidae

1 15

15

9

Micronectidae

12

2

Icaniidae

1

1

Miridae

141

79

Issidae

6

4

Nabidae

4

3

Kinnaridae

1

1

Naucoridae

6

3

Ledridae

2

2

Nepidae

23

5

Lophopidae

2

2

Nerthridae

1

1

Machaerotidae

1

1

Notonectidae

14

4

Meenoplidae

1

1

Ochteridae

2

1

Membracidae

9

6

Pentatomidae

69

46

Ricaniidae

6

2

Piesmatidae

2

2

Tropiduchidae

5

1

Plataspidae

8

4

Pleidae

2

2

Hemiptera (Heteroptera) Alydidae

13

3

Plokiophilidae

1

1

Anthocoridae

17

9

Polyctenidae

2

1

Aphelocheiridae

11

2

Pyrrhocoridae

8

4

Aradidae

59

32

Reduviidae

152

73

Belostomatidae

5

4

Rhopalidae

1

1

Berytidae

3

2

Rhyparochromidae

5

3

Cimicidae

1

1

Saldidae

4

4

Colobathristidae

1

1

Schizopteridae

1

1

Coreidae

20

15

Scutelleridae

8

3

Corixidae

3

2

Tessaratomidae

3

2

Cydnidae

68

26

39

29

Dinidoridae

25

6

Urostylididae

2

2

2

1

Veliidae

29

15

85

25

5

1

Gelastocoridae Gerridae

Tingidae

Velocipedidae  

(Continued)

65

66

Insect Biodiversity: Science and Society

Table 3.2  (Continued)

Family

Number of species

Number of genera

Hemiptera (Sternorrhyncha) Aleyrodidae Aphalaridae

Family

Embolemidae 79

25

Number of species

Number of genera

3

2

Encyrtidae

73

46

1

1

Eucharitidae

15

9

111

57

Eulophidae

79

39

Calophyidae

1

1

Eupelmidae

12

5

Carsidaridae

2

2

Eurytomidae

8

8

Coccidae

25

14

Evaniidae

2

2

Diaspididae

36

17

Figitidae

10

2

Homotomidae

1

1

Formicidae

245

63

Kerriidae

3

2

Gasteruptiidae

3

1

Liviidae

8

3

Halictidae

19

9

Margarodidae

7

4

Ibaliidae

1

1

Meenopliidae

1

1

Ichneumonidae

152

102

Phacopteronidae

2

2

Leucospidae

2

1

Pseudococcidae

Aphididae

31

14

Megachilidae

26

10

Psyllidae

8

8

Megalyridae

1

1

Triozidae

7

3

Meliponidae

1

1

Melittidae

2

2

Hymenoptera (Apocrita) Agaonidae

3

3

Mutillidae

28

19

Alloxystidae

1

1

Mymaridae

5

5

Ampulicidae

2

2

Platygastridae

58

15

Aphelinidae

24

17

Pompilidae

11

8

131

23

Proctotrupidae

18

3

Bethylidae

72

26

Braconidae

212

65

Apidae Aulacidae

Ceraphronidae

2

2

43

31

Rhopalosomatidae

1

1

Roproniidae

1

1

Pteromalidae

4

3

Rotoitidae

1

1

Chalcididae

37

14

Sapygidae

1

1

Chrysididae

7

2

Scelionidae

35

29

Cleptidae

1

1

Scolebythidae

1

1

48

14

1

1

Colletidae

2

2

Scoliidae

13

7

Signiphoridae

Cynipidae

8

8

Sphecidae

39

20

Diapriidae

4

4

Tiphiidae

3

3

Dryinidae

122

18

Torymidae

10

10

Crabronidae

 

 

3  Insect Biodiversity in Indochina

Family

Trichogrammatidae Trigonalidae Vespidae

Number of species

Number of genera

16

7

Cossidae

1

1

Crambidae

143

31

Hymenoptera (Symphyta)

Family

Number of species

Number of genera

31

16

167

75

Ctenuchidae

4

3

Cyclotornidae

1

1

Argidae

2

2

Drepanidae

18

12

Diprionidae

8

6

Elachistidae

1

1

Orussidae

4

2

Endromidae

1

1

Pamphiliidae

2

2

Epermeniidae

2

2

Siricidae Stephanidae Tenthredinidae Xiphydriidae

1

1

Epicopeiidae

2

2

12

3

Epipyropidae

1

1

162

53

13

9

1

1

5

1

Eupterotidae

7

3

1

1

Gelechiidae

56

27

Geometridae

Erebidae Ethmiidae

“Isoptera” Archotermopsidae Hesperiidae

398

94

315

66

Hodotermitidae

1

1

Glyphipterigidae

13

3

Kalotermitidae

15

7

Gracillariidae

25

15

Rhinotermitidae

19

5

Hepialidae

12

6

106

30

Himantopteridae

1

1

Hyblaeidae

1

1

Termitidae Lepidoptera Adelidae

7

2

Immidae

5

2

Agonoxenidae

1

1

Lasiocampidae

25

13

Alucitidae

2

1

Lecithoceridae

135

28

Amphiteridae

2

1

Limacodidae

80

33

Arctiidae

95

45

551

123

Attevidae

2

1

Lymantriidae

54

21

Bedelliidae

1

1

Micropterigidae

1

1

Blastobasidae

1

1

Nepticulidae

3

1

Bombycidae

17

9

Noctuidae

345

185

Brachodidae

7

6

Nolidae

10

6

Brahmaeidae

1

1

Notodontidae

67

37

Callidulidae

3

2

Nymphalidae

715

111

Carposinidae

3

2

Oecophoridae

101

27

Choreutidae

27

4

Opostegidae

2

1

1

1

Palaeosetidae

22

9

Papilionidae

Coleophoridae Cosmopterigidae

Lycaenidae

 

1

1

184

16 (Continued)

67

68

Insect Biodiversity: Science and Society

Table 3.2  (Continued)

Family

Pieridae Psychidae Pterophoridae

Number of species

Number of genera

156

24

17

14

Number of species

Number of genera

23

8

Lepismatidae

3

3

Family

Megaloptera Corydalidae

14

11

126

95

28

7

Machilidae

5

4

3

2

Meinertellidae

3

2

Saturniidae

46

16

Nicoletiidae

7

5

Scythrididae

1

1

Protrinemuridae

2

2

13

9

Pyralidae Riodinidae Roeslerstammiidae

Sesiidae

Microcoryphia and Zygentoma

44

21

Sphingidae

200

66

Ascalaphidae

Thyrididae

31

15

Berothidae

2

2

Tineidae

32

20

Chrysopidae

16

9

300

130

Coniopterygidae

18

8

4

3

Dilaridae

3

2

Tortricidae Uraniidae Urodidae

Neuroptera

1

1

Hemerobiidae

10

6

Yponomeutidae

26

14

Mantispidae

10

5

Zygaenidae

26

16

Myrmeleontidae

Mantodea

45

26

Osmylidae

2

2

Amorphoscelidae

1

1

Psychopsidae

3

2

Empusidae

2

2

Rapismatidae

3

1

Gryllomantidae

1

1

Sisyridae

2

1

Hymenopodidae

17

9

Iridopterygidae

5

3

Aeshnidae

28

13

Jantarimantidae

1

1

Agrionidae

39

10

Amphipterygidae

Liturgusidae Mantidae Metallyticidae

Odonata

7

3

2

2

46

23

Calopterygidae

22

9

2

1

Chlorocyphidae

11

5

3

3

Thespidae

2

2

Chlorogomphidae

Toxoderidae

6

2

Chlorolestidae

1

1

Coenagrionidae

17

8

Bittacidae

4

2

Cordulegastridae

Meropeidae

1

1

Panorpidae

38

3

1

1

Mecoptera

Pseudopolycentropodidae

Corduliidae Disparoneuridae

 

5

2

25

6

1

1

Euphaeidae

19

7

Gomphidae

55

23  

3  Insect Biodiversity in Indochina

Number of species

Number of genera

12

3

1

1

16

12

6

4

Platycnemididae

24

4

Platystictidae

12

2

Protoneuridae

7

1

139

73

Family

Lestidae Libellaginidae Libellulidae Megapodagrionidae

Anostostomatidae

5

2

Chorotypidae

15

4

Dericorythidae

1

1

Erythraeidae

3

1

Eumastacidae

1

1

Gryllacrididae

62

20

206

72

Gryllotalpidae

2

1

Metrodoridae

2

1

Mogoplistidae

24

7

Myrmecophilidae

1

1

Pyrgomorphidae

17

7

Rhaphidophoridae

68

16

Tetrigidae

47

28

Tettigoniidae

251

79

Tridactylidae

2

1

Gryllidae

11

9

Phasmatidae

10

9

Phylliidae

1

1

Pseudophasmatidae

1

1

22

8

2

Trichodectidae

1

1

8

1

Nemouridae

26

6

Peltoperlidae

13

2

Perlidae

99

14

1

1

Amphientomidae

1

1

Caeciliusidae

1

1

Calopsocidae

1

1

Epipsocidae

2

2

Lachesillidae

2

1

Pseudocaeciliidae

1

1

Psocidae

3

3

Psyllipsocidae

1

1

Stenopsocidae

1

1

Inocelliidae

1

1

Mesoraphidiidae

1

1

Ancistropsyllidae

1

1

Ceratophyllidae

2

1

Hystrichopsyllidae

4

2

Ischnopsyllidae

8

6

Leptopsyllidae

1

1

Pulicidae

9

6

Pygiopsyllidae

3

2

Stivaliidae

6

3

Callipharixenidae

1

1

Corioxenidae

1

1

Elenchidae

2

1

Halictophagidae

5

1

Styloperlidae

Raphidioptera

Siphonaptera

Strepsiptera

Enderleinellidae

1

1

Hoplopleuridae

4

1

19

8

3

1

Pedicinidae

50

Polyplacidae

Psocoptera

Phthiraptera

Menoponidae

Philopteridae

Leuctridae

Phasmatodea Diapheromeridae

Number of genera

Plecoptera

Orthoptera Acrididae

Number of species

Family

 

(Continued)

69

70

Insect Biodiversity: Science and Society

Table 3.2  (Continued)

Family

Number of species

Number of genera

Family

Number of species

Number of genera

Myrmecolacidae

5

3

Leptoceridae

250

12

Stylopidae

3

2

Limnephilidae

6

2 1

4

4

Limnocentropodidae

8

Aeolothripidae Merothripidae

1

1

Molannidae

8

2

Phlaeothripidae

44

28

Odontoceridae

32

5

Thripidae

82

41

Philopotamidae

190

5

Phryganeidae

2

2

Phryganopsychidae

3

1

Polycentropodidae

89

9

Psychomyiidae

91

5

Ptilocolepidae

1

1

71

4

2

1

16

1

3

1

22

5

Thysanoptera

Trichoptera Apataniidae

14

3

Brachycentridae

8

1

Calamoceratidae

30

4

Dipseudopsidae

72

4

Ecnomidae

62

1

Rhyacophilidae

Glossosomatidae

51

5

Sericostomatidae

Goeridae

32

3

Stenopsychidae

Helicopsychidae

28

1

Uenoidae

Hydrobiosidae

7

1

Hydropsychidae

202

14

Hydroptilidae

154

19

30

3

Lepidostomatidae

Xiphocentronidae Zoraptera Zorotypidae Total

506

4 16,044

2 5,183

* Termites (“Isoptera”) are given separately to provide additional detail.

and available expertise. The data show that bee­ tles and butterflies make up half of the total number of species known from this region. Below, we briefly discuss the biodiversity of the insect orders, alphabetically, in the Lower Mekong Subregion (Fig. 3.2).

species having been recorded. The fauna is under‐sampled, and no species are currently recognized as shared among countries, although we assume this situation will change as knowl­ edge of the fauna is developed. 3.2.2 Coleoptera

3.2.1 Blattodea

Eight families and 69 species of Blattodea, excluding the termites, are reported from coun­ tries in the Lower Mekong Subregion – 1.5% of the total species of Blattodea in the world. Most of the species are described from Myanmar, Thailand, and Vietnam, whereas Cambodia and Laos remain understudied, with only a few

Beetles are one of the most comprehensively sampled groups in the region. Seventy‐three families have been reported from this region with 3653 species – 1.4% of all beetle species in the world. The greatest numbers of species are recorded from Laos and Thailand  –  1642 and 1706 species, respectively. A detailed study of the Cerambycidae in Laos (1155 species) by

3  Insect Biodiversity in Indochina

(a)

(b)

(c)

(d)

(e)

(f)

Figure 3.2  Diversity of insect species from the Lower Mekong Subregion. (a) Papilio sp. (Lepidoptera). (b) Graphium antiphates (Lepidoptera). (c) Apis dorsata (Hymenoptera). (d) Xylocopa sp. (Hymenoptera). (e) Cucujus sp. (Coleoptera). (f ) Aphis nerii (Hemiptera). Images by authors.

Gressitt et  al. (1970) shows that this family makes up more than one‐quarter of the total Coleoptera from the region. 3.2.3 Dermaptera

Ten families, represented by 152 species (7.7% of the world dermapteran fauna), are found in

the region. The Forficulidae are particularly diverse and widely distributed. The world cata­ log of the Dermaptera (Steinmann 1987) is the main reference, together with recently pub­ lished data. The Dermaptera have been studied in Myanmar, Thailand, and Vietnam to some extent, whereas only two species are known from Cambodia.

71

72

Insect Biodiversity: Science and Society

3.2.4 Diptera

3.2.8 Hymenoptera

About 3640 species of Diptera are reported from the Lower Mekong Subregion  –  2.4% of the total dipteran species in the world. Most reported species are from Myanmar, Thailand, and Vietnam, whereas Cambodia and Laos remain understudied.

The Hymenoptera are an economically impor­ tant group because they are effective pollinators and natural enemies of pest insects. Sixty‐three families are recorded from the countries of the region, with 1955 described species (1.3% of the world’s species in the order). The best‐studied families are the Apidae, Braconidae, Dryinidae, Formicidae, Ichneumonidae, Tenthredinidae, and Vespidae. Two‐thirds of the region’s total species of Hymenoptera are bees. The commer­ cially important honeybees and stingless bees have been particularly well studied, especially in Thailand. The most important source of infor­ mation on the Hymenoptera is that by Ascher and Pickering (2016).

3.2.5 Embiodea

The webspinners are a small group of insects that favor tropical and subtropical climatic conditions. Four subfamilies comprising 22 spe­ cies (4.8% of the world’s species in the order) are found in countries of the Lower Mekong Subregion. Most species are reported from Thailand; none are known from Cambodia.

3.2.9 “Isoptera” 3.2.6 Ephemeroptera

From the countries along the Lower Mekong Subregion, 16 families comprising 127 species (4.2% of the world’s mayfly fauna) have been recorded. Few species are recorded from more than one country, but we expect that most spe­ cies are widespread in the region. No species have been recorded in Cambodia. 3.2.7 Hemiptera

The important references for Heteroptera of the region are those by Schuh (1995), Polhemus and Polhemus (2003), Vitheepradit (2008), and Yasunaga and Duwal (2015). For Sternorrhyncha, they are by Blackman and Eastop (2012) and Martin and Mound (2007), and for Auchen­ orrhyncha, Thai and Yang (2009) and Boulard (2012). Ninety‐one families occur in the coun­ tries of the region, comprising 1940 species (1.9% of the world’s hemipteran species). More than half of these species belong to the suborder Heteroptera. The fauna of Thailand is relatively well studied, whereas that of Cambodia and Myanmar is poorly known.

Five families of termites are recorded from countries of the region, represented by 142 spe­ cies (5.0% of the world’s species of termites). The fauna of Thailand and Vietnam is best known, with only a few species reported from the other countries. 3.2.10 Lepidoptera

Seventy families of Lepidoptera are known from countries in the region, with 4600 species recorded (2.9% of the total butterflies and moths in the world). The largest number of species has been recorded from Thailand and Vietnam (3712 and 1410 species, respectively). The families Arctiidae, Geometridae, Hesperiidae, Leci­ tho­ ceridae, Lycaenidae, Noctuidae, Nymphalidae, Oecophoridae, Papilionidae, Pieridae, Pyralidae, Sphingidae, and Tortricidae are particularly well represented in the region. The “Checklist of Thailand” (Charernsom and Suasa‐ard 2000) is an important source, listing 3547 species, although some difficulties were encountered using this list due to changes in classification and nomencla­ ture. Many investigators have contributed to the

3  Insect Biodiversity in Indochina

understanding of the Lepidoptera of the region, including Dean (1978), Leps and Spitzer (1990), Pitkin et  al. (2007), Spitzer and Jaros (2008), Lien (2010, 2013), Monastyrskii (2010), Polaszek (2010), Monastyrskii and Hollo­way (2013), and Park (2014). 3.2.11 Mantodea

Eleven of 15 families are distributed in countries of the region, and include 90 species, or 3.8% of the world’s species of mantids. As with other orders, the Mantodea are least known for Cam­ bo­dia and Laos. 3.2.12 Mecoptera

Only five families are known in the region, with 44 species (6.5% of world’s mecopteran species). The family Panorpidae is relatively well studied. Most recorded species are from Myanmar, Thailand, and Vietnam. 3.2.13 Megaloptera

neuropteran species). Only a few species are known from Cambodia and Laos (four and two species, respectively). 3.2.16  Notoptera (Grylloblattodea and Mantophasmatodea)

No species are known from the region. Possibly, they could be found above 5000 m in the north­ ernmost mountains of Myanmar. 3.2.17 Odonata

Twenty families are distributed in countries of the region, represented by 306 species (5.4% of the world’s species of odonates). The two fami­ lies, the Agrionidae and Gomphidae, are the most diverse in the region. Most species are found in Thailand, but contrary to the pattern in other groups, the fauna of Cambodia is rela­ tively well known. 3.2.18 Orthoptera

A single family, the Corydalidae, has been recorded from the region, comprising 23 spe­ cies (6.8% of the world’s megalopteran species). All species are recorded from Thailand except Neochauliodes tonkinensis (Weele 1907), which is found in Vietnam.

Seventeen families and 846 described species (3.6% of the world’s orthopteran species) are known from the region. The Acrididae, Gryl­ lidae, Tettigoniidae, and Rhaphidophoridae are the most abundant families. Although the fau­ nas of Thailand and Vietnam are well studied, the two countries share only a few species.

3.2.14  Microcoryphia and Zygentoma

3.2.19 Phasmatodea

Five families are found in the region, with 20 species (3.8% of the world’s species in these orders). The recorded species are all from Myanmar, Thailand, and Vietnam. No species are known to be shared among countries.

Four families are recorded from the region, with 23 species (0.8% of the world’s species in the order), mostly from Vietnam.

3.2.15 Neuroptera

The data reveal 12 families from the region, with 127 described species (2.2% of the world’s

3.2.20 Phthiraptera

Seven families and 86 species (1.7% of the world’s species of lice) are recorded from the region. The largest number of species, 62, is reported from Thailand.

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3.2.21 Plecoptera

3.2.26 Thysanoptera

The stoneflies are a widely distributed group in the Southern and Northern Hemispheres, and are considered important indicators of water quality. Within the region, five families are known, with 147 species (4.2% of the world’s species of stoneflies). The family Perlidae is the most diverse in Thailand and Vietnam; few spe­ cies are known from Laos and Myanmar. The other four families are known only from Thailand and Vietnam. No species have been reported from Cambodia.

Four families are reported from the region, including 131 species (2.3% of the world’s spe­ cies of thrips), most of them from Thailand.

3.2.22 Psocoptera

Nine families and 13 described species (0.2% of the world’s psocopteran species) are known from the region, all from Thailand and Vietnam. 3.2.23 Raphidioptera

The species of this group are typically found in temperate climates and, therefore, would be expected only in the northern parts of the Lower Mekong Subregion. Two species, one in the Inocelliidae and the other in the Mesoraphidiidae, are known from Myanmar. 3.2.24 Siphonaptera

Eight families are known from countries in the region, comprising 34 species (1.7% of the world’s species of fleas), mainly from Cambodia, Thailand, and Vietnam. A single species, Lentis­ tivalius insolli (Traub), is known from Laos and is common in all countries of the region. 3.2.25 Strepsiptera

Six families are known from countries in the region, and include 17 species (2.8% of the world’s strepsipteran species). Most species are known only from Thailand. None are reported from Cambodia or Myanmar.

3.2.27 Trichoptera

Twenty‐eight families are known from the region, with 1484 recorded species (11.5% of the world’s species of caddisflies). The database of Morse et al. (2011) is the most important refer­ ence for Oriental caddisflies. Trichoptera are well studied in Thailand and Vietnam, whereas Cambodia, Laos, and Myanmar remain poorly investigated. 3.2.28 Zoraptera

No extant species of this order are known from the region, but four fossil species are described from Myanmar (Engel and Grimaldi 2002).

3.3 ­Insect Biodiversity and Society in Indochina 3.3.1  Entomophagy in the Lower Mekong Subregion

Entomophagy, or the consumption of insects by  humans, has occurred for various reasons throughout history (Defoliart 2002). Although the consumption of insects has declined with the development and adoption of Western civiliza­ tion and attitudes, some ethnic groups still regu­ larly use insects, without hesitation, as a source of food. In some parts of the world, consumption of insects has been commercialized. Thailand is among the countries where insect foods are pop­ ular locally and commercially and are exported to the rest of the world. The buying and selling of insects also takes place in Cambodia, Laos, Myanmar, and Vietnam (Fig.  3.3). Nearly 164 species are reported as being edible in countries

3  Insect Biodiversity in Indochina

(a)

(b)

(c)

(d)

(e)

(f)

Figure 3.3  Insect foods in local market of Laos. (a) Coleoptera. (b) bamboo caterpillar (Lepidoptera). (c−g) Orthoptera. (h) Hemiptera. Images by authors.

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(g)

(h)

Figure 3.3  (Continued)

of Indochina (Yhoung‐Aree and Viwatpanich 2005), especially those of beetles, crickets, giant water bugs, grasshoppers, mantids, silkworms, and termites (Defoliart 2002). Domestication of certain insects for commer­ cial value, such as honeybees, lac insects, and silkworms, is common and familiar to many people, but the commercial production of insects for consumption is rare. Edible insects are mostly collected in the wild or are semi‐cul­ tivated. During a field survey in Cambodia in 2010, we observed an interesting method of col­ lecting giant water bugs by a local farmer in the Kampong Seila District. A florescent light trap placed in a large bucket was set into the ground and filled with water. By the next morning, the farmer had captured hundreds of live giant water bugs, as well as crickets and various semi­ aquatic insects. In contrast with the larger insect market of Thailand, we observed only a few insect snack shops near temples and in tradi­ tional street markets of other countries in the Lower Mekong Subregion (Fig. 3.3). In ancient times, people consumed insects to satisfy their hunger, but their choice of particu­ lar insect species provided an important source of fiber, minerals, protein, and vitamins (FAO 2013). Although the current trend is to con­ sider insects only as pests, world leaders are looking  to insects as sustainable alternative

food sources. The return to the tradition of insect consumption should be encouraged and promoted by governments at all levels, and should include research and development programs for the commercialization of insect production. Although some universities, such as Khon Kaen  University (Thailand) and the National University of Laos (Faculty of Agri­ culture) in Indochina and a few others around the world, are involved in research on insects of  high nutritional value, these kinds of pro­ grams need to be initiated on a larger scale. As countries in Indochina are economically poor, encouraging the consumption of insects can be a low‐cost method of improving health and liv­ ing standards. 3.3.2  Research Initiatives

The Indochina peninsula has been identified as the core of the “Indo‐Burma Hotspot” (Tordoff et al. 2012), a region with one of the highest lev­ els of biodiversity in the world. Several inter­ national governmental and non‐governmental organizations (e.g., BirdLife, CEPF, UNDP, WCS, and WWF) working on conservation pro­ jects are involved in the exploration of biodiver­ sity in this region. The Korean government recently showed interest in biodiversity research in the Indo‐Burma Hotspot. The International

3  Insect Biodiversity in Indochina

Cooperation Unit on Biodiversity and Environ­ ment Conservation (ICUBEC), a subunit of the  National Institute of Biological Resources (NIBR), supported by the Korean Ministry of Environment, has initiated a project focused on the biodiversity of Cambodia, Laos, Myanmar, and Vietnam, in collaboration with several uni­ versities in Korea and institutions in the host countries (the Forest Ministry in Cambodia; the Department of Forest Resource Management in Laos; the Forest Department in Myanmar; and the Institute of Ecology and Biological Resources in Vietnam). The purpose of ICUBEC is to determine and assess the biodiversity knowledge of countries in the Lower Mekong Subregion, with an emphasis on conservation. To meet this goal, each research group is required to (i) conduct a series of field surveys to evaluate biodiversity, (ii) publish new and important findings, (iii) determine economically important plants and insects, (iv) provide field and laboratory train­ ing for people in the countries of the region, (v) encourage advanced education in biodiversity studies, and (vi) initiate public awareness pro­ grams to enhance forest‐habitat conservation. Since 2010, the Insect Biosystematics Labo­ ratory, Seoul National University, has been involved in ICUBEC, with responsibility for studying the Hemiptera and Hymenoptera from Cambodia, Laos, and Myanmar. During field work, we include government officers and local villagers in our team so that both groups can increase their understanding of survey areas and knowledge of the insect fauna. We have been involved with teaching and training local collaborators in collection methods, data man­ agement, and specimen curation. We also assist in the identification of pest species, suggest con­ trol measures, and provide expertise on eco­ nomically important insects. According to the annual reports of ICUBEC from 2007 to 2016, more than 1000 species of insect have been identified, although thousands of specimens remain unidentified. Many spe­ cies have been recorded for the first time in the region, and several species are new to science.

In one of the groups for which our own team is responsible, the Hemiptera, about 70% of the species that we have collected from Cambodia and Laos are new to those countries. Some data have already been published: Lim and Lee (2011, 2013), Park and Bae (2012), Jung et  al. (2013), Park et  al. (2013), Qi et  al. (2013), Oh et  al. (2015), Bayarsaikhan and Bae (2016), Bayarsaikhan et al. (2016), Lee and Lee (2016), S. Lee et  al. (2016), Y. Lee et  al. (2016), Park et  al. (2016), Win et  al. (2016), Yasunag et  al. (2016a, 2016b), and Duwal et  al. (2017). The majority of the specimens were deposited with the NIBR or cooperating universities, or else as indicated in each paper. A representative of each identified species has been returned to collaborating institutions in Cambodia, Laos, Myanmar, and Vietnam.

3.4 ­Conclusions Although 19,721 species of insects have been recorded from the Lower Mekong Subregion (Table 3.1), the insect fauna remains poorly studied. The current list, however, provides a framework for further biodiversity studies in the subregion as well as in the larger Oriental Region. Given the geographic extent of the sub­ region and its recognition as a biodiversity hot­ spot, the number of species known is probably a small proportion of the number of species actu­ ally present. In general, institutional support for insect studies has not been a high priority for coun­ tries in Indochina, although some government organizations and colleges in Thailand and Vietnam have undertaken research in some groups. Most of the current knowledge of the biodiversity of the area has been contributed by international biologists. The development of local expertise is necessary if a more complete knowledge of insect biodiversity in the area is to be achieved. The data we have assembled show that Thai­ land and Vietnam are comparatively advanced in faunistic studies of certain insect groups,

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whereas the insect faunas of Cambodia, Laos, and Myanmar are more poorly known. For example, orders such as the Mecoptera, Megaloptera, Microcoryphia, Raphidioptera, Plecoptera, Strepsiptera, Trichoptera, and Zygentoma, have not been reported from more than one of these countries.

Acknowledgments We thank the laboratory members and intern students of the Insect Biosystematics Laboratory, Seoul National University, for searching for refer­ ences (Yerim Lee, Hwaseop Song, Geonho Cho, Seunghyun Lee, and Jinyeong Choi) and for assisting in developing the species lists (Sungjik, Deowan Kim, and Sanghyeok Nam). We also thank the NIBR for providing the opportunity to participate in biodiversity surveys of the Lower Mekong Subregion.

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Park, K. T., S. Kim, M. Y. Kim and Y. S. Bae. 2016. Genus Lecithocera Herrich‐Schäffer in Vietnam (Lepidoptera, Lecithoceridae, Lecithocerinae), with descriptions of eight new species. Journal of Asia‐Pacific Entomology 19: 295–305. Pitkin, L. M., H. Han and S. James. 2007. Moths of the tribe Pseudoterpnini (Geometridae; Geometrini); a review of the genera. Zoological Journal of the Linnean Society 150: 343−412. Polaszek, A. 2010. Species diversity and host associations of Trichogramma in Eurasia. Progress in Biological Control 9: 237−266. Polhemus, D. A. and J. T. Polhemus. 2003. A review of Veliinae of Vietnam (Heteroptera: Veliidae) with description of a new Velia species. Journal of the New York Entomological Society 111: 29−40. Qi, M. J., G. M. László, G. Ronkay, Y. S. Bae and H. L. Han. 2013. Description of a new genus Purenola Qi, László, Ronkay, Bae & Han, gen. n. and a new species of the tribe Nolini from Cambodia (Lepidoptera: Nolidae, Nolinae). SHILAP Revista de Lepidopterología 41: 371−376. Schuh, R. T. 1995. Plant Bugs of the World (Insecta: Heteroptera: Miridae): Systematic Catalog, Distributions, Host List, and Bibliography. New York Entomological Society, New York, New York. 1329 pp. Spitzer, K. and J. Jaros. 2008. Annotated checklist of the butterflies (Papilionoidea) of the Nam Cat Tien Reserve (South Vietnam). Tropical Lepidoptera 18: 12−16. Steinmann, H. 1987. A new reclassification of the family Chelisochidae (Dermaptera). Annals Historico‐Naturales Musei Nationalis Hungarici 79: 113−118. Steinmann, H. 1989. World Catalogue of Dermaptera. Series Entomologica 43: 1−934. Thai, P. H. and J. T. Yang. 2009. A contribution to the Cicadellidae fauna of Vietnam (Hemiptera: Auchenorrhyncha), with one new species and twenty new records. Zootaxa 2249: 1−19. Tordoff, A. W., M. C. Baltzer, J. R. Fellowes, J. D. Pilgrim and P. F. Langhammer. 2012. Key

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4 Biodiversity of Arthropods on Islands Rosemary G. Gillespie and Kipling Will Department of Environmental Science, University of California Berkeley, Berkeley, California, USA

The intrigue of islands to naturalists was spawned in the Pacific, initially through the col­ lections of Joseph Banks, who served as natural­ ist, accompanying James Cook on the voyages of the Endeavour (1768–1771) (Whitehead 1969, Diment et  al. 1984). The subsequent work of Charles Darwin (1859) emphasized the signifi­ cance of islands in addressing questions of evolu­ tionary origin: isolated archipelagos can serve as  natural laboratories for studies in evolution and adaptive radiation (Howarth 1990, Thornton 1996) and as microcosms for research in ecologi­ cal processes. Thus, the Galápagos Islands served as a focus in the development of Charles Darwin’s theory of evolution by means of natural selection (Grant and Grant 2007). Likewise, the Spice Islands of Indonesia formed a basis for the devel­ opment of similar theories by Alfred Russel Wallace. More recently, the closed nature of insu­ lar systems, coupled with their relative simplicity, has allowed them to serve as microcosms for understanding fundamental processes in both ecology and evolution.

4.1 ­What is an Island? An island is generally defined as a piece of land, smaller than a continent, which is surrounded by water. As such, because water represents a barrier to humans, the term has been associated

with isolation, and many biological ideas about islands have developed in the context of this iso­ lation. However, isolation is scale and context dependent. A lake in the middle of a continent may be isolated for a fish, but not so much for a horse; an oceanic island may be isolated for a frog or an insect, but not for a whale; and a rock in the middle of a forest may be isolated for an ant, but not for a pig. Thus, the level of isolation is dictated by the impermeability and extent of the isolating matrix relative to the organism in question. But whatever the context, the well‐ defined nature of islands results in properties such as a microcosmal nature and a uniquely assembled biota. In essence, islands can be con­ sidered nature’s test tubes. Each island repre­ sents a trial in an experiment, and each new island is the repeat of one of these experiments. In considering this concept of an island, it becomes clear that almost anything can serve as an island, depending on the scale: water‐filled tree holes can serve as islands for many invertebrates; mountaintops can serve as so‐called “sky islands” for many arthropods (McCormack et  al. 2009); and an animal’s body is an island to the parasites it contains (Reperant 2010, Bossard 2014). Given this context, the purpose of this chapter is to cover a broad range of insular systems as they pertain to arthropods. In general, the composition of biological com­ munities on islands is dictated by (i) the history

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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of the island (i.e., how it was formed), (ii) the degree of isolation of the island relative to the dispersal abilities of an organism, (iii) the area of the island, and (iv) the age of the island, or the time over which its area has been habitable (Gillespie and Roderick 2002). Although organ­ isms differ in the way that they respond to these properties, the diversity of arthropods means that they show the greatest range of responses. 4.1.1  History of the Island

In terms of history, islands can be divided into  two broad categories: (i) fragment islands (Fig. 4.1c,f ) that were formed by separation from the source, and (ii) de novo islands that were cre­ ated without life. In the case of fragment islands that become separated from a larger entity, commonly from a (continental) land mass, the available niche space on that fragment has usually been filled to a large extent before isola­ tion. Accordingly, with increasing isolation, the number of species can only decrease over time. Indeed, because they are usually ecologically

saturated at the time of separation, they tend to  lose species through ecological time, a phe­ nomenon termed “relaxation” (Diamond 1972, Cayuela 2009). Over evolutionary time, the spe­ cies on these islands may change through a pro­ cess termed “relictualization,” with the formation of paleo‐endemics simply a result of extinction of close relative. However, the biological changes on such islands will depend on the interplay between isolation and time. If islands are not extremely isolated, continual immigration and extinction might be expected. But on islands that have been very isolated over extended time periods, as in the case of the ancient islands of Madagascar, New Caledonia, New Zealand, Sri Lanka, and others, paleo‐endemics may increase over time, as for pelican spiders in Madagascar (Wood et al. 2013). In contrast to fragments, life on de novo islands (most commonly islands of volcanic ori­ gin, Fig. 4.1a) is contingently built by colonists from outside sources. Such islands have served as the setting for the well‐known Equilibrium Theory of Island Biogeography, now applied to

Figure 4.1  Different kinds of insular systems. (a) Oceanic island. The island of Pico in the archipelago of the Azores. This archipelago is one of the best studied for arthropods, as evidenced through the work of Borges and colleagues (Borges 1992, Borges and Brown 1999, Cardoso et al. 2010, Triantis et al. 2010a, Triantis et al. 2010b, Cardoso et al. 2011, Gaspar et al. 2011, Meijer et al. 2011). (b) Cave. View towards entrance from within Algar do Carvão, an ancient lava tube on the Azorean island of Terceira, which harbors a number of endemics, including both spiders and insects (Reboleira et al. 2011). (c) Pleistocene fragment island. Agistri in the Sarconic Islands of Greece. Early on (23–12 million years ago), the Greek islands were all connected in a continuous land mass. Sea transgression (12–5 mya) formed a mid‐Aegean barrier, followed by fragmentation and widening of the Aegean, leading to the Pleistocene, which was characterized by eustatic sea‐level change (Triantis and Mylonas 2009). The long history of connection and isolation has shaped the diversity of arthropods known from the region today (Sfenthourakis and Legakis 2001). (d) Forest fragment. Shown is a “kipuka,” or island of forest surrounded by lava, on the island of Hawaii. Arthropods are often isolated in these fragments and show clear genetic differences among kipukas (Vandergast and Gillespie 2004, Vandergast et al. 2004). (e) Unique habitat islands. Mono Lake is an example of a unique habitat – a saline lake – that is isolated from similar such habitats. It harbors distinct assemblages of organisms, particularly notable being the brine shrimps and alkali flies (Herbst 1999). Other habitat types that hold unique assemblages of arthropods include sand dunes (Van Dam and Matzke 2016) and vernal or desert pools (Ward and Blaustein 1994). (f ) Recent fragment islands. Barro Colorado island, in Gatun Lake of Panama, was formed by flooding of the Chagres River in the creation of the Panama Canal (Leigh 2009). As a result, species numbers declined through relaxation of the supersaturated insular biota as it returns to equilibrium. A similar phenomenon has been documented for oaks, which act as islands for leaf‐mining insects (Opler 1974). (g) Sky islands. The American Madrean sky islands of southeastern Arizona and New Mexico have served to isolate many arthropods on the mountain summits. Particularly well known are the jumping spiders in the Habronattus pugillis complex (Masta 2000), scorpions (Hughes 2011), and beetles (Smith and Farrell 2005, Ober and Connolly 2015). All photographs by George K. Roderick, used with permission. (See color plate section for the color representation of this figure.)

4  Biodiversity of Arthropods on Islands

many and diverse insular settings. Given suffi­ cient isolation, these islands also provide the setting for the formation of endemic species (Gillespie and Roderick 2002). Moreover, when isolation is extreme, the few successful colonists are presented with abundant ecological oppor­ tunity; such conditions have paved the way for  adaptive radiation in multiple lineages. Besides Hawaii, other volcanic archipelagos have formed de novo and provided conditions for the formation of neo‐endemics, with exam­ ples from the Marquesas, Societies (Gillespie

et  al. 2008b), and Galápagos islands (weevils, Sequiera et  al. 2008) in the Pacific; the Canaries (Juan et al. 2000) and Azores (arthropods, Borges and Hortal 2009) in the Atlantic; and the Mascarenes (weevils, Kitson et al. 2013) in the Indian Ocean. Although the contrast between de novo and fragment islands might seem clear‐cut, geologi­ cal events subsequent to the formation of some ancient land masses, such as submersion or obduction, can reduce the size of the island or its habitable area to such an extent that it

(a)

(b)

(c)

(d)

(e)

(f)

(g)

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behaves like – and its floral and faunal composi­ tion are largely consistent with  –  a de novo island. For example, neo‐endemics can form on fragment islands, as shown for insects on the islands of Madagascar (Miraldo et al. 2011) and New Zealand (Buckley et al. 2015), where fluc­ tuations in area or resources seem to have inter­ mittently opened up ecological space, and on New Caledonia, where dispersal and endemic radiations on emergent land areas are major forces behind the current patterns of diversity (Grandcolas et  al. 2008, Murienne et  al. 2008, Cruaud et al. 2012). 4.1.2  Degree of Isolation

The traditional view of an island is a land mass surrounded by water. However, a huge range of habitats can be considered within this context, from intermittent tidal islands such as Le Mont‐Saint‐Michel to the remote islands of Polynesia. Islands that are less remote by dis­ tance, age of separation, and type of barrier are more likely to share species and near relatives with the proximal source area and be subject to gene flow from source‐area populations. The level of isolation is dictated by the dispersal abilities of the organism, whether using aerial transport, rafting, or vectors, although these frequently change subsequent to island colo­ nization. Although dispersal strategies associ­ ated with these mechanisms generally evolved for local movement to enable migration, to escape predation, or to locate mates and resources, the same traits can allow long‐­distance dispersal in some taxa (Gillespie et  al. 2012). Moreover, taxa with similar dispersal abilities frequently colonize remote archipelagoes mul­ tiple times from a mainland source (Gillespie et  al. 2008), as shown by fulgoroid planthop­ pers that have colonized different remote islands of Oceania independently (Asche 1997). Arthropods commonly colonize remote islands by aerial transport  –  mostly flying by insects and ballooning by spiders  –  and by ­rafting on debris or wood (Peck 2008), which is  most common among leaf miners, wood

­ orers, and other insects that inhabit plant b debris (Gressitt 1961) or attach their eggs to vegetation (Perkins 1907, Paulay and Meyer 2002). However, a number of arthropods appar­ ently have used vectors (migratory birds in par­ ticular) to reach remote islands (Green and Figuerola 2005). 4.1.3  Area of the Island

Islands range in size from tiny Polynesian motus, Florida Keys, and English eyots, to the large “island” of Greenland. For a given latitude, the relationship between area (A) and the number of species (S) tends to follow the power law relation, S ∞ Az. (MacArthur and Wilson 1967), with an additional effect of topography (Hortal et al. 2009). The premise of the theory is that species diversity on an island is a balance between immigration and extinction. The rate of immigration decreases with increasing dis­ tance from a mainland source, whereas the rate of extinction decreases with increasing island size. As the number of resident species on an island increases, the chance of an unrepre­ sented species arriving on that island decreases, and the likelihood of extinction of any one resi­ dent species increases. The species diversity of a given island can be assessed by projecting to the abscissa of the intersection point of the immigration and extinction curves, whereas the rate of turnover is reflected by the projec­ tion to the ordinates. Thus, species richness, together with immigration, extinction, and turnover rates, are all island–specific parame­ ters that change according to the area of the island and its isolation, with species richness remaining roughly constant over time, whereas species composition is continually changing (Fernandez–Palacios 2009). Although the rela­ tionship between S and A is well established, much discussion has focused on the value of the exponent z, which is a measure of the extent to which increases in area have dimin­ ishing returns in terms of species number and seems to be dictated by environmental varia­ bles and traits of the organisms. Thus, among

4  Biodiversity of Arthropods on Islands

insects, values reported for the exponent range considerably: 0.36 for carabid beetles (Nilsson et al. 1988) and 0.20 for butterflies in the West Indies, 0.14 for Sphingidae in the Malaysian archipelago (Beck et al. 2006), 0.20 for butter­ flies of the West Indies (Davies and Smith 1998), and 0.67 for butterflies from islands in the Baltic Sea (Itämies 1983). Slopes of the ­species–area relationships tend to be steeper on “true” islands relative to habitat islands, presumably because the isolating matrix is more permeable in habitat islands (Franzén et al. 2012). In addition, slopes vary according to traits of the organisms, being higher in fau­ nas with lower abundance and smaller range size (Franzén et al. 2012), in specialist species compared to generalists (Krauss et  al. 2003, Öckinger et al. 2010), in species with low ver­ sus high rates of reproduction (Franzén et  al. 2012), and in monophagous compared with polyphagous species (Steffan‐Dewenter and Tscharntke 2000). Although the size of an island is integral to understanding its biogeography, this in turn is linked to its geological history. Over the past few years, our geological understanding of islands has advanced enormously. For many oceanic archipelagos, the age of each island is often known with some level of precision, and information is generally available for changes in island size, elevation, and associated climatic factors over millennia (Gillespie and Clague 2009). At the same time, information on islands that have long since disappeared provides insights into historical patterns of connected­ ness. Studies on complexities of the splitting, sinking, and uplifting of the ancient landmass of Gondwana have highlighted the potential effect of the dynamic nature of island formation on patterns of biodiversity, as evidenced by arthropods in Southeast Asia (Polhemus and Polhemus 1998) and spiders in the Caribbean (Crews and Gillespie 2010). Thus, biologists have a temporal framework within which to examine ecological and evolutionary processes, and how biodiversity has formed on an island of a given area over time.

4.1.4  Age of the Island

Species diversity tends to increase with island age, at least up to a point. This relationship is well documented in the hotspot archipelago of Hawaii, where the number of species per unit area is much lower on the youngest island (Peck et  al. 1999, Gillespie and Baldwin 2010). For fragment islands, the initial conditions, such as the existing species diversity at the time of frag­ mentation and the available ecological niche space when the island effectively becomes, fully separated from the mainland, may result in a relative increasing or decreasing species diver­ sity (Diamond 1972). The longer the time since fragmentation, the more likely that the autoch­ thonous elements will be overwritten; thus, a continental fragment island such as Madagascar can come to be dominated by dispersal and neo‐ endemics (Yoder and Nowak 2006, Miraldo et al. 2011). For any given island system, the effects of his­ tory, isolation, habitable area, and age are inex­ tricably entwined and combined in unique ways. Thus, every island bears a signature of its past and possesses a mosaic of ancestral and derived features.

4.2 ­Ecological Attributes of Islands 4.2.1  Species Diversity on Islands

Given a suitable environment, the success of any particular colonization event will depend largely on the availability of ecological space and the ability of an organism to reach the island. For islands close to a source of propagules, the pat­ terns of colonization and approach to equilib­ rium have been studied in detail in the context of the Theory of Island Biogeography. As outlined above, this theory relates species and area by the formula S = cAz, (MacArthur and Wilson 1967). The primary predictions of the model are that (i) species numbers on an island should change little once the equilibrium is reached; (ii) there

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should be continual turnover of species, with some becoming extinct and others immigrating; (iii) small islands should support fewer species than do large islands; and (iv) species richness should decline with remoteness of the island because islands farther from the source will have lower rates of immigration.

distance dispersal are wind, birds, and ocean currents, the likelihood of a propagule being dispersed by a vector will depend on its mecha­ nism of association with the vector, coupled with its ability to withstand the environment to which it is exposed during transit, attributes that impose strong filters to the kind of organ­ isms involved (Gillespie et al. 2012).

4.2.2  Island Colonization

Detailed studies of immigration patterns have been conducted on mangrove islands in the Florida Keys, following artificial sterilization. Species of insects and spiders accumulated to an equilibrium number, although the turnover of species was not randomly distributed. Recently formed volcanic islands, such as Motmot Island in Lake Wisdom in Papua New Guinea, Surtsey near Iceland, or Krakatoa in the Java Straits, have been used to calculate immigration rates on eco­ logical timescales. On islands off Papua New Guinea, common early colonizers were largely widespread taxa, including a circumtropical dragonfly, a skipper butterfly, a chironomid midge, a caddisfly, a widespread anthicid beetle, a carnivorous earwig, a lycosid spider, and sev­ eral long‐jawed tetragnathid spiders (Edwards and Thornton 2001). Similar taxa characterized the early arrivals to Krakatoa (Thornton 1996). The surprising number of predatory arthropods on newly emerged islands might be due to al­lochthonous inputs, with recently colonizing spiders and collembolans relying on such inputs where communities lack primary productivity (Ingimarsdóttir et al. 2014). For more remote islands, the major filter is the ability of organisms to reach the island. Long‐distance dispersal, because of its infre­ quent and unpredictable nature, is difficult to study in the context of testable hypotheses (Gillespie and Baldwin 2010). However, although a single, rare, long‐distance dispersal event might be impossible to predict, an understand­ ing of the mechanisms involved in long‐distance dispersal over extended time periods can lend predictability to the process. In particular, assuming the three primary vectors for long‐

4.2.3  Factors Facilitating Establishment

Features of the new environment that affect the ability to establish will impose additional fil­ ters, with successful colonization and estab­ lishment being more likely in environments that approximately match the source environ­ ment (Hirao et  al. 2015). For example, organ­ isms from higher latitudes are most likely to become established at lower latitudes in higher‐ elevation habitats with climatic conditions matching their original environment. Similarly, lower elevations at low latitudes should be more easily colonized by taxa from tropical sites. In addition, for taxa that live inside other organisms or are associated with particular substrates, establishment probably can occur more readily if they arrive with (or after the establishment of ) their substrate, symbiont, or host. Even some highly specialized mutualisms are maintained (or recovered) following inde­ pendent colonization of remote islands by the different partners (Hembry et  al. 2013). An additional factor that can contribute to suc­ cessful establishment is escape from predators, parasites, and competitors, a phenomenon well documented through studies of successful invasions where predators are lacking (Gillespie and Roderick 2002). 4.2.4  Niche Preemption

In a given area, once a niche has been filled, it is apparently more difficult for ecologically similar individuals to enter (MacArthur and Wilson 1967). For example, lineages of Pheidole ants have relatively low dispersal rates (just sufficient to allow occasional long‐distance

4  Biodiversity of Arthropods on Islands

dispersal), and once the genus is established on an island, it is difficult for new congeners to establish and radiate (Economo et  al. 2015a). Similarly, such niche preemption also can con­ tribute to the “progression rule” of successive dispersal from older to younger islands in the Hawaiian Islands (Shaw and Gillespie 2016). Certain taxa can apparently still colonize occupied islands, as evidenced by successful invasions, although the interplay between dis­ turbance and competitive displacement in facilitating the establishment of new colonists is not always clear (Gao and Reitz 2017). 4.2.5  Ecological Release

What happens to a population subsequent to successful colonization? A common outcome is ecological (or competitive) release, with colo­ nizers expanding their range to fill the available ecological space. The extent and duration of any ecological release will depend on the isolation of the habitat: on islands that are close to a source of colonists, niche space will be filled quickly by diverse taxa; on isolated islands, the infrequency of colonization will allow the initial colonists to “explore” the ecological space much more broadly. Regular cycles of distributional change following colonization of islands have been pro­ posed several times in the literature, starting with the idea suggested by E. O. Wilson (1961) for a “taxon cycle” in Melanesian ants in which widespread, dispersive populations give rise to many more restricted and specialized popula­ tions or species. Recent work on Pacific island ants has supported this phenomenon (Eco­nomo and Sarnat 2012, Economo et al. 2015b). How­ ever, although there is a clear trend at least among ants, the consistency of distributional changes associated with the taxon cycle is less predictable (Gillespie et al. 2012).

governing interactions and changes thereof (Warren et al. 2015). The order and timing of arrival of colonists, and their ecological attrib­ utes, can have a large effect on interaction net­ works in all systems (e.g., Fukami et al. 2010). Studies examining the role of interaction net­ works in community assembly have focused on Krakatoa (Whittaker and Fernández‐Palacios 2007) and mangrove islands in the Florida Keys (Simberloff and Wilson 1971), the latter showing that species richness on islands is the result of a dynamic balance between stochastic immigration and extinction, and that the pro­ portion of specialist species increases during food‐web assembly relative to generalist spe­ cies (Piechnik et al. 2008). More recent work is starting to examine how interaction networks develop over evolutionary time among arthro­ pods on more isolated islands (Rominger et al. 2015).

4.3 ­Evolution on Islands 4.3.1 Anagenesis

On islands that are within the geographic dis­ tance to which a species is likely to disperse, the genetic coherence between populations from the source and those on the island can be main­ tained. In such circumstances, new species will tend not to form on the island (unless there is strong selection driving divergence). On islands that are beyond the range within which popula­ tions can maintain genetic contact with source populations, the island population will undergo evolutionary change, over time being replaced by another taxon without branching (anagene­ sis). This phenomenon has been highlighted by recent studies of crickets (Warren et  al. 2016) and spiders (Kuntner and Agnarsson 2011) in the Mascarene Islands.

4.2.6  Networks of Ecological Interactions

The processes that shape interaction networks are poorly understood, and because of this, islands have been used to determine factors

4.3.2 Cladogenesis

Where islands are farther from a source of colonists, fewer representatives from the entire

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source community will establish within the same time period. As a result, there is sufficient time for diversification and formation of new species characterized by tree‐like branching of taxa  (cladogenesis)  within the island (Warren et al. 2016). 4.3.3  Adaptive Radiation

The phenomenon whereby single (or few) colo­ nists, isolated genetically from their source pop­ ulation, give rise to a series of ecologically disparate species, often through a rapid burst of  speciation, is termed adaptive radiation. Adaptive radiation tends to be limited to islands that are beyond the so‐called “radiation zone,” or normal range of dispersal, of a given organ­ ism (Shaw and Gillespie 2016). Species that form through adaptive radiation are typically neo‐endemics, formed in situ and found no­­ where else. Adaptive radiation tends to be associated with the formation de novo of extremely isolated hab­ itats, with subsequent colonization being infre­ quent, giving the few successful colonists sufficient time to explore the ecological space available, diverge, and diversify into multiple species. Among arthropods, well‐known exam­ ples of adaptive radiation include Drosophila in Hawaii (Carson 1987, Magnacca and Price 2015) and multiple other insect (Zimmerman 1970, Howarth and Mull 1992, Liebherr 2015) and spider (Gillespie 2016) lineages on the archipelago. Many more recent radiations are coming to light, such as ant‐nest beetles in Madagascar (Moore and Robertson 2014); stick insects (Bradler et al. 2015) and weevils (Kitson et  al. 2013) in the Mascarene Islands; weevils (Hernández‐Teixidor et  al. 2016) and spiders (Arnedo et al. 2008) in the Canary Islands; spi­ ders in the Juan Fernandez Islands of Chile (Soto  et  al. 2017); darkling beetles in the Galápagos Islands (Finston and Peck 2004); car­ abids in Tahiti (Liebherr 2013) and New Zealand (Goldberg et al. 2014); and leafhoppers (Bennett and O’Grady 2013), bark lice (Bess et al. 2014),

and moths (Haines et al. 2014) in the Hawaiian Islands. Mechanisms of diversification are also com­ ing to light through recent studies of arthro­ pods, highlighting the roles of mating behavior in Drosophila flies, song repertoire in lineages of crickets (Oh et  al. 2013), host associations coupled with sexual signals in sap‐feeding planthoppers (Roesch Goodman et  al. 2012, Goodman et al. 2015), habitat in moths (Haines et al. 2014), and form of plant substrate in many beetles (Mckenna et  al. 2015). Diversification can follow a predictable pattern, at least in some groups; for example, among Tetragnatha spi­ ders, similar ecological sets of species have evolved over and over again on each of the dif­ ferent Hawaiian Islands (Gillespie 2016). Some of these radiations provide spectacular exam­ ples of the rapidity of adaptive diversification. For example, the estimated rate of diversifica­ tion for Hawaiian crickets in the genus Laupala is 4.17 species per million years (Mendelson and Shaw 2005), more than an order of magnitude higher than the estimated average rate of arthro­ pod speciation, 0.16 species per million years (Coyne and Orr 2004). Most island adaptive radiations occur in an archipelago setting, with allopatry between islands implicated in providing sufficient isola­ tion for adaptive radiation. In contrast, how­ ever, a radiation of small flightless weevils in the genus Miocalles (Coleoptera: Curculio­ nidae: Crypto­rhynchinae) has diversified within the single small island of Rapa in the southern Australs of French Polynesia (Paulay 1985). Rapa is home to almost half of the 140 species that occur across the western Pacific and Australia. Here, the beetles collectively feed on 24 genera of native plants and show varying degrees of host specificity, using almost all gen­ era of native plants on Rapa. 4.3.4  Isolation, Hybridization, and Admixture

In addition to rapid genetic and ecological differentiation, some new factors have come to

4  Biodiversity of Arthropods on Islands

light as being associated with adaptive radiation  – in  particular, the potential role of multiple colonizations and admixture in enhanc­ ing variability, as recently shown for spiders on the islands of the Galápagos (Hendrick et  al. 2015). A number of studies have demonstrated how the negative influence of genetic founder effects can be offset if different colonization events result in multiple genotypes in the intro­ duced population, highlighting the potential role of admixture among successively introduced populations in providing the genetic variation to allow adaptive evolution (Gillespie 2016). 4.3.5  Parallel Evolution and Convergence

Many of the best‐known island radiations are characterized by repeated shifts in phenotype and speciation, often associated with marked morphological and ecological convergence. This phenomenon, in which similar sets of ecological forms have evolved largely independently, has been well illustrated in spiders in the Hawaiian Islands (Blackledge and Gillespie 2004, Gillespie 2004).

4.4 ­Evolution in Other Insular Environments 4.4.1  Mountaintops – Sky Islands

The environments of mountaintops are often quite different from the surrounding slopes, and can include alpine grasslands, cloud forests, herbfields, and paramo; these sites tend to be relictual fragments of previously more wide­ spread habitats isolated by changing climate. Perhaps the best‐known sky islands, at least from a biological perspective, are the American Madrean sky islands of southeastern Arizona and New Mexico (Fig. 4.1g). Here, striking phe­ notypic divergence has been found among pop­ ulations of the jumping spider Habronattus pugillis (Masta 2000), as well as among scorpi­ ons (Hughes 2011), cerambycid beetles (Smith

and Farrell 2005), and carabid beetles (Ober and Connolly 2015). Given that these habitats are only about 10,000 years old, diversification has been rapid. 4.4.2 Caves

Caves (Fig. 4.1b) can be considered islands, with the taxa adapted to these systems isolated from the surface environment. Among cave spiders, beta diversity tends to be higher for assemblages of obligate cave dwellers (troglobionts), often with minimal gene flow between caves, whereas alpha diversity and phylogenetic and functional diversities tend to be low, as highlighted by spi­ ders in the caves of Appalachia (Hedin 1997a, 1997b). Cave‐adapted species are found among multiple different kinds of arthropods, and many cave species have evolved suites of mor­ phological traits associated with cave life, includ­ ing reduced or missing eyes, reduced or missing pigment, elaboration of appendages, and hyper­ trophy of extrasensory structures (Culver and Pipan 2014). Moreover, cave‐modified species tend to show recurrent evolution of similar forms, often through repeated evolution from surface forms, as shown in cave moths in Hawaii (Medeiros et al. 2009), carabid beetles in Texas caves (Gómez et  al. 2016), and cave spiders in the Canary Islands (Arnedo et al. 2008). Shifts may be controlled by the same genetic path­ ways, which can span many taxonomic levels; for example, adaptation to cave environments involves a similar mutation at the first step of melanin synthesis in planthoppers (from Hawaii and Croatia) (Bilandzija et al. 2012). 4.4.3  Desert Dunes and Salt Lakes (Fig. 4.1e)

Desert dune systems can also act like islands, although recent studies have shown that the unique assemblages of arthropods on the differ­ ent dunes is a product of rare jump dispersal events rather than vicariance (Van Dam and Matzke 2016). The restricted nature of taxa in  such dune systems has led to the possible

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extinction of several California insects, includ­ ing the Antioch Dunes katydid (Dunn and Fitzpatrick 2012), and to isolated and threatened populations, such as that of Doyen’s Trigonoscuta dune weevil (Williams et al. 1998). Similarly, the archipelago‐like system of salt lakes and isolated drainages in Australia’s interior are home to a radiation of tiger beetles (López‐López et  al. 2016). The deeper cladogenic events found in this study are correlated with the main period of continental acidification, and various molecular data‐based methods show a better fit to geogra­ phy than phenotype, reinforcing the island‐like nature of these lakes. 4.4.4  Habitat Fragments

The process of fragmentation has resulted in insularization at many different scales across space and time. In the Australian landmass, ref­ ugial rainforests dating from the Miocene are found in small pockets along the eastern and southwestern coasts, resulting in deeply diver­ gent arthropod lineages (Rix and Harvey 2012, Toussaint et al. 2014). The diversity of taxa that are confined to these habitats is extraordinarily high, with species often restricted to single mountains; the deep divergence between moun­ tains is due to the ancient fragmentation of for­ est habitats and subsequent diversification. Over the course of tens of millions of years, habitat specialization, coupled with climate change, can create second‐order islands within islands on relatively large, older fragment islands, such as New Zealand (Buckley et  al. 2015) and New Caledonia (Anso et al. 2016). On a more recent timescale, volcanic activity on the young island of Hawaii has created a dynamic mosaic of habi­ tats of different ages as large areas of forest are fragmented when lava flows around them. Fragmentation on scales of hundreds of meters and over several hundred years can be sufficient for genetic differentiation of arthropod popula­ tions (Vandergast et al. 2004), although – because of its transitory nature  –  this fragmentation is likely to be more important in maintaining

genetic diversity than in separation of popula­ tions (Gillespie 2016). Considering habitat islands that have been created within recorded history, many of the ideas originally developed for islands in the sea have been extended to these systems. Most such islands are fragments of habitats that were  historically connected, such as remnant trees and forest patches (Fig. 4.1d). For habitat islands, as for islands in the ocean, ecologi­ cal  and evolutionary processes are governed largely by isolation, time, and the nature of the matrix relative to the dispersal abilities of the organisms in question. Thus, in the context of the framework outlined above, habitat islands generally started with a full complement of species, and they tend to have a relatively small area that is less isolated than for many islands in the ocean because of the shorter distances involved and relative permeability of the sur­ rounding matrix. Habitat islands, because of their discrete nature and ease of manipulation, compared with islands in the sea, have been exploited in the development of many ecologi­ cal principles, including those related to meta‐ population dynamics and physical design of nature reserves.

4.5 ­Characteristics of Island Biodiversity A number of attributes characterize biodiversity on islands, in particular those that are more remote. 4.5.1 Disharmony

On remote oceanic islands, the biota is often considered to be “disharmonic” (Gressitt 1971, Gillespie and Roderick 2002, Gómez‐Zurita 2011). This phenomenon arises partly because taxa differ considerably in their ability to colo­ nize remote islands, leading to a filtering effect, whereby only a small set of taxa ever make it to a more isolated island; thus, large groups such

4  Biodiversity of Arthropods on Islands

as ants (Wilson and Taylor 1967) and mayflies (Zimmerman 1957) apparently have not been able to colonize remote oceanic islands east of Samoa in the Pacific Ocean. Adaptive radiation can accentuate the effect of disharmony when multiple species arise from the few that suc­ cessfully colonize, the result being few families and genera compared with the overall diver­ sity. This pattern, whereby the species diver­ sity  is represented by few higher taxonomic groups  (genera, families) and disproportion­ ate  numbers of species within a few genera, has been noted on many remote islands. Conti­ nental islands, including New Caledonia and New  Zealand, may also have many elements that are in disharmony as well as those that are in harmony. For example, some plants (e.g., Araucariaceae) and insects (e.g., ancestrally flightless carabid beetles; Will 2011) probably represent old harmonic elements, whereas ver­ tebrates (Darlington 1957) and some insect groups (Holloway 1979) seem to be dishar­ monic. Likewise, the Madagascar fauna (spi­ ders) is not disharmonic (Griswold 2004), perhaps due to its age and relative proximity to source areas. 4.5.2 Endemism

On islands formed de novo, the pattern of spe­ cies accumulation will depend on the rate of for­ mation of new species relative to the frequency of island colonization, in extreme cases leading to extensive adaptive radiation as a result of in  situ evolution with associated adaptation to occupy the available ecological space. On remote islands, the frequency of colonization becomes vanishingly rare. On fragment islands the initial state of diversity – number of species and relative saturation of ecological niches – will act to mediate effects of colonization, and the pattern of species accumulation will depend more on the rate of formation of new species in response to local adaptation and population fragmentation. Both situations can lead to very high levels of endemism.

4.5.3  Loss of Dispersal Ability and Flightlessness

Loss of dispersal ability is a common feature of organisms on more remote islands, and notably so in arthropods (Roff 1990, Wagner and Liebherr 1992, Zera and Denno 1997). The phe­ nomenon was first documented by Darwin, who suggested that “powers of flight would be injuri­ ous to insects inhabiting a confined locality, and expose them to be blown to the sea” (F. Darwin 1887, p. 45). However, not all species lose disper­ sal ability on islands. Indeed, loss of dispersal requires that (i) there be some dispersal behavior on which selection can act, and (ii) that there is no opposing selection to maintain dispersal abil­ ity. Thus, some strong‐flying insects can still maintain flightedness even on remote islands. Nevertheless, flightlessness in insects has evolved repeatedly on islands of all types (Darlington 1943, Vogler and Timmermans 2012). 4.5.4 Innovations

The known disharmony of islands seems to stimulate the development of some unusual traits in island groups. For example, expansion of host diversity seems to play a major role in the diversification of many island insects (Percy 2003). In addition, the absence of native social insects on some of the more remote islands of the Pacific might have been largely responsible for the adaptive shift to predation in some ter­ restrial insect groups and the diversity of other predatory arthropods on these islands (Howarth and Mull 1992). One of the most striking inno­ vations is the development of predatory behav­ ior in a lineage of Hawaiian caterpillar moths in the genus Eupithecia (Mironov 2014). Another innovation in Hawaii has been the development of odonates with terrestrial larvae, associated with the paucity of lakes in the upland moist for­ ests of the islands (Tennessen 2009). Islands fre­ quently lack native large mammals and the arthropods that typically depend on them. For example, native dung beetles are absent on more

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recent and remote islands such as Fiji, Hawaii, Samoa, and Tahiti. On older and less remote landmasses with endemic scarab beetles, such as New Caledonia and New Zealand, the beetles appear to have retained ancestral generalist feeding habits of the family or entirely shifted to being generalist feeders using non‐mammalian food sources such as fungi, fruit, and carrion (Jones et al. 2012). Generalized feeding and an ability to rapidly shift to new resources (Hanski et al. 2008) would enhance the chances of prop­ agules establishing and of established popula­ tions surviving significant changes. 4.5.5 Size

Change in body size is also commonly observed on islands, with species showing a tendency toward size extremes, both gigantism and dwarf­ ism, a phenomenon that has been called the “Island Rule”. Gigantism is best known among crickets: for example, wetas in New Zealand (Goldberg et al. 2008). However, it is also known among beetles, in particular long‐horned beetles in Fiji (Yanega et  al. 2004), earwigs on Saint Helena Island, and Indonesian stick insects (Chown and Gaston 2010), as well as sheet web spiders in Hawaii and the Juan Fernandez Islands (Hormiga 2002). In arthropod groups in which vagility is related to body size – for example, loss of flight leads to increase in mass and reduced dispersal, so that small‐sized individuals are more likely to colonize islands – the trend toward increasing size might be driven by the prepon­ derance of small ancestors (Liebherr 1988). However, favorable ecological niches available on newly colonized islands might also lead to size increase irrespective of ancestral size and vagility (Liebherr and Porch 2015). Island dwarf­ ism is less well known in insects, but has been documented in damselflies and dobsonflies on the islands off the coast of Japan (Hayashi 1990). 4.5.6  Reproductive Shifts

Reproductive shifts are well known to be asso­ ciated with island colonization in plants. In

insects, shifts to parthenogenesis on islands are known for some taxa. In particular, the Holarctic carabid Pterostichus empetricola from the Aleutian Islands and adjacent mainland might be the parthenogenetic sister species of the widespread Pterostichus brevicornis (Lindroth 1966, Eremin 1998, Bousquet 2012). The absence of males of P. empetricola and huge biases in the number of females in populations of P. brevicornis suggest that parthenogenesis evolved in these beetles on islands and small gla­ cial refugia islands (Ball 1966, Lindroth 1966). Parthenogenesis is also known in the damselfly Ischnura hastata from the islands of the Azores (Lorenzo‐Carballa and Cordero‐Rivera 2009).

4.6 ­Conservation Although islands have long proven them­ selves  as extraordinary laboratories for study­ ing processes associated with the generation of diversity, they are now contributing to our understanding of processes leading to the loss of diversity (Graham et al. 2017). The vulnerability of islands to extinction is now widely recog­ nized. Islands have suffered much higher rates of extinction than have continents, and although this has been most extensively documented for vertebrates, it is also known in insects, such as the Lord Howe stick insect (Mikheyev et al. 2017), the Saint Helena giant earwig (Labidura herculeana) and the giant carabid beetle, which seem to have gone extinct (Maunder et al. 1995, Ashmole and Ashmole 2004). Data from pre‐ human deposits on islands, long known to con­ tain diverse extinct assemblages of birds, also contain numerous arthropods, many now extinct, on the islands of Hawaii (Liebherr and Porch 2015) and the Australs and Cooks (Craig and Porch 2013). However, one of the most cru­ cial issues in island conservation is the lack of knowledge of the biota, coupled with the gener­ ally restricted areas of endemism, with associ­ ated small population sizes rendering them vulnerable to extinction. Compounding these problems are the large numbers of invasive

4  Biodiversity of Arthropods on Islands

species on islands (Samways 2013). Although these issues are similar to those confronting continental biotas, the microcosmal nature of islands makes them much more acute (Howarth and Ramsay 2012).

of species with small ranges arises because a local­ ized threat can affect their entire distribution, whereas those with low abundances are increas­ ingly susceptible to demographic stochasticity. 4.6.3  Abiotic Factors

4.6.1  Taxonomic Impediments

The biota of islands is often unique. For exam­ ple, the islands of the Pacific have been desig­ nated a biodiversity hotspot. Assessing this diversity, particularly for arthropods, is prob­ lematic. The major impediment is a lack of taxo­ nomic understanding of arthropods on many islands, particularly those that are more remote. New species are being collected at a remarka­ ble  rate in areas such as French Polynesia, Madagascar, and even the relatively well‐­studied Canary Islands, New Zealand, Hawaii, and the Galápagos; yet, the training of arthropod sys­ tematists has lagged behind, resources and job opportunities are lacking, and conveying the importance of fundamental taxonomic dis­ covery to policymakers and the general public remains challenging. Geographical and taxonomic biases have a large effect on the inferred undiscovered diver­ sity of insects (Jones et  al. 2009). Short of extended and extensive surveys across large areas of the most inaccessible parts of the world, addressing such biases is difficult, although some progress has been made using various species‐richness estimators to evaluate the number of species on islands (Hortal et  al. 2006). Further progress can be found in metrics such as completeness at high taxonomic levels, congruence with well‐established ecological relationships, and publication efforts to assess the reliability of species estimates in different groups of insects (Santos et al. 2010). 4.6.2  Restricted Ranges and Small Population Sizes

Species on islands tend to have limited geographic distributions and small population sizes, making them vulnerable to extinction. The vulnerability

Two major factors that disproportionately affect islands are habitat modification and cli­ mate change. Although natural habitat distur­ bances have apparently played a role in fostering diversification on remote islands such as Hawaii (Carson 1990), as evidenced by lava flows serving as a natural barrier to gene flow (Vandergast et al. 2004), recent modifica­ tions as a result of human activities have occurred much more rapidly (Florencio et  al. 2016). Moreover, because human activities tend to be associated with non‐native species, any habitat modification tends to facilitate invasion. Thus, although natural fragmenta­ tion as a result of lava flows still occurs on the youngest island of Hawaii, the taxa that colo­ nize the new habitat are frequently not native species (Gruner 2005), a consequence that seems to arise simply from the higher prop­ agule pressure presented by alien species as compared with native species. Climate change is a major environmental con­ cern worldwide. Islands and other insular envi­ ronments, such as mountaintops, tend to be highly climate‐sensitive, and a major concern is that climate change will compress ranges fur­ ther (Triantis et al. 2010b), leading to increased extinction rates of mountain species that have limited opportunities for migration (Harter et  al. 2015). Other concerns of climate change include increased extinction rates of mountain species that have limited opportunities for dis­ persal. The high‐elevation cloud forests that characterize many oceanic islands have a nar­ row geographical and climatological niche that can be eliminated with even a slight climatologi­ cal change. Climate change might also precipi­ tate declines in forests due to floods, droughts, or increased incidence of pests, pathogens, or fire. In addition, increases in the frequency or

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intensity of hurricanes can cause disturbance that favors invasive species. Small islands in particular share challenges associated with cli­ mate change and their exceptionally fragile environments (Konoe 2014). They are highly vulnerable to coastal erosion and saltwater intrusion into freshwater lenses, they have lim­ ited resources, and they are often isolated and exposed to cyclones and sea‐level rise. In addi­ tion, small islands frequently have high ende­ mism rates, but are often highly deteriorated, as a result of which their resilience to new aggres­ sions is limited. Furthermore, small island econ­ omies often rely on the quality of their natural environment, notably through tourism, fish­ ing,  and subsistence farming; a degradation of  their  environment could deeply affect local communities. 4.6.4  Invasive Species

The isolation of islands makes them more vul­ nerable to invasive species and other stresses, with extinctions attributed to alien species inva­ sion having been noted in several archipelagos (Wagner and Van Driesche 2010). Multiple fac­ tors interact to dictate invasion success, the most important being the following: (i) species diversity, which can serve as an ecological bar­ rier to invasion, an effect that has been used to explain the higher frequency and influence of invasions on islands; (ii) disturbance and the opening of ecological space; (iii) propagule pres­ sure, in particular the relative abundance of native versus non‐native propagules; (iv) char­ acteristics of propagules (more generalist in habitat requirements); and (v) novelty of pertur­ bations and “naïveté” of native biota. Generalist predators or competitors tend to have the great­ est impact as invaders, resulting in cascading effects. For example, the crazy ant Anoplolepis gracilipes has invaded Christmas Island, result­ ing in complete modification of the ecosystem (O’Dowd et al. 2003). Abundant, small, human commensals, such as ants and mosquitoes, are continually expanding their range onto oceanic islands and they apparently have a major adverse

effect on the native biotas on islands. Notable examples come from the effects on the native arthropods of ants in Hawaii (Krushelnycky and Gillespie 2008, 2010) and yellow jackets in New Zealand (Brockerhoff et al. 2010). Numerous insects have damaging effects on island vertebrates, perhaps the best known being mosquitoes as carriers of the organisms that cause diseases such as avian malaria and pox in Hawaii (LaPointe et  al. 2012) and the Galápagos (Levin et al. 2013). In the same way, introduced vertebrates such as cats, cane toads, coqui frogs, possums, and rats can have tremen­ dously damaging effects on native island arthro­ pods, although few studies have documented the effects.

4.7 ­Conclusion Islands have long provided intriguing insights into almost every aspect of arthropod biodiver­ sity, and indeed have spawned foundational understanding of ecological and evolutionary principles. The current rapid change affecting these systems provides us with an immediate relevance and need to develop a more complete understanding of the biodiversity in these sys­ tems, make use of the opportunity for under­ standing the shifting dynamic and, most critically, explore ways of mitigating the effects and potentially restoring ecosystems before they have gone beyond a state from which they will no longer be able to return.

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5 Beneficial Insects in Agriculture: Enhancement of Biodiversity and Ecosystem Services Matthew S. Jones and William E. Snyder Department of Entomology, Washington State University, Pullman, Washington, USA

When we think of insects in agriculture, we often concentrate on their negative effects. After all, herbivorous pests are estimated to consume 13% of annual crop production, at a cost of more than US$18.77 billion in lost production per year in the United States alone (and untold costs in terms of unmet food needs by local human populations) (Pimentel 2005, Losey and Vaughan 2006, Zhang et al. 2007). Some insects transmit pathogens of livestock and crop plants (Agrios 2005, Wales et al. 2010, Maclachlan 2011, Perry et  al. 2013), whereas others are livestock parasites (Lefèvre et  al. 2010). In some cases, these pathogens and parasites can also infect humans (Dorny et al. 2009, Newell et al. 2010, Thompson 2013). So, insects inflict considerable harm on agricultural production and, thus, on the world economy and human well‐being. Although these harms are undeniable, perhaps less appreciated are the many benefits that insects provide for agricultural production. More than 35% of the world’s food crops, and 15–30% of the average human’s diet, consist of plants that are pollinated by bees, flies, wasps, and other pollinators (McGregor 1976, Klein et  al. 2007). Likewise, predatory and parasitic insects and spiders consume large quantities of pest insects, delaying or sometimes eliminating the need for insecticide applications (Stern et al. 1959, Cohen et al. 1994). Insects are also key facilitators in the decomposi-

tion of dead plants and animals, returning nutrients to the soil for uptake by actively growing crop plants (Yokoyama et  al. 1991, Beare et  al. 1997, Bang et al. 2005). A subset of insect decomposers feeds on animal feces, improving soil fertility while removing a key source of contamination of fresh produce by human pathogens (Fincher 1981, Nichols et al. 2008, Jones et al. 2015). Collectively, these benefits for human food production are known as “ecosystem services” (Costanza et  al. 1997, de Groot et al. 2002), and modern agricultural ­systems are deeply dependent on them. We review the many ways that insect biodiversity influences each of these crucial insect‐mediated ecosystem services. In most cases, greater biodiversity increases the beneficial effects of insects, although in a few (relatively rare) instances, insect biodiversity degrades ecosystem services. At its simplest level, biodiversity is needed for all ecological roles of insects to be filled; at least one pollinator species, one predator, and one decomposer would be needed to minimally fulfill all of the crucial insect roles described above. We begin by describing how insect biodiversity at a finer scale might also influence the efficiency with which ecosystem services are delivered. We next describe harmful biodiversity effects, before closing with a discussion of how land‐use intensification harms insect biodiversity and how this damage might be reversed.

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

Insect Biodiversity: Science and Society

5.1 ­Components of Biodiversity: Species Richness, Species Evenness, and Species Identity Ecologists have recognized three primary components that must be measured and understood when quantifying biodiversity: the number of species, or species richness; the relative abundances of species, or species evenness; and the identities of the species in a community (Fig. 5.1). The first of these, species richness, is ideally tabulated by counting every species in a community (although in practice this must be done with care; Gotelli and Colwell 2001, Colwell 2009). Species richness is generally thought to be important simply because with more species present, a wider variety of ecological roles, or niches, probably will be filled (Kakehashi et al. 1984, Tilman 2000, Tilman et al. 2002, Cardinale et al. 2003). Evenness is colloquially referred to as the “­balance

of nature,” and is thought to be important because communities with greater evenness are more able to fill each of a diverse array of niches (Crowder et  al. 2010). Species identity is a key factor to consider because when ecological communities are more biodiverse, they are more likely, by chance alone, to include a particularly influential single species (Denoth et al. 2002). For example, a diverse biocontrol community is more likely to include an effective predator than is a less diverse community (Myers et al. 1989, Straub and Snyder 2006, Long and Finke 2014).

5.2 ­Why Does Insect Biodiversity Matter to Agriculture? Ecologists have long considered how diversity influences agricultural production, noting that the type of explosive pest‐insect outbreaks High

Low (b)

(c)

(d)

Evenness

(a)

Richness

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Figure 5.1  Species richness describes the number of different species present, irrespective of abundance, with low species richness (a) characterized by few species, and high species richness (b) characterized by many species. Species evenness describes the relative abundances of species in an assemblage, with low evenness (c) characterized by relatively more of particular species than others, and high evenness (d) characterized by relatively even abundances of the species present. Here we illustrate how, even when the number of individuals in an assemblage is held constant, levels of species richness and evenness can vary dramatically. Original figure by author.

5  Beneficial Insects in Agriculture

c­ ommonly seen in modern monoculture ­farming systems are rarely seen in more diverse natural communities (Pimentel 1961, Chapin et  al. 2000,  Macfadyen et  al. 2009). Root (1973) first described a specific mechanism that could underlie this observation: diverse plant communities could encourage an equally diverse and abundant community of predatory insects; in turn, predators with greater evenness and richness might lead to more dead pests. Similar issues have again surfaced in recent years on a global scale, as ecologists ponder the ecosystem effects of humans driving many species to extinction (Chapin et al. 2000, Green et al. 2005). This interest has spurred work on two specific mechanisms that can lead to a positive relationship between biodiversity and ecosystem functioning (roughly analogous to ecosystem ­ services)  –  complementarity and the sampling effect (Wardle 1999, Sokol‐Hessner and Schmitz 2002, Straub and Snyder 2006). We next describe how each of these mechanisms operates, alongside examples of each at work in pollinator, predator, and decomposer communities. We finish by discussing several mechanisms that can lead to negative biodiversity–service relationships, which run counter to, and sometimes might entirely erase, positive biodiversity effects. 5.2.1 Complementarity

Greater species richness and greater evenness among the species are thought to improve the delivery of ecosystem services through the same mechanism  –  fostering greater complementarity among species (Hooper et  al. 2005, Duffy et al. 2007). The perceived importance of complementarity derives from early ideas in ecology, particularly that to coexist, two species must occupy different niches. Thus, when two species occupy different niches, they complement one another to accomplish more ecological work in total than either species could accomplish on its own (Fridley 2001, Ives et al. 2005). Although pollinators, predators, and decomposers have many obvious differences from one another in the agroecosystem services

they deliver, species in each of these different functional roles c­omplement one another in similar ways. We next describe complementarity as it is most ­commonly known among agriculturally beneficial insects, acting through complementary ­ differences in spatiotemporal occurrence and activity, and through complementary foraging behaviors. 5.2.1.1  Temporal Complementarity

At the broadest temporal scale, insects might complement one another by exhibiting different, and at least partially non‐overlapping, differences in seasonal activity. For example, Neuenschwander et al. (1975) found seasonal complementarity in a community of predators attacking pea aphids on alfalfa. Several species of lady beetles provided strong impacts during relatively mild weather earlier in the season, whereas predatory bugs ­ were most active during the hottest part of the summer. Thus, only a diverse predator community provided attacks on aphids throughout the growing season. Similar to the benefits offered by the presence of different predators throughout the year, different pollinator species also often operate at different, and complementary, times of the year. For example, bumblebees (Bombus spp.) are active at lower temperatures (Fründ et al. 2013), earlier in the year, when honeybees (Apis mellifera) will not forage (Heinrich 2004). Thus, in blueberry crops a diversity of pollinators is important, each active at different temperatures, to enable efficient pollination throughout the bloom period (Tuell et al. 2009). A similar situation exists in dung beetle communities, wherein for rapid processing of dung throughout the pasture grass‐growing season and for maximizing grass yields, one species needs to be active early in the year, a second species active early in midyear, and a third species active later in the year. Bertone et al. (2006) collected dung beetles from North Carolina cattle pastures over an 18‐month period. ­ Although fewer beetles were collected during the winter, dung beetles were active all year round.

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Four  dominant species of dung beetles (Onthophagus hecate, Onthophagus pennsylvanicus, Onthophagus taurus, and Aphodius pseudolividus) were most abundant from March to October when fly breeding was highest (the flies breed in dung and are cattle pests) and forage production was greatest, whereas a host of other species provided manure removal throughout the rest of the year. In this system, year‐round cycling of feces into the soil could be realized only when early‐ and late‐season species co‐occurred. At a narrower scale – a single day – predation of a lepidopteran pest (Helicoverpa zea) of corn was maximized through the combined effects of two natural enemies, a lady beetle that foraged diurnally and a nocturnal predatory bug (Pfannenstiel and Yeargan 2002) (Fig. 5.2). Only with both enemies present were the moths deprived of temporal refuge from predation (Fig.  5.2c). Similarly, bees structure their foraging and pollination ser-

vices based on the time of day. Almost all species of bees differed significantly from each other in their preferred time of pumpkin‐flower visitation in the tropics (Hoehn et al. 2008). The species that visited flowers earliest in the morning were Apis cerana, Xylocopa confusa, and Xylocopa dejeani, whereas Ceratina cognata and Xylocopa nobilis appeared significantly later each afternoon. Therefore, a diverse community of pollinators, including both morning‐ and afternoon‐active species, was needed to ensure complete pollination of flowers opening throughout the day. Predators and pollinators can differ in their daily activity patterns, providing temporal complementarity on a relatively fine scale. 5.2.1.2  Spatial Complementarity

Agriculturally beneficial insects complement one another across time and space. Straub and Snyder (2008), for example, found that aphid pests of

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Figure 5.2  Temporal and spatial complementarity. (a–c) Temporal complementarity is the mechanism by which a lepidopteran corn pest (Helicoverpa zea) is most effectively controlled (Pfannenstiel and Yeargan 2002). During the day, a lady beetle is an effective predator of the moth eggs (a); at night, a predatory bug is an effective egg predator (b). Predation is maximized through the combined effects of two natural enemies; only with both enemies present are the moths deprived of a daily (temporal) refuge from predation (c). (d–f ) On collard plants, spatial complementarity among predator species leads to effective control of pest aphids only when multiple predator species co‐occur (Straub and Snyder 2008). This level of control is because some predator species forage mostly on leaf edges (d), while other predator species access aphids at the center of leaves (e). Only with a diverse predator community (f ) are all of the spatial refuges of the aphids removed. Original figure by author.

5  Beneficial Insects in Agriculture

collards (Brassica oleracea) were most effectively controlled by a diverse group of species of natural enemies, because some predators foraged mostly on leaf edges, whereas others accessed aphids at the center of leaves (Fig.  5.2d–f). Thus, only a diverse predator community eliminated the aphid’s entire spatial refuge (Fig. 5.2f). Similarly, sit‐and‐wait ambush spiders (Pisaurina mira) and actively hunting spiders (Phidippus rimator) provide predation in different parts of old‐field plant communities. The sit‐and‐wait P. mira occupies the upper canopy, and the actively hunting P. rimator the entire middle of the canopy; for this reason, both spider species must co‐occur for their grasshopper prey to lose a spatial refuge from predation (Schmitz 2009). In summary, when predators occupy spatially distinct hunting niches, they create a “no place to run, no place to hide” scenario for the prey. Pollinators also exhibit complementarity in a similar manner to that of predators. Bee species forage at different heights; therefore, multiple bee species are needed to pollinate flowers across the range of flower heights on each plant. Presumably, in the absence of the full complement of pollinator species, overall seed set, and thus fruit quality, would be reduced (Hoehn et  al. 2008). Blüthgen and Klein (2011) determined that different bee species pollinate either more exposed or more sheltered stigmas within the same individual flower. This type of architectural complementarity is important for the complete pollination of aggregate fruits; spatially uneven pollination can lead to asymmetrical or smaller fruits with reduced market value. Dung beetles also exhibit spatial complementarity in their feeding sites. The feeding behavior of dung beetles falls into three basic categories: tunnelers (paracoprids), dwellers (endocoprids), and rollers (telecoprids) (Giller and Doubet 1989). Tunnelers consume the inside of a dung pat and burrow into the soil directly beneath the pat, dwellers consume the pat and deposit eggs in the manure or in the soil near the surface, and rollers tend to break the pat into brood balls that are rolled to a suitable site and buried before ­oviposition. Vandamme (2014) found that when

species with different nest‐building behaviors were allowed access to cattle, horse, and sheep feces, there was a significant increase in the ­consumption and removal of the feces, compared with when only one functional group was allowed access. Although the above examples can be categorized as either spatial or temporal complementarity, several examples exhibit simultaneous complementarity across temporal and spatial niche axes. For example, the Colorado potato beetle (Leptinotarsa decemlineata), a key herbivorous pest of potato (Solanum tuberosum), is best controlled by diverse communities of natural enemies that attack the pests across space and time (Ramirez and Snyder 2009). The beetle has a complex life cycle; the larvae feed on plant foliage above ground before later burrowing into the soil to pupate. Above ground, the larvae are attacked by a diverse community of predatory insects, whereas below ground the larvae and pupae are attacked by a diverse community of insect pathogens. These predators and pathogens, attacking the beetles at different times and in different places, combine to kill more potato beetles than could either class of natural enemies on its own. The stress of escaping predators early in the life cycle apparently weakens the immune response that the beetles later rely on to combat infections by pathogens (Ramirez and Snyder 2009). 5.2.1.3  Behavioral Complementarity

Communities of insects that provide biocontrol, pollination, and decomposition services often represent species with distinctly different foraging behaviors. In turn, these behavioral differences can often lead to complementarity in the subset of prey, flowers, or detritus that different species use, resulting in greater overall delivery of ecosystem services when many species are present. For example, in communities of predators attacking the lepidopteran Cnaphalocrocis patnalis in rice (Oryza sativa), differences among predator species in the preferred life stage that is attacked lead to complementarity  –  and thus

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improved pest suppression  –  with increasing predator diversity (Wilby et al. 2005). This complementarity can be attributed to a species‐rich predator community providing predator species that separately focus attacks on adults, first‐ instar larvae, or fourth‐instar larvae; in the absence of predator species with these different food preferences, one or more stages of the pest are safe from predation (Wilby et al. 2005). Among pollinators, species often differ in the morphology of flowers in which each species prefers to forage. For example, despite blooming at the same time, watermelon and sunflower attract different flower‐visiting species, in part due to differences in flower morphology and pollen and nectar availability, which together influence flower attractiveness to different pollinator species (Klein et al. 2008). A diverse community of pollinator species, therefore, is needed to ensure that plant species with flowers of different sizes, colors, and shapes will be pollinated. Giller and Doubet (1989) found that a diverse community of dung beetles was needed to fully process dung pats as they aged. Soon after feces deposition, the pats were attacked by members of the tribe Coprini that prefer fresh feces ( 40 spp.) are assigned to the genus Phyllium, which seems to be paraphyletic with respect to members of the genus Chitoniscus, which again is recovered as polyphyletic in molecular studies (Buckley et al. 2009a, Bradler et al. 2015). 11.2.6 Heteropteryginae

The Heteropteryginae (by some referred to as Heteropterygidae, Zompro 2004a) are stout and robust phasmatodeans of the Oriental Region. There are more than 100 described species in approximately 30 genera ranging from small (Epidares from Borneo, ~4 cm) to large and bulky forms (Heteropteryx from Peninsular Malaysia, 13–16 cm). Most taxa are wingless ground‐dwellers that are well camouflaged in leaf litter. The Heteropteryginae were traditionally divided into four subgroups (Günther 1953, Klante 1976), of which the Malagasy Anisacanthini were excluded from the group by Zompro (2004a). Molecular data have supported this view (Bradler et al. 2015, Goldberg et  al. 2015), but monophyly of the remaining Heteropteryginae, consisting of the Datamini, Heteropterygini, and Obrimini, is not well established either. Some molecular studies showed the group to be polyphyletic (Whiting et al. 2003, Buckley et al. 2009a) or monophyletic based on low support (Tomita et al. 2011; Kômoto et  al. 2012; Bradler et  al. 2014, 2015; Goldberg et al. 2015). Morphological evidence is also vague, with derived characters present only in some groups, leading to a deficiently worked‐out ground pattern of the Heteropteryginae. All taxa insert their eggs into the soil, but only females of the Heteropterygini and Obrimini possess a

s­econdary ovipositor consisting of an enlarged epiproct (supraanal plate) and an elongated, tapered operculum (abdominal sternum VIII) (Bradler 2009). By contrast, the Datamini and Obrimini bear a pair of prominent sensory areas on the basisternite of the prothorax, which is missing in the Heteropteryginae (Zompro 2004a, Bradler 2009). All Datamini and the majority of Obrimini are wingless, with the exception of Miroceramia from Sulawesi, the taxonomic position of which is contentious (Bradler 2009), and Pterobrimus from Fiji, with diminutive wing remnants, whereas members of the Heteropterygini always bear wings. Of all the conceivable relationships among those three lineages of the Heteropteryginae, two have been favored in the past (Fig. 11.5a,b), based on morphological or molecular data, with either the Heteropterygini as sister to Obrimini + Datamini (together: Obriminae, Rehn and Rehn 1938) (Zompro 2004a, Goldberg et  al. 2015) or the Datamini as sister to Heteropterygini + Obrimini (Klante 1976 who also included Anisacanthini, cf. Fig. 11.5c, Bradler 2009, Bradler et  al. 2015). The latter topology would indicate multiple losses of wings among the Heteropteryginae or the regain of wings after ancestral loss in the Heteropterygini, as proposed before (Whiting et al. 2003, Bradler 2009). Usually, wings in the Heteropterygini are short and only used for defensive sound production, with a certain area in the hindwing forming a timbal organ (Carlberg 1989, Bradler 2009). This highly derived modification is found in both sexes of Haaniella and in females of Heteropteryx. By  contrast, the male of the large arboreal Heteropteryx from Peninsular Malaysia is capable of flight and exhibits the most plesiomorphic condition of tegmina m ­ orphology among extant stick insects. The f­orewings are not shortened and cover most of the abdomen and the equally well‐ developed hindwings at rest, and the radial sector or posterior radius of the tegmina is two‐branched; these two traits are unique among extant forms but are repeatedly reported from extinct stem‐ group phasmatodeans (Willmann 2003, Bradler

11  Biodiversity of Phasmatodea Obriminae

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Figure 11.5  Alternative phylogenetic hypotheses of Heteropteryginae subgroups, following different authors. (a) Zompro 2004a, Goldberg et al. 2015. (b) Bradler 2009, Bradler et al. 2015, Hennemann et al. 2016b. (c) Klante 1976. (d) Bradler (unpublished). The Heteropteryginae are sometimes also referred to as the Heteropterygidae, with the corresponding subgroups being the Heteropteryginae, Obriminae, and Dataminae (based on rank escalations proposed by Zompro 2004a). Original by authors.

2009, Shang et al. 2011, Wang et al. 2014). Because females of Heteropteryx are canopy‐dwellers, which still deposit eggs into the ground, this species most probably became secondarily arboreal. This scenario is also supported by the phylogenetic position of Heteropteryx; all ambiguity aside, this genus is always recovered as a subordinate taxon among the Heteropteryginae, suggesting that the male wing of Heteropteryx might be the result of an atavistic character reversal.

11.2.7 Diapheromerinae

The Diapheromerinae are the most species‐rich lineage of stick insects in the New World, with more than 350 species described in approximately 40 genera, ranging from small (Ocnophiloidea, 4 cm) to large (Megaphasma, 15 cm) and giant‐ sized species (Bacteria and Otocrania, > 22 cm). All species are anareolate with long antennae. The majority of taxa are slender and wingless, with

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some having winged males (Bacteria and Pterolibethra). The Diapheromerinae comprise three subgroups, two of which are minor, the Ocnophilini and Oreophoetini, with less than 40 species described in each. By far the most diverse group is the Diapheromerini, containing more than 300 species. Whether these groups reflect monophyletic taxa is not clear. Preliminary studies suggest either a sister‐group relationship between Ocnophilini + Oreophoetini and Diapheromerini (Bradler et  al. 2014) or Ocnophilini as sister to Diapheromerini, with Oreophoetini nested within the latter, rendering Diapheromerini paraphyletic (Bradler et  al. 2015). One monophyletic group  among Diapheromerini taxa is the Eusermyleformia (Diapheromera, Pseudosermyle, and allied genera), characterized by the absence of the male vomer, specialized cerci that form claspers for copulation, and abdominal stigma VIII shifted backward (Bradler 2009). Some analyses recover paraphyletic Diapheromerinae at the base of the euphasmatodean tree (Whiting et al. 2003; Bradler et al. 2003, 2014). One peculiar trait that could be interpreted as being particularly plesiomorphic among the Euphasmatodea is the large gonoplac of the female ovipositor (Bradler 2009). By contrast, other studies suggest the Diapheromerinae to be a monophyletic and subordinate lineage among the Phasmatodea (Buckley et al. 2009a, Bradler et al. 2014, Goldberg et  al. 2015). The phylogenetic status and taxonomic boundaries of the Diapheromerinae are in need of clarification and await a more comprehensive survey. 11.2.8 Pseudophasmatinae

Besides the Diapheromerinae, the Pseudophasmatinae (referred to by some as Pseudophasmatidae, Zompro 2004a) is the dominant group among Neotropical stick insects. This subfamily comprises approximately 300 described species in more than 50 genera that are distributed across South and Central America, the Caribbean Islands, and the southern part of the United States. Members of the Pseudophasmatinae are small‐ to medium‐sized

and often winged, but there are also numerous brachypterous and apterous forms, frequently with well‐developed defensive glands and aposematic coloration (Eisner 1965). Several species exhibit extreme mate‐guarding behavior, with small males being carried for several weeks on the back of the much larger females (Thornhill and Alcock 1983). Günther (1953) considered Agathemera, Dajaca (Aschiphasmatinae), and taxa now assigned to the Heteronemiinae to belong to the Pseudophasmatinae. Besides those few formerly misplaced taxa, the bulk of the species form a well‐supported monophylum based on molecular data (Whiting et al. 2003; Buckley et  al. 2009a; Bradler et  al. 2014, 2015). They are also characterized by derived mouthparts; for example, the maxillae bear a lancet‐shaped galealobulus, and the trichome area of the galea is shifted onto the inner side (Bradler 2009). This trait combination is also found in Prisopus, Melophasma, and allies, which were excluded from the Pseudophasmatinae by Zompro (2004a), who erected a new family for these taxa: Prisopodidae. Zompro (2004a) also assumed a closer relationship of the Prisopodidae to the Oriental Aschiphasmatinae and Korinninae and to the Malagasy Damasippoididae, but these assumptions have all been refuted by recent molecular studies (Bradler et al. 2015, Goldberg et al. 2015). The Prisopodidae genus Melophasma has been unambiguously placed in the Pseudophasmatinae clade (Goldberg et al. 2015). 11.2.9 Palophinae

With barely 20 species described in two genera, the Palophinae are not diverse, but comprise the largest stick insects in continental Africa, often exceeding 20 cm in body length, with the female of Bactrododema hippotaurum measuring more than 26 cm (Sjöstedt 1913). Its members are winged and bear ocelli and a characteristic pair of closely spaced, prominent processes on the vertex of the head. The phylogenetic position of the Palophinae is nebulous, with molecular analyses placing Bactrododema either as sister group to the New World Cladomorphinae (Buckley

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et  al. 2009a) or to the Asian Necrosciinae (Bradler et  al. 2015). The Palophinae do not show any obvious affinities to other African phasmatodeans. 11.2.10  The African Clade

At present, approximately 105 described species form a lineage of African‐European stick insects, of which more than 75 are distributed in Madagascar alone (Table 11.1). The described Malagasy species might represent only a small percentage of the actual species richness. Four distinct taxonomic groups are currently recognized on Madagascar, the Achriopterini, Anisacanthidae, Antongiliinae, and Damasippoididae (Cliquennois 2007), and one taxon incertae sedis, Spathomorpha (Cliquennois 2005). These groups are endemic to Madagascar and the Comoros. The Malagasy stick insects are morphologically diverse and range from small to giant‐sized (between 30 and 240 mm), with most members being flightless (either wingless or brachypterous). Recent taxonomic revisions of Malagasy subgroups (Cliquennois 2006, 2008) contain descriptions of several new taxa, but are devoid of any formal phylogenetic analysis. New data suggest that these taxa are more closely related to each other than previously thought, as the result of one or two colonization events and subsequent adaptive radiation in Madagascar and the Comoro Islands, but without colonization of the Seychelles or Mascarene Islands (Bradler et  al. 2015). Affinities of Malagasy taxa to those from either the Oriental Region or the Neotropical Region have been suggested in the past (Hennemann and Conle 2004, Zompro 2004a). Molecular data have now revealed affinities to African and Mediterranean stick insects traditionally assigned to the Bacillinae (Bacillus, Clonopsis, Macynia, Phalces, and Xylica) and some African members of the Gratidiini, which uniformly are small‐ to medium‐sized and wingless stick insects with often remarkably short antennae (Bradler et  al. 2015; S. Bradler et  al., unpublished data). Most taxa in this African clade are areolate, but some anareolate species

are closely related to these taxa, such as the anareolate Achriopterini forming the sister group of the areolate Anisacanthidae (Bradler et  al. 2015), and the anareolate Spathomorpha being probably related to the areolate Antongiliinae (Cliquennois 2007). The same pattern is found in the exhaustively studied European phasmatodeans, which apparently form another well‐supported clade (Scali et  al. 2013), comprising the areolate genera Bacillus and Clonopsis and the anareolate Leptynia and Pijnackeria. 11.2.11 Gratidiini

The Gratidiini comprise 17 genera and approximately 180 described species, with the huge majority (> 120 spp.) assigned to the genus Clonaria (formerly Gratidia), members of which are impressive grass mimics in the African savanna. The Gratidiini are gracile, medium‐sized stick insects usually ranging between 5 and 8 cm, with short antennae. The Gratidiini can also be found across a wide geographical area ranging from Europe (Leptynia and Pijnackeria) to Southeast Asia (Clonaria and Sceptrophasma), but there are fewer species across this range when compared with the numerous species in Africa. One peculiar trait of this group is a derived egg‐laying behavior displayed by the females of many (but not all) Oriental and African taxa: The female bends its abdomen over the thorax and head with the abdominal tip positioned between the short antennae, which then assist with placing the elongated adhesive eggs on blades of grass or other suitable plant substrate (Brock and Shlagman 1994). The phylogenetic affinities of the Gratidiini have been poorly investigated, with few Asian and African taxa included in past studies. Oriental Gratidiini show strong affinities to the Asian Clitumnini + Medaurini, either as the sister group (Buckley et al. 2009a, Bradler et al. 2014) or nested within it (Whiting et al. 2003, Bradler et al. 2015). By contrast, the Mediterranean Leptynia and Pijnackeria and the African Clonaria spp. seem to be more closely related to the Bacillinae (Scali et al. 2013,

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Bradler et al. 2015), thus being part of the large African clade discussed above. In consequence, the Gratidiini might not be monophyletic and the derived egg‐laying strategy has evolved two times independently, which is in need of more extensive documentation. 11.2.12 Clitumnini

The Clitumnini comprise approximately 240 described species of anareolate, slender, and wingless stick insects of medium to large size, geographically distributed from India to Southeast Asia. Most recently, a Vietnamese genus (Lobofemora) has been reported with winged males (Bresseel and Constant 2015). Of the 15 currently recognized genera, Ramulus is by far the most diverse among the Clitumnini, containing more than 170 of the known species. Ramulus comprises the most gracile stick insects (Ramulus nematodes) and numerous taxa from more northern Oriental regions such as China, Korea and Japan. The Clitumnini are probably the sister group of the Medaurini (Bradler et al. 2014) or Medaurini + Asiatic Gratidiini (Whiting et al. 2003, Bradler et al. 2015). 11.2.13 Medaurini

The Medaurini are a species‐poor group of medium‐sized, anareolate, and wingless stick insects that comprise five genera and approximately 30 described species that inhabit more or less the same Southeast Asiatic regions as do the Clitumnini, which seem to be closely related (Whiting et al. 2003; Bradler et al. 2014, 2015). 11.2.14 Pharnaciini

This group of anareolate stick insects from Southeast Asia consists of giant‐sized, strongly elongated canopy‐dwellers, with Phobaeticus kirbyi (32.8 cm) from Borneo considered to be the largest insect in the world for a long time (Brock 1999, Bradler 2003), until recently replaced by Phobaeticus chani (35.7 cm), which was also described from Borneo (Hennemann

and Conle 2008). Five genera and nearly 50 species are currently recognized. Females are always apterous, whereas males can be either apterous or brachypterous, rarely fully winged. Both sexes are devoid of ocelli. Molecular studies have placed the Pharnaciini as either related to Cladomorphinae + Stephanacridini + Lanceocercata (Whiting et al. 2003; Kômoto et  al. 2011, 2012; Tomita et  al. 2011; Bradler et al. 2014) or with closer affinities to the Clitumnini + Medaurini (Bradler et  al. 2015). From a morphological point of view, the latter placement appears to be more convincing, as the Clitumnini, Medaurini, and Pharnaciini (together: Clitumninae sensu Hennemann and Conle 2008) share the same derived morphology of the male terminalia. Abdominal tergum X is longitudinally split into movable hemitergites with a ventral articulation and spines on the inner sides, serving as claspers for holding the female abdomen during copulation (Hennemann and Conle 2008, Bradler 2009). 11.2.15 Cladomorphinae

The anareolate New World Cladomorphinae form a diverse group of more than 100 described species in 30 genera ranging from giant‐sized canopy dwellers with well‐developed wings (Diapherodes gigantea, measuring up to 19 cm long) to minute, wingless, ground‐dwelling forms (Tainophasma, ~3 cm body length). Ocelli are absent even in volant forms. The Cladomorphinae underwent a largely unexplored adaptive radiation mainly in continental Central America and the Caribbean, with numerous species distributed across the Greater and Lesser Antilles. The Cladomorphinae have been repeatedly corroborated as being related to the Lanceocercata (Whiting et al. 2003, Buckley et al. 2009a), or more precisely as the sister taxon of the Australasian clade Lanceocercata + Stephanacridini (Buckley et  al. 2010; Bradler et al. 2014, 2015). Yet the taxonomic boundaries of the Cladomorphinae are largely unexplored, and the possibility cannot be excluded that some members belong to the Diapheromerinae. However, the idea that a subgroup of the

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Cladomorphinae, the Hesperophasmatini, forms a subordinate lineage among the areolate Pseudophasmatidae (Zompro 2004b) has been repeatedly falsified based on morphology (Bradler 2009, Hennemann et  al. 2016a) and molecular data (Buckley et  al. 2009a; Bradler et  al. 2014, 2015). 11.2.16 Stephanacridini

With barely 40 described species in eight genera, the Stephanacridini are another species‐poor clade of anareolate large‐ to giant‐sized stick insects from the Oriental Region (Hennemann and Conle 2006). Extending from Malaysia over New Guinea and Australia to the Fiji islands, the Stephanacridini are unambiguously placed as the sister group of the diverse Lanceocercata, which show a comparable geographic distribution (Buckley et  al. 2009a, 2010; Bradler et  al. 2014, 2015; Goldberg et  al. 2015). Females are either brachypterous or fully apterous, whereas males have well‐developed wings but no ocelli (Hennemann and Conle 2006, Bradler 2009). Females of the Stephanacridini bear an elongated operculum or subgenital plate (abdominal sternum VIII) with similarly elongated ovipositor valves (gonapophyses VIII) (Bradler 2009). Males possess a vomer, which became lost in its sister group Lanceocercata. 11.2.17 Lanceocercata – The “Marsupials” Among the Phasmatodea

The morphologically and ecologically diverse clade Lanceocercata contains a wide array of Australasian phasmatodeans conventionally thought to be unrelated to one another, comprising species of the Eurycanthinae, ­ Pachymorphinae, Phasmatinae, Platycraninae, Tropidoderinae, and Xeroderinae, with all these traditional subfamilies recovered as polyphyletic (Bradler 2009; Buckley et al. 2009a, 2010; Bradler et al. 2015). The Lanceocercata comprise approximately 220 species in 70 genera. Originally proposed based on a combination of derived characters from the male and female

t­ erminal region (Bradler 2001), the Lanceocercata have been repeatedly corroborated in phylogenetic studies ever since (Whiting et  al. 2003; Bradler 2009; Buckley et al. 2009a, 2010; Kômoto et al. 2012; Dunning et al. 2013; Bradler et al. 2014, 2015; Goldberg et  al. 2015). The Lanceocercata radiated not only in Australia and adjacent areas (e.g., New Guinea), but also from as far west as the Mascarene Archipelago and the Seychelles in the Indian Ocean and as far east as New Caledonia, Fiji, and New Zealand (Buckley et al. 2010, Bradler et  al. 2015), and exhibit morphological and ecological parallelisms comparable with those found between placental mammals and marsupials (Springer et al. 1997). The Lanceocercata and the remaining Euphasmatodea underwent parallel adaptive radiations with astounding examples of convergence (Fig. 11.1, Fig. 11.2, Fig. 11.6). The Lanceocercata comprise giant, winged stick insects of the canopy, with well‐developed ocelli, such as the Australian Acrophylla, exceeding 26 cm in body length, which is paralleled by the equally large, morphologically and ecologically similar African Bactrododema (Palophinae). The leaf‐imitating forms include Malandania and Tropidoderus in the Lanceocercata (imitating Eucalyptus leaves) and true leaf insects in the Phylliinae (Fig. 11.2). Gracile flyers are also found in the Lanceocercata (e.g., Carlius), which are strikingly similar to the  forms in the Necrosciinae (e.g., Sipyloidea). In  addition, the small wingless Australian Lanceocercata with exceedingly short antennae (e.g., Pachymorpha) are highly reminiscent of the Afro‐Oriental Gratidiini (e.g., Sceptrophasma, Clonaria). Examples among diminutive spiny trunk‐dwellers with a body length of less than 4 cm include Cnipsus in the Lanceocercata and some Neopromachus spp. in the Lonchodinae. The most impressive convergence is found among the “tree lobsters” or “land lobsters” (Fig. 11.6)  –  large, stout, flightless, ground‐dwelling, brownish phasmatodeans, which have a dorsoventrally flattened body and sturdy legs, and congregate in large numbers and close spatial proximity in tree hollows and cavities during the day (Buckley et al. 2009a). Females deposit their eggs into the

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Figure 11.6  Female “tree lobsters.” (a) Eurycantha horrida (Lonchodinae) from Papua‐New Guinea (photograph by Michael F. Whiting, Provo). (b) Dryococelus australis (Lanceocercata) from Lord Howe Island (photograph by Thomas Reischig, Göttingen). Traditionally these ground‐dwelling phasmatodeans were considered to be closely related and were referred to as the Eurycanthinae. However, these forms are unrelated to each other and the result of convergent evolution.

soil, and the greatly enlarged and armed hind legs of some males probably evolved as a response to ground‐hunting predators, and might also be used against other males. Tree lobsters were traditionally assigned to the subfamily Eurycanthinae (Günther 1953), but molecular analyses of its members revealed that tree lobster ecomorphs evolved three times independently, two times in the Pacific Lanceocercata and once in the New Guinean Lonchodinae (Buckley et  al. 2009a, 2010). The overall uniformity of tree lobsters with regard to habitus and behavior is probably the product of similar selective pressure associated with adaptations to ground‐dwelling life. Beyond those extrinsic factors, such phenotypic similarity is probably also generated by intrinsic factors such as shared trajectories in the underlying developmental architecture (Brakefield 2006), which

might provide constraints on the direction of stick insect evolution. The most famous tree lobster is Dryococelus australis from Lord Howe Island (Fig. 11.6b), which was assumed to have become extinct when black rats were introduced to the island in 1918, but was rediscovered on a remote offshore rock, Ball’s Pyramid, in 2003 (Priddel et al. 2003; see also Cranston, volume 1, chapter 6). D. australis descended from tree‐dwelling Australian Lanceocercata (Buckley et al. 2009a, 2010). Another lineage of tree lobsters, Canachus and  related genera, originated independently as part of a New Caledonian radiation of the Lanceocercata (Buckley et  al. 2010). In consequence, a number of convergent forms not only evolved between the Lanceocecata and non‐­ lanceocercatan stick insects, but also within separate  Lanceocercata lineages. For instance, the

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Mascarene Apterograeffea, traditionally viewed as a member of the Platycraninae (coconut stick insects and allies), exhibits the same enlarged genae (“cheeks”) to accommodate the massive mandibular muscles that are necessary for feeding on hardy palm or pandanus leaves, as do the unrelated Australian and Pacific ecotypes, such as Graeffea and Megacrania (Bradler et al. 2015). The females of the Mascarene Rhaphiderus spp., hitherto assigned to the Tropidoderinae, are wingless, leaf‐imitating forms similar to the Australian Malandania and Tropidoderus (Buckley et  al. 2010). On Mauritius, Epicharmus, traditionally assigned to the Xeroderinae, is a stout, winged stick insect with lobes on the legs and abdomen, which enhance crypsis of this bark‐dwelling species; it is most reminiscent of the unrelated Australian Xeroderus or New Caledonian Leosthenes (Bradler et al. 2015). The relationships within the Lanceocercata have been repeatedly explored in recent years, leading to some robust phylogenetic hypotheses. Dimorphodes (subfamily Xeroderinae) from New Guinea and adjacent landmasses constitutes the sister group to all other members of the Lanceocercata, which comprise some well‐ supported lineages, such as a New Caledonia + New Zealand clade (with possibly two independent colonizations of the latter) and a fully unexpected Mascarene lineage (Buckley et  al. 2009a, 2010; Dunning et al. 2013; Bradler et al. 2014, 2015). 11.2.18 Lonchodinae

The Lonchodinae are an anareolate, species‐ rich lineage of generally wingless and slender stick insects of average size (7–11  cm, Fig.  11.1a). Larger and more robust forms are found among the New Guinean tree lobsters (females of Eurycantha measuring up to 16 cm, Fig. 11.6a). More than 330 Lonchodinae species are described in 40 genera, extending from the Seychelles and India throughout Southeast Asia to Northern Australia. The Lonchodinae include the Indian stick insect Carausius morosus, also known as the laboratory stick

insect. For more than a century, C. morosus has served as a model organism in an impressive number of biological studies dealing with physiological color change (e.g., Atzler 1930, Bückmann and Dustmann 1962, Willig 1969), developmental biology (e.g., Stratakis 1976, Bücker et  al. 1986, Masetti and Giorgi 1989), neurobiology (e.g., Tichy 1979, Ashcroft and Stanfield 1982), and in numerous studies on leg movement, locomotion, and related bionics (e.g., Cruse 1976, Cruse and Saxler 1980, Büschges 1990, Dürr et al. 2004, Schmitz et al. 2015, Theunissen et al. 2015). The majority of the Lonchodinae stick insects are bush‐ or tree‐dwelling, with the exception of the Eurycanthomorpha (Bradler 2002), a monophyletic New Guinean subgroup of the Lonchodinae (Bradler et al. 2014), which became adapted to a bark‐ and ground‐dwelling life style. Members of the Eurycanthomorpha developed a secondary ovipositor consisting of an enlarged abdominal tergum X and operculum (abdominal sternum VIII) for placing eggs into soil or substrate alike (Fig. 11.6a, Bradler 2009). The most specialized Eurycanthomorpha are found among the tree lobsters in the genus Eurycantha, which, with respect to body form and biology, are highly reminiscent of the tree lobsters in the Lanceocercata (Buckley et al. 2009a, Fig. 11.6). The phylogeny of the Lonchodinae remains largely unresolved. Preliminary molecular analyses provided tentative support for a sister‐group relationship between the Lonchodinae and Necrosciinae (Bradler et  al. 2014). Within the Lonchodinae, the Eurycanthomorpha are either the sister taxon of all remaining Lonchodinae (Buckley et al. 2009a, Bradler et al. 2015, Goldberg et al. 2015) or a subordinate lineage of the latter (Bradler et al. 2014). Furthermore, a few taxa that were previously placed in the Necrosciinae (Baculofractum and Leprocaulinus), owing to their winged males, form a subordinate group in the otherwise apterous Lonchodinae (Bradler et al. 2014). This phylogenetic pattern indicates a potential regain of wings after ancestral loss, an evolutionary scenario suggested before for phasmatodeans in general (Whiting et al. 2003).

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11.2.19 Necrosciinae

The Necrosciinae are the most species‐rich traditional subfamily, currently comprising approximately 700 described species in nearly 90 genera from the Oriental Region, including approximately 30 Australian species (Brock and Hasenpusch 2007) and one invasive species, Sipyloidea sipylus, in the Mascarene archipelago and Madagascar (Cliquennois 2012). Members of this lineage are mostly gracile flyers of medium size with anareolate, slender legs and long antennae. Few species are wingless or brachypterous and can reach larger body sizes; for example, some females of Phaenopharos exceed 14 cm in body length. Sellick (1997) considered the Necrosciinae (referred to as Necrosciidae by the author) to be polyphyletic, and described the group as being extremely heterogeneous in light of egg‐capsule morphology and egg‐laying strategies. The available cladistic morphological analyses (Tilgner 2002, Bradler 2009) did not recover monophyletic Necrosciinae. Not surprisingly, to date not a single apomorphic character supporting monophyly of the Necrosciinae has been reported. Nevertheless, monophyly of the Necrosciinae has been repeatedly corroborated in molecular studies (Whiting et al. 2003, Buckley et al. 2009a, Bradler et al. 2015). More comprehensive studies showed that a few genera were taxonomically misplaced and belong to the Lonchodinae, and some wingless taxa formerly assigned to Lonchodinae (Neohirasea and allies) are in fact members of the Necrosciinae (Bradler et  al. 2014). Most surprisingly, the Necrosciinae also encompass the species‐poor Korinninae. This is particularly noteworthy because the Korinninae and Necrosciinae were considered to belong to the two different suborders of Phasmatodea, the Areolatae and Anareolatae. The subfamily Korinninae was erected by Günther (1953), based on the genera Korinnis and Kalocorinnis, which currently comprise only seven described species from Borneo, Thailand, and the Philippines (Bragg 2001, Zompro 2004a, Gottardo 2008). Originally considered to be an

“isolated” taxon without any obvious relationships to other phasmatodean subfamilies (Günther 1953), recent classifications placed the Korinninae either as the sister group to the areolate Oriental Aschiphasmatinae (Bragg 2001) or as sister to the areolate Neotropical Prisopus + related genera, the Prisopodinae sensu Zompro (2004a). Molecular analyses refute any close relationship of the Korinninae to either of these groups (Goldberg et al. 2015). The Korinninae have evolved a unique egg‐laying strategy among the Phasmatodea. In contrast to the majority of stick and leaf insects that disperse single eggs, females of the Korinninae produce a complex egg‐case or ootheca (Fig. 11.7), which is glued to arboreal substrate (twigs and leaves) and carries more than 30 eggs in a highly ordered arrangement (Goldberg et  al. 2015). Ootheca formation bears a number of adaptive advantages likely related to protection against parasitoids and desiccation, and to allocation of specific host plants. The predominantly volant and highly mobile members of the Necrosciinae are often food‐plant specialists (Blüthgen et  al. 2006, Junker et al. 2008).

11.3 ­The Phasmatodean Fossil Record The stick and leaf insects exhibit a remarkably poor fossil record. Besides a well‐preserved male leaf insect, E. messelense, from the Eocene of Germany (˜47 mya) (Wedmann et al. 2007), no adult fossil specimen of any extant euphasmatodean lineage has been described so far. Juvenile stick insects, mostly areolate, that cannot be assigned to any known taxon are repeatedly found in Baltic and Dominican amber inclusions (Tilgner 2001, Poinar 2011). A number of fossilized eggs were reported from the Eocene of Oregon (Clark Sellick 1994) and Dominican amber (Poinar 2011), which have been assigned to extant genera. There has been an ongoing debate over whether any pre‐Cenozoic taxa are related to

11  Biodiversity of Phasmatodea Empty egg chambers

(a)

(d)

Egg-like chamber

(b)

Operculum Embryo

(e) Anterior Operculum

(c)

2 mm

Twig

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Figure 11.7  Micro‐computed tomography scans of the ootheca of an undescribed stick insect (Euphasmatodea: Korinninae) from Vietnam. (a) Top view. (b) Lateral view. (c) Longitudinal section. (d,e) Cross sections. After Goldberg et al. 2015. Data obtained and processed by Peter Michalik, Greifswald, Germany.

the extant crown group at all (Ren 1997, Tilgner 2001, Béthoux and Nel 2002, Willmann 2003). Although numerous fossils have been reported as stick insects in the past, most of these specimens are so fragmentary, often based on wings alone (Nel et al. 2004), that their true affinities must remain questionable (Bradler and Buckley 2011). A presumed euphasmatodean stick insect was recently described from Upper Cretaceous amber (Myanmar, ~99 mya) by Engel et  al. (2016), who interpreted the extremely small individual as an adult male whose body size (8 mm) significantly falls below

that of any extant phasmatodean. Given these uncertainties, the number of genuine fossil stick insects must be even smaller than documented in textbooks (e.g., Grimaldi and Engel 2005). Some reported stick insects are possibly not phasmatodeans (Nel et  al. 2010, Bradler and Buckley 2011). Some remarkably well‐preserved and particularly informative fossils were described more recently from the Mesozoic of China (early Cretaceous, 120–130 mya), combining derived wing characters with phasmatodean apomorphies such as the male vomer, unsegmented cerci, and the female operculum

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(Nel and Delfosse 2011, Wang et al. 2014). These Chinese Cretaceous taxa are interpreted to be true stem‐phasmatodeans and are referred to as the Susumanioidea, which are probably paraphyletic with respect to the Euphasmatodea (Wang et al. 2014). The morphology of the preserved Susumanioidea sheds light on the early evolution of stick insects, which appear to have been well‐ adapted to pre‐angiospermous forests. The fossils show gymnosperm leaf mimicry and, unlike extant phasmatodeans, indicate the presence of well‐developed, long tegmina and a low degree of sexual dimorphism (Wang et al. 2014). Strikingly similar members of the Susumanioidea have now been described from the early Eocene of the United States and Canada (Archibald and Bradler 2015). Another taxon with likewise plesiomorphic characters is the Archipseudophasmatidae from Baltic amber (Zompro 2004a), which was considered to belong to the Euphasmatodea (Wedmann et  al. 2007), but might be a stem‐ phasmatodean instead. This expanded presence of stick insects largely resembling Mesozoic taxa into the early Eocene comes as a major surprise, indicating a transitional phase in which they might have persisted during the radiation of the Euphasmatodea. These ancient stick insects are not seen after the early Eocene and were replaced by modern forms in a world of changing forest communities, climates, and plant–insect interactions (Archibald and Bradler 2015).

11.4 ­Phasmatodea as Research Tools The fascinating and variable biology of stick insects has made them excellent model systems for investigating a number of evolutionary phenomena, including speciation and reproductive isolation (e.g., Nosil et al. 2012, Schwander et al. 2013, Kelly 2014, Soria‐Carrasco et  al. 2014, Myers et al. 2016), evolution of parthenogenesis and alternative reproductive strategies (e.g., Scali et  al. 2003, Morgan‐Richards et  al. 2005, Ghiselli et  al. 2007, Buckley et  al. 2008,

Schwander and Crespi 2009, Milani et al. 2010, Schneider and Elgar 2010), and more recently the evolution of cold tolerance (Dunning et al. 2014, Dennis et al. 2015). The phylogenetically relictual Californian genus Timema has yielded significant insights into speciation processes (e.g., Soria‐Carrasco et  al. 2014). Speciation studies in Timema have focused on host‐plant specificity. In particular, the species T. cristinae Vickery exhibits ecotypes that have adapted to different host plants. Geographically separate populations specializing on the same host plant showed weaker reproductive isolation than did pairs of populations specialized on different host plants (Nosil 2007). This observation suggested a role for ecological processes in driving reproductive isolation. Soria‐ Carrasco et  al. (2014) tested this and other hypotheses through genome resequencing of population pairs adapted to different host plants as well as translocated populations. They demonstrated selection acting on different regions of the genome between these replicate pairs, suggesting the development of independent evolutionary solutions or responses to a range of selective pressures. However, they also observed selection acting on the same genomic regions in repeated comparisons, showing that evolution was also acting in a consistent manner. Included in these genomic regions were genes annotated to functions such as metal ion binding, which is associated with insect diet, suggesting selection on these traits (Soria‐Carrasco et  al. 2014). These studies on Timema have had implications far beyond the Phasmatodea, as they have elegantly demonstrated that parallel processes can drive speciation. In addition to host‐plant adaptation, mate recognition has received increasing attention as an important factor in stick insect speciation (Kelly 2014; Myers et al. 2015, 2016). In a study of genital morphology in the New Zealand stick insect Clitarchus hookeri, Myers et  al. (2016) showed that mutilating the male claspers and covering the female opercular organs reduced the efficiency of coupling. In particular, modification of male claspers led to increased r­ ejection

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by females. Through Y‐maze choice experiments, Myers et  al. (2015) also showed that males of C. hookeri are probably detecting females through long‐range pheromone signals, a finding supported through the examination of antennal morphology. Myers et  al. (2015) also demonstrated a scramble competition system for mates in C. hookeri, and Kelly (2014) showed a similar mating system in the related New Zealand stick insect Micrarchus hystriculeus. These studies on the biology of mate recognition are revealing the critical phenotype systems that define stick insect species. What is now required is the unification of these studies with genomic approaches to unpick the genetic basis of these traits and the finer details of the speciation process in stick insects. Stick insects are well known for their highly variable reproductive systems. In particular, they seem to be highly prone to evolving parthenogenesis. Parthenogenesis has been studied in a wide variety of genera, including Timema (e.g., Schwander et  al. 2011 2013), Acanthoxyla (Morgan‐Richards and Trewick 2005, Buckley et  al. 2008, Myers et  al. 2013), Clitarchus (Buckley et  al. 2010, Morgan‐Richards et  al. 2010), Extatosoma (Schneider and Elgar 2010, Burke et al. 2015), Leptynia (Ghiselli et al. 2007), Clonopsis (Milani et al. 2010), and most notably in the southern European Bacillus (e.g., Scali et  al. 2003). One feature in common across many of these taxa is the role of hybridization in generating new parthenogenetic species. This was originally inferred from parthenogenetic Bacillus species that showed heterozygous allozyme profiles inherited from two parental species (reviewed by Scali et  al. 2003). Similar patterns have been shown in the Mediterranean taxa Leptynia and Clonopsis and the New Zealand Acanthoxyla (Fig. 11.1b, e.g., Buckley et  al. 2008). In Acanthoxyla, cloning and sequencing of single‐copy nuclear genes (Buckley et al. 2008) suggested hybrid origins of parthenogenetic lineages. However, the pattern is typically more complicated than a single origin of a parthenogenetic lineage following an  independent evolutionary trajectory. In

Acanthoxyla, Bacillus, Clonopsis, and Leptynia, evidence has been found of complicated patterns of reticulation whereby gene flow ­ ­repeatedly occurs from sexual relatives into parthenogenetic lineages (e.g., Scali et  al. 2003, Morgan‐Richards et al. 2005, Ghiselli et al. 2007, Buckley et al. 2008, Milani et al. 2010). The evolutionary history of many of these genera forms a complicated network rather than a bifurcating phylogeny. The application of genome‐wide markers and eventually whole‐genome sequencing is likely to reveal further complexity in the evolutionary history of these networks. Many stick‐insect species also display geographical parthenogenesis, where some populations are unisexual and others bisexual. In the New Zealand Argosarchus and Clitarchus, populations found today in the post‐glacial landscapes of the South Island tend to be asexual, whereas populations in the northern former refugia tend to be sexual (Buckley et al. 2009b, 2010; Morgan‐Richards et  al. 2010). A similar pattern is also seen in North American Timema (Law and Crespi 2002a). As in many other animal species, these stick insects are showing an association between parthenogenesis and the colonization of marginal or new habitats (e.g., Kearney 2005). The asexual populations of Argosarchus and Clitarchus do not show the elevated heterozygosity typically associated with hybrid lineages of stick insects (Scali et al. 2003, Buckley et al. 2008) and, therefore, there is probably some demographic factors favoring their geographic distribution. These stick‐insect species are therefore excellent candidates for testing general hypotheses on the evolution of geographic parthenogenesis. There has also been recent research on the evolution of adaptations that allow stick insects to thrive in a wide variety of habitats. In addition to the genomic studies that are revealing the likely key role played by host‐plant adaptation in some species of Timema (e.g., Soria‐Carrasco et  al. 2014), studies on the New Zealand stick insects are showing the genetic basis of adaptation to extreme environments. Physiological studies (Dennis et  al. 2015) have shown that

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­ifferent New Zealand alpine species have d adapted both freeze‐avoidance and freeze‐tolerance strategies for surviving in harsh alpine conditions. Furthermore, studies of gene expression demonstrated that different genes are expressed in response to cold shock in these species, and that patterns of positive selection are not the same among species (Dunning et al. 2013, 2014). Studies on the New Zealand species are showing that evolution often proceeds down different pathways even among closely related species with similar genetic backgrounds. A recent advance in understanding the biology and evolution of the Phasmatodea has been the introduction and application of genomic tools (e.g., Dunning et  al. 2014; Shelomi et  al. 2014a, 2014b; Soria‐Carrasco et al. 2014; Dennis et  al. 2015; Wu et  al. 2016). These studies are uncovering the genetic basis of key traits in the Phasmatodea and their evolution. Coupling these studies with continued research into morphology, behavior, and physiology will reveal more about their unique traits and will allow us to test general hypotheses in evolutionary biology. Genome‐scale data sets will also resolve key regions of the Phasmatodea phylogeny, for which previous data sets have yielded conflicting or inconclusive results (e.g., Whiting et  al. 2003; Bradler 2009; Buckley et al. 2009a; Bradler et al. 2014, 2015). In conclusion, it is an exciting time for phasmatodean biology. The next few years will see significant advances in knowledge and understanding.

11.5 ­Importance to Human Society Phasmatodeans are a significant component of the herbivore fauna in tropical forests (Blüthgen et al. 2006). Although the local density of these insects is often relatively low, their generally large size means that they tend to make up a large proportion of the vegetation‐damaging animal biomass. In the crown of dipterocarp trees, a few large stick insects made up an estimated 19% of the animal biomass (Ellwood and Foster 2004).

A few species are of economic importance. In Australia, Anchiale austrotessulata, Didymuria violescens, and Podacanthus wilkinsoni defoliate certain eucalyptus trees at times, which are valuable timber trees (Rentz 1996). In North America, outbreaks of Diapheromera femorata have been reported in two‐year cycles, leading to a significant decrease of black oaks in the infested area, whereas outbreaks of Graeffea crouanii in South Pacific plantations can cause severe losses of coconut palms (Bedford 1978). Phasmatodeans are considered to be among the easiest insects to raise in captivity (Rentz 1996). These impressively large and charismatic insects are often exhibited in zoos and museums. They are able to thrive on a wide variety of alternative food plants, and numerous species are kept live in culture in school classes and by a large community of enthusiastic amateur entomologists (Brock 1999, Seiler et  al. 2000). The contributions of amateur taxonomists, in particular members of the British Phasmid Study Group and the Dutch‐Belgian PHASMA society (Rentz 1996, Brock 1999), play a crucial role in describing the phasmatodean diversity and add significantly to the research outlined above.

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Bradler, S. 2009. Phylogenie der Stab‐ und Gespenstschrecken (Phasmatodea). Species, Phylogeny and Evolution 2: 3–139. Bradler, S. and T. R. Buckley. 2011. Stick insect on unsafe ground: does a fossil from the early Eocene of France really link Mesozoic taxa with the extant crown group of Phasmatodea? Systematic Entomology 36: 218–222. Bradler, S., M. F. Whiting and R. Klug. 2003. Basal diversification and the evolution of wings within stick insects. Entomologische Abhandlungen 61: 132–133. Bradler, S., J. A. Robertson and M. F. Whiting. 2014. A molecular phylogeny of Phasmatodea with emphasis on Necrosciinae, the most species‐rich subfamily of stick insects. Systematic Entomology 39: 205–222. Bradler, S., N. Cliquennois and T. R. Buckley. 2015. Single origin of Mascarene stick insects: ancient radiation on sunken islands? BMC Evolutionary Biology 15: 196. Bradley, J. C. and B. S. Galil. 1977. The taxonomic arrangement of the Phasmatodea with keys to the subfamilies and tribes. Proceedings of the Entomological Society of Washington 79: 176–208. Bragg, P. E. 2001. Phasmids of Borneo. Natural History Publications Borneo, Kota Kinabalu, Malaysia. 772 pp. Brakefield, P. M. 2006. Evo‐devo and constraints on selection. Trends in Ecology and Evolution 21: 362–368. Bresseel, J. and J. Constant. 2014. Giant Sticks from Vietnam and China, with three new taxa including the second longest insect known to date (Phasmatodea, Phasmatidae, Clitumninae, Pharnaciini). European Journal of Taxonomy 104: 1–38. Bresseel, J. and J. Constant. 2015. The new genus of stick insect Lobofemora from Vietnam, with the description of three new species (Phasmida: Phasmatidae: Clitumnini). European Journal of Taxonomy 115: 1–25. Brock, P. D. 1999. Amazing World of Stick and Leaf Insects. The Amateur Entomologists’ Society, Orpington, Kent, England. 165 pp. Brock, P. D. and J. Hasenpusch. 2007. Studies on the Australian stick insects (Phasmida),

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Tomita, S., K. Yukuhiro and N. Kômoto. 2011. The mitochondrial genome of a stick insect Extatosoma tiaratum (Phasmatodea) and the phylogeny of polyneopteran insects. Journal of Insect Biotechnology and Sericology 80: 79–88. Trewick, S. A., M. Morgan‐Richards and L. J. Collins. 2008. Are you my mother? Phylogenetic analysis reveals orphan hybrid stick insect genus is part of a monophyletic New Zealand clade. Molecular Phylogenetics and Evolution 48: 499–808. Vallotto, D., J. Bresseel, J. Constant and M. Gottardo. 2016. Morphology of the terminalia of the stick insect Dajaca napolovi from Vietnam (Insecta: Phasmatodea). Entomological Science 19: 376–382. Van de Kamp, T. and F. H. Hennemann. 2014. A tiny new species of leaf insect (Phasmatodea, Phylliidae) from New Guinea. Zootaxa 3869: 397–408. Vera, A., L. Pastenes, C. Veloso and M. A. Méndez. 2012. Phylogenetic relationships in the genus Agathemera (Insecta: Phasmatodea) inferred from the genes 16S, COI and H3. Zoological Journal of the Linnean Society 165: 63–72. Vickery V. R. 1993. Revision of Timema Scudder (Phasmatoptera: Timematodea) including three new species. Canadian Entomologist 125: 657–692. Vickery, V. and C. Sandoval. 2001. Descriptions of three new species of Timema (Phasmatoptera: Timematodea: Timematidae) and notes on three other species. Journal of Orthoptera Research 10: 53–61. Wang, M., O. Béthoux, S. Bradler, F. Jacques, Y. Cui and D. Ren. 2014. Under cover at pre‐ angiosperm times: a cloaked phasmatodean insect from the Early Cretaceous Jehol biota. PLoS ONE 9: e91290. Wedmann, S., S. Bradler and J. Rust. 2007. The first fossil leaf insect: 47 million years of specialized cryptic morphology and behavior. Proceedings of the National Academy of Sciences USA 104: 565–569.

11  Biodiversity of Phasmatodea

Whiting, M. F., S. Bradler and T. Maxwell. 2003. Loss and recovery of wings in stick insects. Nature 421: 264–267. Willig, A. 1969. Die Carotinoide und der Gallenfarbstoff der Stabheuschrecke, Carausius morosus und ihre Beteiligung an der Entstehung der Farbmodifikationen. Journal of Insect Physiology 15: 1907–1927. Willmann, R. 2003. Die phylogenetischen Beziehungen der Insecta: Offene Fragen und Probleme. Verhandlungen Westdeutscher Entomologentag 2001: 1–64. Windsor, D. M., D. W. Trapnell and G. Amat. 1996. The egg capitulum of a Neotropical walkingstick, Calynda biscuspis, induces aboveground egg dispersal by the ponerine ant, Ectatomma ruidum. Journal of Insect Behavior 9: 353–367. Wu, C., R. N. Crowhurst, V. G. Twort, A. B. Dennis, S. Liu, R. D. Newcomb, H. A. Ross and T. R. Buckley. 2016. De novo transcriptome

analysis of the common New Zealand stick insect Clitarchus hookeri (Phasmatodea) reveals genes involved in olfaction, digestion and sexual reproduction. PLoS ONE 11: e0157783. Zompro, O. 2003. Eine generische Revision der Insektenordnung Phasmatodea: Areolatae, einschließlich der Einführung einer neuen Ordnung der Insekten. Ph.D. dissertation, University of Kiel, Kiel, Germany. Zompro, O. 2004a. Revision of the genera of the Areolatae, including the status of Timema and Agathemera (Insecta, Phasmatodea). Verhandlungen des Naturwissenschaftlichen Vereins in Hamburg (NF) 37: 1–327. Zompro, O. 2004b. A key to the stick‐insect genera of the ‘Anareolatae’ of the New World, with descriptions of several new taxa (Insecta: Phasmatodea). Studies on Neotropical Fauna and Environment 39: 133–144.

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12 Biodiversity of Dermaptera Fabian Haas Leipzig, Germany

The Dermaptera, or earwigs, are a small group of medium‐sized insects (typically 10–15 mm long), with more than 1900 described, living species (Table 12.1), although only a few are known to the general public. Forficula auricularia, for example, is a synanthropic species that appears in agricul­ tural and domestic environments. Most other species inhabit a wide range of natural habitats. The characteristic habitus of earwigs evolved early, and fossils are easily identified as Dermaptera, based on knowledge of extant insects. The pres­ ence of ocelli, five tarsomeres, and annulated cerci are plesiomorphic character states found only in fossil specimens or, for annulated cerci, in a few extant taxa. Dermaptera have well‐developed hindwings, each of which folds into a wing pack­ age. They also have several special characters, such as densely folded wings, forceps‐like cerci, and maternal care of eggs and first‐instar nymphs. These characteristics are found in all species, although they show some variability within the group. Some species are ovoviviparous, laying eggs from which nymphs immediately hatch, or they are or viviparous, actually giving birth to nymphs.

12.1 ­Epizoic Dermaptera The most significant departures from typical earwigs are the Arixeniidae and Hemimeridae, two distinct and unrelated epizoic groups (i.e.,

living non‐parasitically on other animals). Both families convergently evolved a close relation­ ship with mammals. The exact nature of this relationship previously was unclear, and so they were considered parasites (e.g., Popham 1984), implying a negative effect on the host animal. A closer examination, however, could find no neg­ ative or positive effects on the hosts; thus, these dermapterans should be considered commen­ salistic or epizoic. The Hemimeridae, first described by Walker (1871), occur in Africa south of the Sahara on hamster rats (Beamys and Cricetomys) and are reported from many locations across their range. They have long, oval, flattened bodies, short antennae, thin filiform cerci, and a lack of dense setation. The annulation (segmentation) of the cerci has been discussed as a possible ple­ siomorphic character (Popham 1984), although this could not be confirmed. The legs are short, compared with those of free‐living Dermaptera, but are highly adapted for moving in the dense fur of hamster rats (Haas and Gorb 2004). Each of the three tarsomeres has pads with dense setation that assists attachment to the fur. This characteristic is of special importance because individuals cannot survive for an extended period without a host. The gut contents indicate that members of the Hemimeridae feed on skin flakes and possibly fungus growing on the skin of hamster rats or around the nests. Ashford (1970) kept live hamster rats with Hemimerus

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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Table 12.1  Families and described, valid species of living Dermaptera. Family

No. of genera

No. of described species

Figures

Karschiellidae

2

12

—

Diplatyidae

9

143

12.1f

Pygidicranidae

18

182

12.1c, 12.2e

Anisolabididae

37

393

12.1e, 12.2g

Labiduridae

7

73

12.1b, 12.2a,b,d,f,h

Apachyidae

2

15

—

40

499

—

Forficulidae

67

ca. 500

Chelisochidae

17

95

Eudermaptera Spongiphoridae

12.1g,h, 12.2c, 12.3 12.1d

Arixeniidae

2

5

12.1a

Hemimeridae

2

12

—

203

ca. 1,930

—

Total

The Arixeniidae and Hemimeridae have been regarded as orders and suborders, but they now are considered to belong within the Forficulina, which constitute the “typical” earwigs. Therefore, they are included in this table as families (Kocarek et al. 2013, Naegle et al. 2016).

for about four and a half months and observed many aspects of their biology. The highly specialized life history and strong dependency on hamster rats led to modifica­ tions in the reproductive strategies. The Hemimeridae are viviparous. Rather than laying eggs in a nest like other Dermaptera, they give birth to fully developed and mobile first instars. The structure of the ovaries was examined more than 100 years ago (Heymons 1912) and indi­ cated a close relationship with the higher Dermaptera (Jarvis et  al. 2005). This relation­ ship was confirmed molecularly by Kocarek (2013) and Naegle et  al. (2016), who showed that the Hemimeridae are nested within the Eudermaptera. Chromosomal data also are available (White 1971). An overall description of hemimerids has been given by Jordan (1909), and a detailed description of the female ­abdomen has been provided by Klass (2001). Rehn and Rehn (1935, 1937) also gave useful ­descriptions. A recent glimpse of hemimerid

­istributions and faunistics was provided by d Cook and Richardson (2010). The second, strongly divergent taxon in the Dermaptera is the Arixeniidae (Fig. 12.1a), at one time considered a separate order or subor­ der of Dermaptera. Jordan (1909a, 1909b) first described the taxon and gave a detailed morpho­ logical description. Nakata and Maa (1974) pro­ vided an excellent review. The Arixeniidae live in association with bats in caves in Malaysia (e.g., the Malay Peninsula and Sarawak), Indonesia (Java and Sumatra), and the Philippines. Their structure is highly derived, with long legs that allow them to move around rapidly. The anten­ nae are long, the tegmina (i.e., forewings) and hindwings are absent, and the cerci are short. The body is densely pubescent. Members of the Arixeniidae have a significant risk of being separated from their hosts, often dropping to the ground and being forced to find a new bat for food and a hiding place. Therefore, rapid locomotion is an advantage, as it enables

12  Biodiversity of Dermaptera (a)

(e)

(f)

(b)

(g)

(c) (h)

(d)

Figure 12.1  (a) Arixenia esau (Arixeniidae) from Borneo, with a unique life history and long, slender legs. (b) Female of Labidura riparia (Labiduridae) in defensive posture in Belgium. Labidura riparia prefers sandy underground habitats such as beaches and riverbanks. (c) A representative of Echinosoma sp. (Pygidicranidae) with uniquely strong, short bristles (i.e., modified setae). (d) Schizoproreus volcanus (Chelisochidae). The Chelisochidae form a small taxon with a preference for warm, humid tropics. The exception is Chelisoches morio, a tramp species of worldwide distribution. (e) Anisolabis maritima (Anisolabididae), a widespread, generalized earwig on which many behavioral and physiological studies have been conducted. Many of the Anisolabididae resemble this species, and identifications require examination of genitalia. (f ) Nymph of Diplatyidae from Brunei, with long, annulated cerci, a plesiomorphic character state in the Dermaptera. (g) Forficula senegalensis (Forficulidae) killed by an unknown fungus in Kenya. (h) Forficula auricularia feeding on grapes after they had been opened by wasps in Germany. (a,c–f ) Photographs by Petr Kocarek, University of Ostrava. (b,g,h) Photographs by Fabian Haas. (See color plate section for the color representation of this figure.)

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them to quickly cover the several meters from cave floor to ceiling. Members of the Arixeniidae feed on glandular secretions and skin flakes, and shelter in the skin folds of the bats. They are found on cave floors and walls, suggesting a weaker link to their host. Survival without the host is possible for a considerable time. Their occurrence in several widely separated caves suggests that they at least occasionally cling to their hosts during flight. The females are viviparous, and the ovarial structure resembles that of higher Dermaptera (Jordan 1909, Tworzydlo et  al. 2013). First instars, rather than eggs, are deposited by the female. The chromosomes have been examined (White 1972). Like other highly specialized taxa (e.g., para­ sites), the high degree of morphological differen­ tiation makes taxonomic assignment difficult. Thus, the Arixeniidae and Hemimeridae have conveniently been regarded as suborders (Arixeniina and Hemimerina). However, this clas­ sification expresses a probable close relationship to the suborder Forficulina, but does not explain their evolution (i.e., the last common ancestor) or which taxon is their sister group. Molecular data provided a new set of tools for addressing these issues, and Jarvis et  al. (2005), Kocarek et  al. (2013), and Naegle et al. (2016) suggested that the Arixeniina and Hemimerina are, indeed, derived Eudermaptera (Spongiphoridae, Chelisochidae, and Forficulidae), with Arixeniina and Chelisochidae as sister groups and Hemimerina and Forficulidae as sister groups. Accordingly, the two taxa are considered families and named with  the appropriate suffix: Arixeniidae and Hemimeridae.

12.2 ­Structure and Function The structure of Dermaptera is fairly uniform (Fig. 12.1b–e). Earwigs have long, filiform antennae; chewing, prognathous mouthparts; compound eyes (with the exception of a few cave‐dwelling species); and no ocelli. The head is well articulated and moveably attached to the

prothorax, which in turn is well articulated with the fused mesothorax and metathorax. Each thoracic segment bears a pair of walking legs that insert laterally; the arrangement is an ideal adaptation for pushing into crevices or between leaves. The legs show no modifications for dig­ ging, jumping, or catching prey. The tarsus always consists of three tarsomeres. The second tarsomere is modified as a straight, narrow attachment pad in the Chelisochidae, or as a heart‐shaped attachment pad in the Forficulidae. In Hemimerus, the tarsomeres are adapted to clinging to and moving in the hair of their hosts (Haas and Gorb 2004). In a few taxa (Tagalina and Geracinae), an arolium is present. Two claws are present. The meso‐ and metathorax together form the flight apparatus. The tegmina‐bearing meso­ thorax is rather short and narrow, whereas the metathorax, which bears the wings (i.e., hind­ wings), is larger, indicating that the major flight muscles are in the metathorax. The metathorax is strongly slanted so that the pleural sulcus is almost horizontal, with the wing articulation in the anterior corner of the thorax and the coxal articulation points located posteriorly. The abdomen follows the thoracic slanting, with each tergum of half‐segment width ante­ rior to the sternum of the same segment, result­ ing in a zig‐zag pattern of sclerite margins in lateral view. The stigmata (i.e., openings of the tracheal system) are hidden. The abdominal ter­ gites can bear folds on each side, beneath which defensive glands are situated. The sternites are not specialized, with the exception of the ulti­ mate sternites, which have protrusions in the males of some species. In males and nymphs, 10 tergites are visible. In females, tergites VIII and IX are narrow and hidden under tergite VII. The cerci are present as non‐annulated for­ ceps in adults. In the nymphs of Karschiellidae and Diplatyidae (Fig. 12.1f ), the cerci are annu­ lated and resemble filiform antennae (Haas et al. 2012). If the shape of the cerci is sexually dimor­ phic, then the male – similar to the situation for mandibles and horns in the Lucanidae and Scarabaeidae (Coleoptera) (Emlen and Nijout

12  Biodiversity of Dermaptera

2000, Emlen 2008) – has the stronger and more elaborate shape. Cerci are awl‐shaped or with strong teeth or flanges on the dorsal, ventral, or medial sides, consequently appearing branched. They can be straight or undulated and symmet­ rical or asymmetrical, and can be about as short as one abdominal tergite or about as long as the abdomen. In females and nymphs, the cerci are short and straight, with few, if any, teeth. Cerci are used in many behavioral contexts, including defense, predation, and male–male interactions (Briceno and Eberhard 1995). During mating, the male twists the highly mobile abdomen and supports the straight female abdo­ men from below to assist insertion of its genita­ lia (Fig. 12.2a,b) (Walker and Fell 2001).

12.3 ­Locomotion The typical form of locomotion in Dermaptera is walking. Active flight is observed in some species, such as those that come to light traps, but might be the exception. Locomotion in ear­ wigs is an adaptation for moving on fairly flat, firm surfaces. The strong slant of the pleural sulcus posterior to the coxal articulations ena­ bles the hind legs to push horizontally and thereby move the body in narrow spaces between leaves and bark, under stones, and in the soil. These are the typical hiding places dur­ ing the daytime, and predatory species find their prey in these locations. Although up to 40% of earwigs have at least partly reduced hindwings (Wagner and Liebherr 1992), flight is used to travel greater distances, such as between dung heaps in the case of Labia minor. Flight, however, is not used as an escape mechanism because unfolding of the wings needs considerable preparation, and is accom­ plished either with the cerci or by flapping the wings a number of times (Haas et al. 2012). The hindwings (Fig. 12.2c) are highly derived structures in the Dermaptera, unique among all insects in several characters (Haas and Kukalova‐ Peck 2001). Each consists of a narrow spar at the anterior costal margin and a large fan in the

­ osterior area. The fanwise folded area is crossed p by two transverse folds. In all other insects, the fan is never folded transversely beyond its fan­ wise folding. If there is a transverse folding in the wing in other taxa, it never includes the fan. In addition to this transverse and fanwise folding, the dermapteran hindwing folds along a longitu­ dinal crease, so that in total, four major folding lines are present. The venation is highly derived, with the major braches of the anal vein radiating from the center of the wing rather than from its base. An extensive study of the venation shows some minimal variation, while the relationship between veins and folding lines is invariable. The hindwing is folded into a wing package with about one‐tenth of the surface area of the wing. The mechanics of folding and unfolding were studied by Haas et al. (2000), demonstrating the presence of resilin, an elastic protein with rubber‐ like properties in insect cuticle, at highly stressed points in the wing. The cerci are used to unfold the wing package against the intrinsic elasticity. Two stiffening mechanisms in the hindwing lock it in the flat, unfolded position and make it stiff enough for flight. After flight, the stiffening mechanisms are released, and the hindwing folds back into its folded position as a wing package. The unfolding of the wing was considered to depend on interaction with the cerci; however, Haas et al. (2012) showed earwigs flapping with the wing package, which then unfolds and stiffens after 2–4 strokes. The earwigs were observed in a natural setting (Fig. 12.3).

12.4 ­Distribution Dermaptera as a whole are associated with warm, somewhat humid climates (Fig. 12.4). Accordingly, the greatest diversity is found in the humid trop­ ics, such as Brazil (147 species; Haas 2012), Nicaragua (42; Maes and Haas 2006), and Australia (88), whereas the temperate regions (e.g., Germany, 9; Korea, 22; United States, 28) have only a limited number of species (Haas 2015). If a species is found in extreme climates, such as F. auricularia in Iceland (Tuxen 1938), it is

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Figure 12.2  (a,b) Labidura riparia (Labiduridae) in copulation. (c) Right hindwing of Allodahlia scabriuscula (Forficulidae). (d) Labidura herculeana, the only extinct species of earwig. Specimen in the Copenhagen Natural History Museum (Zoological Museum). Total length ca. 80 mm. (e) Freshly molted adult and nymphal exuviae of Cranopygia marmoricrura (Pygidicranidae) from Borneo. The fine white filaments in the exuviae are the inner lining of large tracheae. Note the well‐pigmented compound eyes, their large size indicating a predaceous life history. (f ) Male of L. riparia in defensive posture in Belgium. (g) Ctenisolabis sp. from Borneo, an exceptionally small member of the Anisolabididae. (h) Body types of the Labiduridae, other than that of the cosmopolitan and well‐studied L. riparia, include the smaller body of Nala tenuicornis. (a,b,c,d,f ) Photographs by Fabian Haas. (e,g,h) Photographs by Petr Kocarek, University of Ostrava.

Figure 12.3  Wing unfolding of a male of Timomenus lugens (Forficulidae), showing continuous frames (333.33 ms) of a movie clip with frame rate of 30 fps. Photo by Arlo Pelegrin (Haas et al. 2012).

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Figure 12.4  Species numbers of earwigs per country.

restricted to an appropriate microclimate, such as a human settlement or an oasis in a desert (Weber 1954). Regions with areas of higher humidity, such as the mountains in Tanzania or country‐wide in Uganda, have a significantly higher number of species than do drier areas such as Kenya. Tanzania has many isolated mountains, and endemic spe­ cies occur on these mountains, further increasing the biodiversity (Haas and Klass 2003). With few exceptions, Dermaptera are not synanthropic, and they have a weak propensity to distribute and extend their range, possibly related to a lack of flight activity in many species. The exception to this rule is F. auricularia, which has attained cosmopolitan distribution, and in some regions it is considered an invasive alien species (e.g., Canada and the United States, Capinera 2011; Mexico, Pavón‐Gozalo et  al. 2011). This

species lives near or in human settlements and thus is easily transported. Furthermore, a num­ ber of alien species are found in European green­ houses (Kocarek et al. 2015, Matzke and Kocarek 2015). Dermaptera also occur on remote islands such as Hawaii (Brindle 1980) and the Galapagos Archipelago (Peck 2001). A few species colonize caves throughout the world, and the eyes are reduced or lost in these species (Assam and Burma, Chopard 1924; Cuba, Brindle and Decu 1977; Canary Islands, Ashmole et  al. 1992; Hawaii, Brindle 1980). Labidura riparia (Fig. 12.1b, Fig. 12.2a,b,f) has attained cosmopolitan distribution. It prefers sandy habitats, which can be found on river banks and seashores, making the species prone to dispersal with drifting materials. It has reached many islands (and inland areas) (Hawaii, Brindle 1980, Nishida 1994; Papua New Guinea, Edwards

12  Biodiversity of Dermaptera

and Thornton 2001). The now‐extinct Labidura herculeana on St Helena in the Atlantic Ocean (Brindle 1970) probably evolved from L. riparia. Labia minor, a species living in dung heaps from horses and cows, possibly extended its range from Asia around the globe via horses and cows (Picker and Griffiths 2011). Labia minor is a predaceous species and finds an ample supply of maggots and other insects in dung heaps.

12.5 ­Development and Reproduction Development in Dermaptera is hemimetabolous, with an egg stage, 4–6 nymphal instars, and the adult stage. Two species, Marava arachidis and Chaetospania borneensis, independently evolved (ovo)viviparity (Kocarek 2009). Viviparity in the two epizoic taxa (Arixenia and Hemimerus) ensures that the host is immediately available to the first‐instar offspring. After copulation (Fig. 12.2a,b), females build a nesting site under stones, in crevices, or in exist­ ing tunnels. Up to 50 eggs 1.0–2.5 mm in diam­ eter are laid, and after 8–20 days, the first instars hatch using an egg‐tooth to penetrate the egg shell. In some species of Tagalina and Diplatys, the eggs are glued to a surface with an egg cup (Matzke and Klass 2005, Shimizu and Machida 2011). The female regularly cleans the eggs, pre­ venting fungal growth. It actively defends and reassembles the clutch after disturbances, and relocates the nest to a new site if conditions are unsuitable. Nymphs disperse before or after their first molt. They molt 4–6 times within 2–8 months to become adults. The gender and wing buds are vis­ ible from the third instar onward, and with the exception of the Diplatyidae and Karschiellidae, the cerci are short and straight, resembling those of the adult female. In nymphs of the Diplatyidae (Fig. 12.1f) and Karschiellidae, the cerci are annu­ lated (Haas et al. 2012), and all but the basal annu­ lus is lost during the final molt. The adult lives about 6–7 months, and so the life span is about one year. Figure 12.2e shows a freshly molted adult

of Cranopygia marmoricrura (Pygidicranidae) with its exuviae. The life cycle is influenced by sea­ sonality (dry–wet or cold–warm).

12.6 ­Behavior Maternal care and mating have attracted most of the research on dermapteran behavior, and little is known of the remaining behavioral repertoire. Dermaptera show aggregation behavior, whereby conspecific animals gather together. This behav­ ior is mediated by an aggregation pheromone (Sauphanor 1992, Hehar et al. 2008). 12.6.1  Mating Behavior and Maternal Care

Kamimura (2014) and Kamimura and Lee (2014a, 2014b) examined courtship behavior and the role of the variable number and direc­ tion of the penes. During courtship, the male rotates its highly flexible abdomen and pushes its cerci underneath the female abdomen to gain access to the female genitalia (Fig. 12.2a,b). However, it rarely holds or grasps the female with its forceps; only males of Pseudomarava prominens are known to use their forceps to grasp the female (Briceño and Eberhard 1995, Kamimura 2014). Actual copulation in earwigs can take several hours and can be repeated before the male and female separate. Suzuki et al. (2005) examined maternal care in Anechura harmandi and found that matriphagy increases the survival rates of nymphs. However, matriphagy and semelparity are the exception in Dermaptera, as many females can have more than one brood if climatic conditions allow. Females of Anisolabis maritima provision food for their off­ spring and increase their survival rate (Suzuki 2010, 2011), and females of F. auricularia regurgi­ tate food to the nymphs (Stärkle and Kölliker 2008). Yet, cannibalistic behavior exists (Dobler and Kölliker 2011). Maternal care increases ­survival of the offspring and is not restricted to cleaning the eggs (Boos et al. 2014). Herter (e.g., 1967)  conducted extensive ­observations on the

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r­eproductive behavior in the 1960s and 1970s, and Kölliker and Vancassel (2007) studied familial relationships. Wong et  al. (2013) contributed a review of the evolution of parental care in insects, and Butnariu et al. (2013) studied the behavior of Doru lineare. 12.6.2 Defense

The defensive behavior of dermapterans includes threatening the attacker with raised forceps and pinching it (Fig. 12.2f ). In rare cases, earwigs can penetrate human skin, and a small droplet of blood may be the trophy (Bishopp 1961). However, defensive behavior is more effective against insects. Attack behavior is strengthened by the presence of abdominal defensive glands in some species, such as L. riparia, which spray their contents at predators. The glands are situ­ ated in the folds on the abdominal tergites of a variable number of segments (tergites III to V). The substance has antimicrobial activity, pre­ venting infection during aggregation in the win­ ter (Gasch et al. 2013). 12.6.3 Feeding

Occasional feeding observations and gut‐con­ tent examinations have been conducted on the Dermaptera. Forficula auricularia is omnivo­ rous, feeding on soft plant parts and insects, and L. riparia is mainly carnivorous on arthro­ pods. Other species feed on a mixture of insects, plant materials, and spores and parts of fungi. Members of the Karschiellidae are reported, based on one review, to feed on ants. The influ­ ence of feeding on the development and ovipo­ sition of F. auricularia in North America has been examined by Berleur et  al. (2001). In Sudan, Forficula senegalensis has been studied in some detail, revealing that it feeds mainly on pollen and insects, but not on green leaves; the species, therefore, can no longer be considered a pest of millet (Boukary et al. 1997). The effects of diet on development also have been investi­ gated (Boukary et al. 1998). Original taxonomic descriptions occasionally note feeding habits,

and gut‐content examinations also give some insights (e.g., Haas 1995).

12.7 ­Parasitism and Symbiosis In insects, the gut and the fat body are common sites for symbionts and parasites, but there are few reports for earwigs. However, apicomplexan parasites (Gregarina spp.) were found in Vostox brunneipenis (Clopton 2008). The genus of gre­ garines was erected by Dufour in 1828 after find­ ing them in specimens of F. auricularia. Tachnid flies are parasitoids of earwigs. Ocytata pallipes and Triarthria setipennis, for example, parasitize F. auricularia and have been used for biological control of this earwig (Kuhlmann 1993, 1994). A fairly unique form of specialized predation on earwigs is that of the formicid Leptogenys sp., which feeds exclusively on the earwigs Gonolabis electa and Paralabis sp. (Steghaus‐Kovac and Maschwitz 1993). Otherwise, many groups of animals, such as amphibians, birds, reptiles, and spiders, prey on earwigs, although they do not specialize on them. Fungi occasionally infest earwigs (Fig. 12.1g).

12.8 ­Fossils and Research History Earwigs typically have low population densities and are rare in the fossil record. Only about 100 fossil specimens are known to date. They are recorded from the Triassic Period (ca. 252–201 million years ago (mya)) (Grimaldi and Engel 2005), and as amber fossils from the Cretaceous Period (ca. 145–66 mya) (Engel and Grimaldi 2014). Their stem‐group, Protelytroptera, achieved significant biodiversity as early as the Permian Period (298–252 mya), and some rep­ resentatives indicate the evolution of the highly complex folding pattern in the hindwings of earwigs (Haas and Kukalova‐Peck 2001).

12  Biodiversity of Dermaptera

Early Dermaptera show the typical habitus: long, slender bodies, with a prognathous head and flexible abdomen. The cerci in early fossil adults and nymphs are long and filiform. Each tarsus has five, rather than three, tarsomeres, and three ocelli are present in these early ear­ wigs. Fossil dermapterans from younger strata are indistinguishable from recent Dermaptera. The Dermaptera have received little but con­ stant attention in the entomological commu­ nity. They cannot be considered an “orphan group” in which no research was conducted over decades. The most notable names in der­ mapteran research are Malcolm Burr (1878– 1954), Walter D. Hincks (1906–1961), and Alan Brindle (1915–2001), who wrote the still‐excel­ lent identification keys to the group. Both Hincks and Brindle worked at Manchester Museum, which has the largest collection of Dermaptera in the world (Miles 2015). Other important contributions came from Borelli, Günther, de Bormans, and Zacher. The number of collected specimens per species is generally low; a species can be known from as few as 10 specimens. Nonetheless, a good, systematic framework now exists for new discoveries and biodiversity research. At present, there is con­ siderable interest in the Dermaptera, covering almost all aspects, including structure, function, and behavior.

12.9 ­Overview of Taxa The phylogeny of the Dermaptera has been stud­ ied several times over the past 20 years. All studies have some weaknesses, such as a focus on one character system (for instance, wings, thus exclud­ ing the many non‐winged species, e.g., in Haas and Kukalova‐Peck 2001). In other cases, the sam­ pling was rather incomplete, with the Arixeniidae and Hemimeridae missing from earlier molecular data sets. Colgan et  al. (2003) were the first to apply molecular techniques, followed by Jarvis et  al. (2005). Two of the most recent molecular studies are those of Kocarek et  al. (2013) and Naegle et al. (2016), which d ­ emonstrated that the

Arixeniidae and Hemimeridae nest within the “general” Dermaptera. Another significant phylogenetic obstacle is the uncertainty of monophyly of the so‐called “fami­ lies.” For Anisolabididae, Labiduridae, and Pygidicranidae, no strong autapomorphic charac­ ters could be established suggesting monophyly. This condition applies to four out of nine studied taxa (Haas and Kukalova‐Peck 2001). Allostethus indicum was removed from its traditional family, the Labiduridae. The situation is particularly vague for the Anisolabididae and Spongiphoridae. The few groups with convincing autapomorphies are the Eudermaptera, Apachyidae, Chelisoch­ idae, Forficulidae, Arixeniidae (Arixenia), and Hemi­meridae (Hemimerus). Dermapteran phylogeny is, thus, somewhat unstable, and the distinction between lower and higher Dermaptera is arbitrary, although based on similarities in lifestyle and distribution. 12.9.1  Lower Dermaptera

The Karschiellidae, Diplatyidae, and Pygidi­ cranidae are often considered Lower Derma­ ptera.  They are usually large and strongly built insects, and in the first two groups, the nymphal cerci are annulated. Little is known about their reproduction. Diplatys flavicollis and Tagalina papua use egg cups to glue their eggs to the sub­ stratum (Matzke and Klass 2005, Shimizu and Machida 2011). Many species seem to be preda­ ceous and carnivorous. They are distributed mostly in the warm tropics, with the Karschiellidae occurring only in Africa south of the Sahara, although a few taxa occur in the Nearctic and Palearctic Regions (e.g., Diplatys in Egypt, Challia on the Korean peninsula, and Pyragropsis in the southern United States). With the exception of the highly modified male genitalia of the Karschiellidae, the Lower Dermaptera have two penes pointing posteriorly. The number of genital structures (i.e., the penis and sclerotized distal part of the ejaculatory duct, called the virga) and their orientation are extensively used in species identification (e.g., in the classic work on African earwigs by Brindle 1973, 1978). The Karschiellidae

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and Diplatyidae were often grouped as subfami­ lies in the “Pygidicranidae.” 12.9.2  Higher Dermaptera

The Anisolabididae (Fig. 12.2g,h), Apachyidae, Labiduridae (Fig. 12.2h), and Eudermaptera are often considered Higher Dermaptera, but their monophyletic status is not firmly established. Many of these species occur in temperate regions of the world, such as Europe and North America. They have two penes, with one directed anteriorly and one posteriorly, or have one penis completely reduced. Labidura riparia is commonly found on beaches and in sandy ground throughout the world, as are Euborellia spp. The Apachyidae are limited to the Old World tropics and are extremely flat, with an extension of the tenth abdominal tergite. The Eudermaptera consist of the Chelisochidae, Spongiphoridae (both occur mainly in warm cli­ mates), and Forficulidae. Eudermaptera consti­ tute a monophyletic group, with many derived character states, and can be considered the most evolved taxon in the Dermaptera. Nonetheless, they do not show significant morphological spe­ cializations. The Chelisochidae are recognized by a long, slender extension of the second tar­ somere ventral to the third tarsomere, whereas in all Forficulidae (nymphs and adults), this exten­ sion is heart shaped.

12.10 ­Societal and Scientific Importance 12.10.1  Plant Pests, Biological Control Agents, and General Nuisances

A few species of Dermaptera are known as pest species, mainly F. auricularia in greenhouses and on flower farms, where they feed on inflo­ rescences and reduce the market value of the flowers. However, their mandibular strength is fairly weak, being incapable of penetrating the skin of grapes; instead they feed on grapes opened by wasps (Fig. 12.1h). Forficula auricularia

occasionally is a nuisance when it hides in large numbers in homes and garden greenhouses (Capinera 2011). Earwigs do not pose a health or hygienic risk. In orchards and gardens, F. auricularia is welcomed because it feeds on aphids and other pest insects (Suckling et al. 2006). Pests and natural enemies have the capability to reproduce quickly in large numbers, using a natu­ ral resource in the short time that it is available. Earwigs lack this capability, with maximum batch sizes of 50 eggs and two batches each year. Trials to use them as natural enemies have failed because of their wide range of feeding habits; none of the spe­ cies are highly specialized on particular organ­ isms. However, they can be important predators on pests in a natural context, such as F. auricularia on aphids (Suckling et al. 2006) and A. maritima on the red palm weevil Rhyncophorus ferrugineus in Saudi Arabia (Al‐Dosary 2009). 12.10.2  Medical, Veterinary, and Forensic Importance

Folk legend claims that earwigs crawl into human ears, penetrate the tympanum, and lay eggs in the brain. This reputation is only a myth and, although there are a few reports of earwigs being found in human ears, this habitat does not serve as a breeding site (Berenbaum 2007). Earwigs simply prefer warm, humid, narrow spaces. The forceps of earwigs are strong enough to painfully pinch the finger of an insect collector, sometimes drawing a drop of blood. No veterinary issues have been reported. However, mass occurrences of earwigs in poul­ try operations have been reported, where they feed on fly larvae and other insects in the bird feces (Berti et al. 1996). Cadavers constitute a rich and easily accessi­ ble source of nutrients, and they attract insects in a specific sequence, allowing determination of the time of death. Labidura riparia and other species have been found under cadavers (Wolff et al. 2001 and personal observations in Magadi, Kenya), but they do not seem to be strictly related to the cadaver; rather, they might be sim­ ilar to birds by profiting from the large numbers

12  Biodiversity of Dermaptera

of flies on which they will feed. No information on the time of death can be inferred from their presence (Segura et al. 2009). 12.10.3  Invasive Alien Species

Invasive alien species pose a major problem for ecosystems. Owing to their limited reproduc­ tion and distributional capabilities, dermapteran species do not pose such problems. However, Chelisoches morio and F. auricularia have been transported and established around the globe. Mass reproduction of F. auricularia has been reported from Canada and the United States, but no reports indicate that they would domi­ nate the local ecosystem. Pavón‐Gozalo et  al. (2011) reported the invasion of Mexico by F. auricularia. Some crop damage by this species has been reported (Capinera 2011). Another tramp species is L. minor, with conge­ neric species in Asia. Possibly a Palearctic species by origin, it is found in association with dung heaps. Thus, this species might have benefited from the spread of agriculture and domestication of animals over the past few thousand years. Despite its ability to live in a wide range of habitats, L. riparia is negatively influenced by the invasive fire ant Solenopsis invicta in the southern United States (Calixto et  al. 2006). Thus, earwigs can be subject to the detrimental effects of invasive alien species. 12.10.4  Pollination and Other Ecological Services

Accidental pollination by earwigs might occur, but low densities and low specificity of habitat and life history contradict at least some essen­ tial ecosystem services. Pseudoscorpions and mites occasionally are found on earwigs, using them as a means of transport (phoresy) (Seeman 2007, Chmielewski 2009). 12.10.5  Research Tools

A significant body of research exists on earwig physiology, although earwigs never achieved

the status of Rhodnius for hormone research, let alone Drosophila for genetic research. Selected contributions can be mentioned. In general, only about five species have been examined in detail. Dermapteran neurosecretory cells were examined by Awashti (1976). Gäde (1999) found adipokinetic peptide in Dermaptera. Sayah (2002) examined the corpus allatum in adults of L. riparia, and Sayah and Lavendure (2001) localized a bombyxin‐like peptide in the brain. Caussanel (1991) examined the ecophysiology of behavioral and social phenomena in L. riparia. Sauphanor (1992) found an aggregation pheromone in the European earwig, and exam­ ined pesticide side effects on the same species (Sauphanor et  al. 1993). Rankin et  al. (1997) tested the effects of diet and mating status on the hormonal system of Euborellia annulipes, and Rankin et al. (2004) examined the effects of the male nerve‐cord status on female mating behavior. Ozeki (1958, 1964, 1977) studied the regeneration of antennae, hormonal aspects of molting, and ovarian development. Leader and Bedford (1972) scrutinized the hemolymph of A. maritima. Bertella et al. (1995) examined the cuticulogenesis and embryonic development of L. riparia, and Caussanel et al. (1986) examined oviposition control. Vancassel and Quris (1994) studied diapause as an indicator of reproductive cohorts. Resistance of eggs to low and high tem­ peratures was the focus of Chauvin et al. (1991). Zinsmeister (1973) studied RNA and protein synthesis in earwig ovaries. Karyotypes of the Dermaptera have been reviewed by Mittal and Sakai (1996), and White (e.g., 1971, 1972) examined the karyotypes of many species. Spermatogenesis is known in five species (Morgan 1928). Wan et al. (2012) pub­ lished the first complete mitochondrial genome of an earwig species (Challia fletcheri). The connection of environment and genetic deter­ minants was examined by Briceño and Eberhard (1987) and Tomkins (1999). Adaptation of the eyes in Labidura truncata was the focus of McLean et  al. (1977). Marais et  al. (2005) discussed gas exchange in the

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Dermaptera in a wider phylogenetic context. Pass (1988) took an interest in the circulatory system of Chelidurella acanthopygia, F. auricularia, and L. riparia. Brousse‐Gary (1983) found glandular systems on the tibia of L. riparia, and described sensilla in the vagina of the same spe­ cies (Brousse‐Gary 1985). Peters and Latka (1986) showed the presence of chitin in the peri­ trophic matrix of F. auricularia. For research questions on the evolution of parental care, Roulin et  al. (2014) stated that: “[t]he European earwig (Forficula auricularia) is an established system for studies of sexual selection, social interactions and the evolution of parental care.”

Probably, the “computer bug” (i.e., referring to an insect) comes closest to this in English. In classic English, “earwig” (apart from the animal) means a flatterer or someone who whispers into your ear and insinuates things. “To earwig” is a syno­ nym for “to eavesdrop.”

12.10.6 Conservation – Vanishing Species

­References

Labidura herculeana of St Helena in the South Atlantic Ocean is the only documented case of an extinct, modern earwig (Fig. 12.2d). The spe­ cies resembles a large version of L. riparia, with an overall length of 80 mm. It was distributed exclusively in a limited habitat on St Helena. Rats and habitat destruction caused the species to disappear in the late 1960s or 1970s, and it is now officially rated extinct (IUCN 2016).

Al‐Dosary, M. M. 2009. Morphological characterization of the antennal sensilla of the earwig Anisolabis maritima (Dermaptera: Carcinophoridae) with reference to their probable functions. Saudi Journal of Biological Sciences 16: 17–22. Ashford, R. W. 1970. Observations on the biology of Hemimerus talpoides (Insecta: Dermaptera). Journal of Zoology 162: 413–418. Ashmole, N. P., P. Oromi, M. J. Ashmole and J. L. Martin. 1992. Faunal succession on lava flows and in caves of the Canary Islands. Biological Journal of the Linnean Society 46: 207–234. Ashworth, A. C. 1973. The climatic significance of a Late Quaternary insect fauna from Rodbaston Hall, Staffordshire, England. Entomologica Scandinavica 4: 191–205. Awashti, V. B. 1976. Neurosecretory cells and aorta as neurohaemal organ in the earwig, Euborellia annulipes Lucas (Dermaptera: Labiduridae). International Journal of Insect Morphology and Embryology 5: 253–260. Berenbaum, M. 2007. Lend me your earwigs. American Entomologist 53: 196–197. Berleur, G., J. Gingras and J. C. Tourneur. 2001. Influence of diet on development and oviposition of Forficula auricularia

12.10.7  Cultural Legacy

Colloquial names for the Dermaptera exist in many languages and are either derived from “ear” or from the conspicuous forceps (Berenbaum 2007). “Ohrwurm” in German might also refer to a catchy tune that refuses to leave one’s mind. In current British English, “earworm” means the same (R. J. Wootton and C. Lyal personal com­ munication). The word “Wurm” in German refers to undetermined causes that spoil some­ thing; for example, a toothache is due to a “tooth worm,” and a stomach ache is due to a “stomach worm.” Although this usage has mostly been lost with the realization of the actual causes, the gen­ eral expression is still in use: for example, “Da ist der Wurm drin” indicates that something just does not work, but you do not know why.

­Acknowledgments I thank Petr Kocarek (University of Ostrava) for permission to use his excellent images of Dermaptera and Arlo Pelegrin for taking the photos of dermapteran wing unfolding.

12  Biodiversity of Dermaptera

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Tervuren, Series no 8, Sciences Zoologiques 205: 1–335. Brindle, A. 1978. The Dermaptera of Africa. Pt II. Annales du Musée Royal de l’Afrique Centrale Tervuren, Series no 8, Sciences Zoologiques 225: 1–204. Brindle, A. 1980. The cavernicolous fauna of Hawaiian lava tubes. 12. A new species of blind troglobitic earwig (Dermaptera: Carcinophoridae), with a revision of the related surface living earwigs of the Hawaiian Islands. Pacific Insects 21: 261–274. Brindle, A, and V. Decu. 1977. Dermaptera from caves in Cuba. Résultats des Expéditions Biospéologiques Cubano‐Roumanies a Cuba 2: 373–375. Brousse‐Gaury, P. 1983. Sur l’existence d’une glande tibiale chez Labidura riparia Pallas et Forficula auricularia L. (Insecte, Dermaptère) [On the existence of a tibial gland in Labidura riparia Pallas and Forficula auricularia L. (Insecta, Dermaptera)]. Comptes Rendus de l’Academie des Sciences Serie III Sciences de la Vie 292: 1169–1172. Brousse‐Gaury, P. 1985. Découverte de sensilles dans le vagin de Labidura riparia Pallas (Dermaptere, Labiduridae). Annales Sciences Naturelles, Zoologique, Botanique et Animales 7: 103–112. Butnariu, A. R., A. Pasini, F. S. Reis and E. Bessa. 2013. Maternal care by the earwig Doru lineare Eschs. (Dermaptera: Forficulidae). Journal of Insect Behavior 26: 667–678. Calixto, A., A. Dean, A. Knutson and M. Harris. 2006. Density changes of two earwigs, Labidura ripria (Pallas) and Euborellia annulipes (Lucas) following fire ant reduction in Mumford, Texas. Southwestern Entomologist 31: 97–101. Capinera, J. L. 2011. European earwig Forficula auricularia Linnaeus (Insecta: Dermaptera: Forficulidae). University of Florida/Institute of Food and Agricultural Sciences Extension EENY‐483 (IN875). Department of Entomology and Nematology, University of Florida, Gainesville, Florida. 5 pp. Caussanel, C. 1991. Ecophysiologie des comportments des Dermaptères existance de

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relationships within Neoptera (Insecta). European Journal of Entomology 98: 445–509. Haas, F., S. Gorb and R. J. Wootton. 2000. Elastic joints in dermapteran hind wings: materials and wing folding. Arthropod Structure & Development 29: 137–146. Haas, F., J. T. C. Hwen and H. B. Tang. 2012. New evidence on the mechanics of wing unfolding in Dermaptera (Insecta). Arthropod Systematics and Phylogeny 70: 95–105. Hehar, G., R. Gries and G. Gries. 2008. Re‐ analysis of pheromone‐mediated aggregation behaviour of European earwigs. Canadian Entomologist 140: 674–681. Herter, K. 1967. Weiteres zur Fortpflanzungsbiologie des Ohrwurmes Forficula auricularia L. Zoologische Beiträge NF 13: 213–244. Heymons, R. 1912. Über den Genitalapparat und die Entwicklung von Hemimerus talpoides Walker. Zoologische Jahrbücher Supplement 15: 141–184. IUCN. 2016. The IUCN Red List of Threatened Species. Version 2015‐4. http://www. iucnredlist.org [Accessed 25 June 2016]. Jarvis, K. J., F. Haas and M. Whiting. 2005. A phylogeny of earwigs (Insecta: Dermaptera) based on molecular and morphological evidence: reconsidering the classification of Dermaptera. Systematic Entomology 30: 442–453. Jordan, K. 1909a. Description of a new kind of apterous earwig, apparently parasitic on a bat. Novitates Zoologicae 16: 313–326 + plates. Jordan, K. 1909b. Notes on the anatomy of Hemimerus talpoides. Novitates Zoologicae 16: 327–330. Kamimura, Y. 2014. Pre‐ and postcopulatory sexual selection and the evolution of sexually dimorphic traits in earwigs (Dermaptera). Entomological Science 17: 139–166. Kamimura, Y. and C. Y. Lee. 2014a. Mating and genital coupling in the primitive earwig species Echinosoma denticulatum (Pygidicranidae): implications for genital evolution in dermapteran phylogeny. Arthropod Systematics and Phylogeny 72: 11–21.

Kamimura, Y. and C. Y. Lee. 2014b. Genital morphology and mating behaviour of Allostethus (Dermaptera), an earwig genus of enigmatic phylogenetic position. Arthropod Systematics and Phylogeny 72: 331–343. Klass, K. D. 2001. The female abdomen of the viviparous earwig Hemimerus vosseleri (Insecta: Dermaptera: Hemimeridae), with a discussion of the postgenital abdomen of Insecta. Zoological Journal of the Linnean Society 131: 251–307. Kocarek, P. 2009. A case of viviparity in a tropical non‐parasitizing earwig (Dermaptera Spongiphoridae). Tropical Zoology 22: 237–241. Kocarek, P., V. John and P. Hulva. 2013. When the body hides the ancestry: phylogeny of morphologically modified epizoic earwigs based on molecular evidence. PLoS ONE 8: e66900. Kocarek, P., L. Dvorak and M. Kirstova. 2015. Euborellia annulipes (Dermaptera: Anisolabididae), a new alien earwig in Central European greenhouses: potential pest or beneficial inhabitant? Applied Entomology and Zoology 50: 201–206. Kölliker, M. and M. Vancassel. 2007. Maternal attendance and the maintenance of family groups in common earwigs (Forficula auricularia): a field experiment. Ecological Entomology 32: 24–27. Kuhlmann, U. 1993. Techniques for rearing tachinid parasitoids of the European Earwig Forficula auricularia. Biocontrol Science and Technology 3: 475–480. Kuhlmann, U. 1994. Ocytata pallipes (Fallén) (Dipt., Tachinidae), a potential agent for the biological control of the European earwig. Journal of Applied Entomology 117: 262–267. Maes, J. M. and F. Haas. 2006. Dermaptera de Nicaragua. Revista Nicaraguense de Entomologia 66: 1–127. Marais, E., C. J. Klok, J. S. Terblanche and S. L. Chown. 2005. Insect gas exchange patterns: a phylogenetic perspective. Journal of Experimental Biology 208: 4495–4507. Matzke, D. and K. D. Klass. 2005. Reproductive biology and nymphal development in the basal

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earwig Tagalina papua (Insecta: Dermaptera: Pygidicranidae), with a comparison of brood care in Dermaptera and Embioptera. Entomologische Abhandlungen 62: 99–116. Matzke, D. and P. Kocarek. 2015. Description and biology of Euborellia arcanum sp. nov., an alien earwig occupying greenhouses in Germany and Austria (Dermaptera: Anisolabididae). Zootaxa 3956: 131–139. McLean, M. and G. A. Horridge. 1977. Structural changes in light‐ and dark‐adapted compound eyes of the Australian earwig Labidura riparia truncata (Dermaptera). Tissue and Cell 9: 653–666. Miles, C. 2015. The Earwig Collection (Dermaptera) of the Manchester Museum, UK, with a complete type catalogue. European Journal of Taxonomy 141: 1–138. Mittal, O. P. and S. Sakai. 1996. Karyotype evolution in Dermaptera. Pp. 237–249. In S. Sakai (ed). Taxonomy of the Dermaptera. Proceedings of 20th International Congress of Entomology. Firenze, Italy. Morgan, W. P. 1928. Comparative study of the spermatogenesis of five species of earwigs. Journal of Morphology and Physiology 46: 241–273. Naegle, M. A., J. D. Mugleston, S. M. Bybee and M. F. Whiting. 2016. Reassessing the phylogenetic position of the epizoic earwigs (Insecta: Dermaptera). Molecular Phylogenetics and Evolution 100: 382–390. Nakata, S. and T. C. Maa. 1974. A review of the parasitic earwigs (Dermaptera: Arixeniina; Hemimerina). Pacific Insects 16: 307–374. Nishida, G. M. 1994. Hawaiian terrestrial arthropod checklist. Hawaii Biological Survey 94‐04: 80. Ozeki, K. 1958. Effect of corpus allatum hormone on development of male genital organs of the earwig, Anisolabis maritima. Scientific Papers of the College of General Education, University of Tokyo 8: 69–75. Ozeki, K. 1964. Studies on the ecdysal line of the earwig, Anisolabis maritima, during normal metamorphosis and that experimentally modified. Scientific Papers of the College of

General Education, University of Tokyo 14: 103–113. Ozeki, K. 1977. Supernymph formation by implantation of corpus allatum during the penultimate stage of the earwig, Anisolabis maritima. Scientific Papers of the College of General Education, University of Tokyo 27: 17–24. Pass, G. 1988. Functional morphology and evolutionary aspects of unusual antennal circulatory organs in Labidura riparia (Pallas) (Labiduridae), Forficula auricularia L. (Forficulidae) and Chelidurella acanthopygia Gené (Forficulidae) (Insecta: Dermaptera). International Journal of Insect Morphology and Embryology 17: 103–112. Pavón‐Gozalo, P., B. Milá, P. Aleixandre, J. A. Calderón, A. Zaldívar‐Riverón, J. Hernandez‐ Montoya and M. García‐París. 2011. Invasion of two widely separated areas of Mexico by Forficula auricularia (Dermaptera: Forficulidae). Florida Entomologist 94: 1088–1090. Peck, S. B. 2001. Smaller Orders of Insects of the Galapagos Islands, Ecuador: Evolution, Ecology, and Diversity. NRC Research Press, Ottawa, Ontario, Canada. 278 pp. Peters, W. and I. Latka. 1986. Electron microscopic localisation of chitin using colloidal gold labeled with wheat germ agglutinin. Histochemistry 84: 155–160. Picker, M. and C. Griffiths. 2011. Alien & Invasive Animals. A South African Perspective. Struik Nature, Cape Town, South Africa. 224 pp. Popham, E. J. 1984. The genus Hemimerus, insect parasites of the giant rat. Nyala 10: 39–42. Rankin, S. M., H. B. Dossat and K. M. Garcia. 1997. Effects of diet and mating status upon corpus allatum activity, oocyte growth, and salivary gland size in the ring‐legged earwig. Entomologia Experimentalis et Applicata 83: 31–40. Rankin, S. M., M. A. Innocenti, C. A. Eicher and D. Furst. 2004. The effect of ventral nerve cord severance and male castration on female mating behavior, clutch size, and maternal care in the ring‐legged earwig. Comparative

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Biochemistry and Physiology, Part A: Molecular and Integrative Physiology 139: 533–541. Rehn, J. A. G. and J. W. H. Rehn. 1935. A study of the genus Hemimerus (Dermaptera, Hemimerina, Hemimeridae). Proceedings of the Academy of Natural Sciences of Philadelphia 87: 457–508. Rehn, J. A. G. and J. W. H. Rehn. 1937. Further notes on the genus Hemimerus (Dermaptera, Hemimerina, Hemimeridae). Proceedings of the Academy of Natural Sciences of Philadelphia 89: 331–335. Roulin, A. C., M. Wu, S. Pichon, R. Arbore, S. Kühn‐Bühlmann, M. Kölliker and J‐C. Walser. 2014. De novo transcriptome hybrid assembly and validation in the European earwig (Dermaptera, Forficula auricularia). PLoS ONE 9: e94098. Sauphanor, B. 1992. Une pheromone d’agregation chez Forficula auricularia [An aggregation pheromone in the European earwig Forficula auricularia]. Entomologia Experimentalis et Applicata 62: 285–291. Sauphanor, B. L., L. Chabrol, F. Faivre d’Arcier, F. Sureau and C. Lenfant. 1993. Side effects of diflubenzuron on a pear psylla predator Forficula auricularia. Entomophaga 38: 163–174. Sayah, F. 2002. Ultrastructural changes in the corpus allatum after azadirachtin and 20‐ hydroxyecdysone treatment in adult females of Labidura riparia (Dermaptera). Tissue and Cell 34: 53–62. Sayah, F., and A. M. Laverdure. 2001. Immunohistochemical localization of a bombyxin‐like peptide in the brain‐ retrocerebral complex of the insect Labidura riparia. Invertebrate Reproduction and Development 39: 1–8. Seeman, O. D. 2007. A new species of Paradiplogynium (Acari: Diplogyniidae) from Titanolabis colossea (Dohrn) (Dermaptera: Anisolabididae), Australia’s largest earwig. Zootaxa 1386: 31–38. Segura, N. A., W. Usaqúen, M. C. Sánchez, L. Chuaire and F. Bello. 2009. Succession pattern of cadaverous entomofauna in a semi‐rural area

of Bogota, Colombia. Forensic Science International 187: 66–72. Shimizu, S. and R. Machida. 2011. Reproductive biology and postembryonic development in the basal earwig Diplatys flavicollis (Shiraki) (Insecta: Dermaptera: Diplatyidae). Arthropod Systematics & Phylogeny 69: 83–97. Stärkle, M., and M. Kölliker. 2008. Maternal food regurgitation to nymphs in earwigs (Forficula auricularia). Ethology 114: 844–850. Steghaus‐Kovac, S. and U. Maschwitz. 1993. Predation on earwigs: a novel diet specialization within the genus Leptogenys (Formicidae: Ponerinae). Insectes Sociaux 40: 337–340. Suckling, D. M., G. M. Burnip, J. Hackett and J. C. Daly. 2006. Frass sampling and baiting indicate European earwig (Forficula auricularia) foraging in orchards. Journal of Applied Entomology 130: 263–267. Suzuki, S. 2010. Progressive provisioning by the females of the earwig, Anisolabis maritima, increases the survival rate of the young. Journal of Insect Science 10: 184. Suzuki, S. 2011. Provisioning mass by females of the maritime earwig, Anisolabis maritima, is not adjusted based on the number of young. Journal of Insect Science 11: 160. Suzuki, S., M. Kitamura and K. Matsubayashi. 2005. Matriphagy in the hump earwig, Anechura harmandi (Dermaptera: Forficulidae), increases the survival rates of the offspring. Journal of Ethology 23: 211–213. Tomkins, J. L. 1999. Environmental and genetic determinants of the male forceps length dimorphism in the European earwig Forficula auricularia L. Behavioral Ecology and Sociobiology 47: 1–8. Tuxen, S. L. 1938. Orthoptera and Dermaptera. In: The Zoology of Iceland. Volume 3, part 38. Levin & Munksgaard, Copenhagen, Denmark. 5 pp. Tworzydlo, W., E. Kisiel and S. M. Bilinski. 2013. Embryos of the viviparous dermapteran, Arixenia esau develop sequentially in two compartments: terminal ovarian follicles and the uterus. PLoS ONE 8: e64087.

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Vancassel, M. and R. Quris. 1994. Differential release of diapause in the earwig Forficula auricularia as indicator of respective contribution of two cohorts to the reproductive generation. Acta Oecologica 15: 63–70. Wagner, D. L. and J. K. Liebherr. 1992. Flightlessness in insects. Trends in Ecology and Evolution 7: 216–219. Walker, F. 1871. Description of Hemimerus talpoides. P. 2. In: Catalogue of the Specimens of Dermaptera Saltatoria in the Collection of the British Museum, Part V, Supplement. British Museum, London, UK. Walker, K. A. and R. D. Fell. 2001. Courtship roles of male and female European earwigs, Forficula auricularia L. (Dermaptera: Forficulidae), and sexual use of forceps. Journal of Insect Behavior 14: 1–17. Wan, X., M. I. Kim, M. J. Kim and I. Kim. 2012. Complete mitochondrial genome of the free‐ living earwig, Challia fletcheri (Dermaptera:

Pygidicranidae) and phylogeny of Polyneoptera. PLoS ONE 7: e42056. Weber, N. A. 1954. The insect fauna of an Iraq oasis, the city of Bagdad. Entomological News 65: 202. White, M. J. D. 1971. The chromosomes of Hemimerus bouvieri Chopard (Dermaptera). Chromosoma 34: 183–189. White, M. J. D. 1972. The chromosomes of Arixenia esau Jordan (Dermaptera). Chromosoma 36: 338–342. Wolff, M., A. Uribe, A. Ortiz and P. Duque. 2001. A preliminary study of forensic entomology in Medellin, Colombia. Forensic Science International 120: 53–59. Wong, J. Y., J. Meunier and M. Kölliker. 2013. The evolution of parental care in insects: the roles of ecology, life history and the social environment. Ecological Entomology 38: 123–137. Zinsmeister, P. P. 1973. RNA and protein synthesis in the earwig ovary. Journal of Insect Physiology 19: 1765–1770.

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13 Biodiversity of Grylloblattodea and Mantophasmatodea Monika J. B. Eberhard 1, Sean D. Schoville 2 and Klaus‐Dieter Klass3 1

Zoological Institute and Museum, University of Greifswald, Greifswald, Germany Department of Entomology, University of Wisconsin‐Madison, Madison, Wisconsin, USA 3 Senckenberg Natural History Collections Dresden, Museum für Tierkunde, Dresden, Germany 2

The Grylloblattodea (ice crawlers or rock crawlers) and Mantophasmatodea (heelwalk­ ers  or gladiators, also called “(African) rock crawlers” by Engel and Grimaldi 2004) are the two most recently described extant insect orders, with formal descriptions published in 1914 (Walker 1914) and 2002 (Klass et  al. 2002), respectively. They are also the two smallest insect orders, each comprising only a handful of described species (33 and 19, respec­ tively). Both taxa include moderately sized insects (1.0–4.5 cm in length) that lack wings, which were secondarily lost. They show lim­ ited morphological and ecological variability and are geographically restricted (Klass et  al. 2003, Schoville 2014, Wipfler et  al. 2015): extant Grylloblattodea to the northeastern Palearctic (including Japan) and northwestern Nearctic regions, and extant Mantophasmato­ dea to the Ethiopian Region (Fig. 13.1, Fig. 13.2, Fig. 13.3, Fig. 13.4). The distribution patterns of both groups are probably based on relictual taxa of  small population size, having under­ gone renewed diversification in fairly recent times (probably in the Miocene to Pleistocene epochs), which led to the current pattern of  strong, mainly regional endemism, with

syntopic occurrence (occurrence of different species at the same locality) being exceptional. Since the discovery of the Mantophasmatodea, numerous studies have provided evidence of a sister‐group relationship to the Grylloblattodea, based on morphology (Klass et al. 2003; Uchifune and Machida 2005; Beutel and Gorb 2006; Wipfler et al. 2011, 2014, 2015), DNA‐sequence data (Terry and Whiting 2005, Cameron et  al. 2006, Kjer et al. 2006, Djernæs et al. 2012, Wang et  al. 2013, Misof et  al. 2014), and neuropep­ tide  sequences (Gäde and Šimek 2010). The clade Mantophasmatodea + Grylloblattodea was named Xenonomia (Terry and Whiting 2005), derived from the Greek xenos (“stranger” or “outsider”) and onoma (“name”), reflecting the fact that both ordinal names include portions of the names of other orders to which they are not  closely related. Some authors combine the Grylloblattodea and Mantophasmatodea into one order, the Notoptera (Grimaldi and Engel 2005, Wang et  al. 2013), mainly to enable the inclusion of the extant wingless taxa in a single order, together with numerous winged fossils putatively related to them. This link to winged fossils, however, is multilayered and problem­ atic, for which reason we use the names

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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Figure 13.1  Currently known distribution of Grylloblattodea (circles) and Mantophasmatodea (squares). For the latter, the occurrence of fossil taxa (represented by a cross in a circle) is also included. Original by authors.

Grylloblattodea and Mantophasmatodea only for the wingless crown‐group taxa. Genomic approaches suggest a split between the two orders in the late Jurassic, about 153 million years ago (mya) (Misof et al. 2014), although at this time insects resembling typical Manto­ phasmatodea probably already existed (Huang et al. 2008). Given their close relationship, the two orders have little in common morphologically. In the context of variation observed across polyneop­ teran orders, body parts such as the head, anten­ nae, legs, genitalia, and cerci are significantly different; similarity in other body parts is mostly plesiomorphic (as in the biting mouthparts). Suggested synapomorphies of the taxa (e.g., Wipfler et al. 2015) are mostly subtle (e.g., the occurrence of particular muscles, and the struc­ ture of circulatory organs (antennal hearts) at the internal base of the antennae). Similarities in thoracic morphology, mostly correlated with winglessness, are partly apomorphies peculiar to these two taxa, suggesting that their last com­ mon ancestor was already wingless. Although research on both orders has in­ ­ creased in recent years, our knowledge of their behavior, biodiversity, ecology, and physiology remains limited, providing a plethora of exciting research topics for future investigations.

13.1 ­Grylloblattodea The Grylloblattodea were first described by the Canadian entomologist E. M. Walker, based on  two female specimens collected in 1913 on Sulphur Mountain, Banff, Canada (Walker 1914). Within a decade, species were discovered in California and Japan (Caudell 1923, Caudell and King 1924), suggesting a much broader distribution of this rare insect order. All extant members of the order are classified in a single family, Grylloblattidae (whose association with fossil taxa is doubtful). Its species occur in western North America, primarily in montane regions of California, the Pacific Northwest, and the northern and central Rocky Mountains including Canada (Fig. 13.2), as well as north­ eastern Asia, including Japan, the Sino‐Korean peninsula, far‐eastern Russia, and the Altai‐ Sayan mountains of southern Siberia (Fig. 13.3). 13.1.1  Morphology and Biology

The widely disjunct species have relatively minor morphological differences. Walker (1914) argued that grylloblattids represent a mixture of mor­ phological features common to different “orthop­ teroid” lineages, to such a degree that he named them by combining the Latin words for crickets

13  Biodiversity of Grylloblattodea and Mantophasmatodea

Figure 13.2  Detailed distribution map of Grylloblattidae in North America (genus Grylloblatta). Original by authors.

(gryllus) and cockroaches (blatta). Anatomi­ cal  studies have since identified a variety of plesiomorphic and apomorphic characters and shown that gryllob­lattids possess highly derived character states (autapomorphies), including, for example, reduced compound eyes (composed of 60  ommatidia or less), an internal spina on the

metathoracic sternum, a median eversible sac on the first abdominal sternum, a lacinia with a proximal tooth, a mono‐layered acrosome in the spermatozoon, and a loss of the ocelli, musculus craniohypopharyngealis, and musculus labroe­ pipharyngealis (Matsuda 1976, Dallai et al. 2005, Wipfler et al. 2011). In the male postabdomen, a

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Figure 13.3  Detailed distribution map of genera of Grylloblattidae in Asia. Original by authors.

complex of correlated apo­morphies includes the asymmetrical subgenital coxal lobes of segment IX (gonocoxites), a process formed by the ven­ tromesal end of abdominal tergite X, and the clasping function enabled by these morphologi­ cal traits. The Grylloblattidae (Fig. 13.5a–d) occur in rocky habitats, typically where cool tempera­ tures and high humidity prevail, primarily in mountainous landscapes or caves. Their com­ mon names, ice crawlers or rock crawlers, are based on their strong association with snow or rocky substrates. They forage nocturnally on snow, both in Siberia and North America, and so there has been considerable interest in their phys­ iological ability to tolerate cold tem­ peratures (Mills and Pepper 1937, Edwards and  Nutting 1950, Henson 1957). The genus Grylloblatta has been reported foraging at temperatures ranging from −2 to 7 °C (Campbell 1949, Kamp 1963, Schoville and Graening 2013), although they are known to choose temperatures between −3.5 and 5 °C in thermal gradient experiments (Mills and Pepper 1937, Henson 1957). Tests of critical temperature limits for Grylloblatta suggest a

narrow thermotolerance  to temperatures between −4.0 ± 0.8 and 27.0 ± 0.7 °C, while their supercooling points are at the same temperature (−3.9 ± 1.0 °C) as  their lower lethal limit (Mills and Pepper 1937, Morrissey and Edwards 1979, Schoville et al. 2015). Grylloblattids possess prognathous mouth­ parts that they use to scavenge for organic debris and small insects. In montane habitats, they forage for insects deposited and frozen on snow surfaces as aeolian (windswept) debris (Edwards 1987). At present, little is known about their trophic ecology, especially whether they are prey for other sympatric predatory insects (e.g., carabid beetles) or exhibit canni­ balism. Cannibalism might be one mechanism that explains the highly skewed sex ratio (toward females, which are larger), although the cause is  unknown. Edwards (1982) suggested that females of Grylloblatta are more active foragers owing to their higher energy demands than the smaller males, but because the skewed sex ratio is evident across different seasons (Schoville and Graening 2013), some other factor might account for the biased sex ratio.

13  Biodiversity of Grylloblattodea and Mantophasmatodea

Figure 13.4  Detailed distribution map of genera of Mantophasmatodea in southwestern Africa (i.e., Tanzaniophasmatidae not included). Original by authors.

Adults of the Grylloblattidae are about 2.0–4.3 cm long, and females are larger than males. Their body coloration varies from white, golden, or rufous, to brown, which can be uniform or bicolorous on the dorsal and ven­ tral  surfaces. Grylloblattids live 5–10 years (Nagashima et  al. 1982, Visscher et  al. 1982), although the upper bound is based on the num­ ber of instars and the molting rate observed in a laboratory setting. Nymphs molt at least seven times, and the instar can be identified by the number of cercomeres (ranging from two in first‐instar nymphs to 8–12 in adults). Nymphs

and adults are typically collected at the same time, so generations are overlapping. During mating, males and females interact by waving their antennae, suggesting chemorecep­ tion plays an important role in mate recognition (Nagashima et  al. 1982). Males typically chase and catch females with their mandibles, by grip­ ping onto various body parts, then slowly mov­ ing their grip to the prothorax. The male holds the female in place, twists his abdomen under­ neath her abdomen (always to the right side of the female due to his asymmetric genitalia), and uses the gonocoxites to hold her ovipositor in

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(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

Figure 13.5  (a–d) Grylloblattodea. (a) Female of Grylloblatta campodeiformis occidentalis. (b) Female of Grylloblattella sp. (c) Male of Galloisiana sp. (d) Female of Galloisiana yuasai. (e–h) Mantophasmatodea. (e) Female of Karoophasma biedouwense (Austrophasmatidae). (f ) Male of K. biedouwense. (g) Mating pair of K. biedouwense. (h) Female of Viridiphasma clanwilliamense. Photographs by authors. (See color plate section for the color representation of this figure.)

13  Biodiversity of Grylloblattodea and Mantophasmatodea

place while he inflates an eversible sac into her gonopore. Copulation can last from 30 minutes to more than 4 hours. Females may mate multi­ ple times, but it is possible that they store sperm and fertilize multiple egg clutches following a single copulation event. Oviposition occurs 10–50 days after copula­ tion, and 20–30 eggs are laid singly, usually in soil or on stones, with the deposition of each egg taking up to 3 minutes (Nagashima et  al. 1982). Thus, oviposition can be prolonged over a period of up to 10 days. Females lay up to 145 eggs over the course of their adult life, in cases where six separate clutches have been laid. Eggs are coal black and ellipsoidal, about 1.6 mm long and 0.75 mm wide. 13.1.2  Overview of Taxa

There are currently 33 described species (and four subspecies) of extant Grylloblattodea, with five genera placed in a single family, the Grylloblattidae (Table 13.1). Species have been defined based on morphological criteria, with mainly male genitalia, female ovipositor shape, and pronotum shape serving as diagnostic crite­ ria. Among the useful morphological characters of the male genitalic region, the shape of the subgenital plate, the shape of the right and left gonocoxites, and the shape of the primary copu­ latory sclerite and secondary accessory sclerites serve to differentiate species. The length, width, and curvature of the female ovipositor, as well as the pronotum shape, have also been used to dif­ ferentiate sympatric species, although they have not been studied extensively. Other notable fea­ tures that differentiate species include eye con­ dition (some taxa have reduced or completely absent eyes), body size, coloration, and anten­ nomere count. Genetic data seem to readily dis­ tinguish species, and have been important in identifying sympatric taxa (Jarvis and Whiting 2006, Schoville and Roderick 2010, Schoville et  al. 2013). These studies have suggested that many more species remain undescribed, based on extensive genetic differentiation among iso­ lated populations.

Genera can be distinguished by the pronotal shape, number of cercomeres (7–12), shape of the lacinia (one or two teeth), pubescence and setae (heavy and more spinose in Asian genera), differentially elongated third antennomere, com­ pound eye shape (large and round in Gryllo­ blatta), leg proportions (thinner and elongate in Gryl­loblatta), and shape and size of the ventro­ lateral pads on the tarsomeres. Genetic data are not clear on the relationships among genera, as different analyses support different relationships among the Asian and North American lineages (Jarvis and Whiting 2006, Schoville et al. 2013). Molecular and morphological analyses disagree on the systematics of Asian grylloblattids. In par­ ticular, molecular data suggest that Galloisiana is paraphyletic if the genus Nam­kungia is recog­ nized (Schoville and Kim 2011, Schoville et  al. 2013). Several morphological characters (the shape of the male supra‐anal plate, the presence of macrotrichiae on the cervical sclerite, and 12 cercomeres) support Nam­kungia (Storozhenko and Park 2002), although a revision of all available material has not been undertaken. Furthermore, within the genus Galloisiana, the species Galloisiana yezoensis represents a highly diver­ gent lineage  that is genetically equidistant from North American and all other Asian lineages (Scho­ville et al. 2013). The genus Grylloblatta (ice crawlers) includes 15 species (Table 13.1; plus three additional subspecies) described from California, Idaho, Montana, Oregon, Washington, Alberta and British Columbia (Fig. 13.2). At present, the highest species diversity of Grylloblatta occurs in California, but there are many known popu­ lations with extensive genetic differentiation throughout Washington and Oregon that might represent undescribed species (Jarvis and Whiting 2006). They are typically collected in summer months at high elevations, whereas they can be found year‐round in humid low‐ele­ vation caves. Ice crawlers are known from few collection sites, are typically low in abundance, and therefore have been considered for pro­ tected status owing to their rarity (Schoville and Graening 2013).

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Table 13.1  Extant species and subspecies of Grylloblattodea.

Taxon

Type locality

Reference including original description

Grylloblattidae Galloisiana chujoi

Megi‐shima Island, Shikoku, Japan

Gurney 1961

Galloisiana kiyosawai

Hirayu‐Onsen, Honshu, Japan

Asahina 1959

Galloisiana kosuensis

Gosu Cave, Chungcheongbuk‐do, South Korea

Namkung 1974b

Galloisiana nipponensis

Chuzenji, Honshu, Japan

Caudell and King 1924

Galloisiana notabilis

Nagasaki, Kyushu, Japan

Silvestri 1927

Galloisiana odaesanensis

Mt Odae, Gangwon‐do, South Korea

Kim and Lee 2007

Galloisiana olgae

Olga Mountains, Primorsky Krai, Russia

Vrsansky and Storozhenko 2001

Galloisiana sinensis

Changbai Shan, Jilin Province, China

Wang 1987

Galloisiana sofiae

Mt Myoyang, North Korea

Szeptycki 1987

Galloisiana ussuriensis

Lazovka River, Primorsky Krai, Russia

Storozhenko 1988

Galloisiana yezoensis

Mt Daisetsu , Hokkaido, Japan

Asahina 1961

Galloisiana yuasai

Tokugo‐Toge, Honshu, Japan

Asahina 1959

Grylloblatta barberi

North Fork Feather River, California, USA

Caudell 1924

Grylloblatta bifratrilecta

Sonora Pass, California, USA

Gurney 1953

Grylloblatta campodeiformis campodeiformis

Sulphur Mountain, Alberta, Canada

Walker 1914

Grylloblatta campodeiformis athapaska

Summit Lake, British Columbia, Canada

Kamp 1979

Grylloblatta campodeiformis nahanni

Mt McDame, British Columbia, Canada

Kamp 1979

Grylloblatta campodeiformis occidentalis

Mt Baker, Washington, USA

Silvestri 1931

Grylloblatta chandleri

Eagle Lake, California, USA

Kamp 1963

Grylloblatta chirurgica

Ape Cave, Washington, USA

Gurney 1961

Grylloblatta chintimini

Marys Peak, Oregon, USA

Marshall and Lytle 2015

Grylloblatta gurneyi

Lava Beds National Monument, California, USA

Kamp 1963

Grylloblatta marmoreus

Marble Mountains, California, USA

Schoville 2012

Grylloblatta newberryensis

Newberry Volcano, Oregon, USA

Marshall and Lytle 2015

Grylloblatta oregonensis

Oregon Caves National Monument, Oregon, USA

Schoville 2012

Grylloblatta rothi

Happy Valley, Oregon, USA; Neotype: Cultus Mountain, Oregon, USA

Gurney 1953; Marshall and Lytle 2015

Grylloblatta scudderi

Whistler Mountain, British Columbia, Canada

Kamp 1979

Grylloblatta sculleni

Scott Camp, Oregon, USA

Gurney 1937

13  Biodiversity of Grylloblattodea and Mantophasmatodea

Table 13.1  (Continued) Reference including original description

Taxon

Type locality

Grylloblatta siskiyouensis

Oregon Caves National Monument, Oregon, USA

Schoville 2012

Grylloblatta washoa

Echo Summit, California, USA

Gurney 1961

Grylloblattella cheni

Akekule Lake, Xinjiang Province, China

Wang and Yang 2010, in Bai et al. 2010

Grylloblattella pravdini

Teletskoye Lake, Altai Republic, Russia

Storozhenko and Oliger 1984

Grylloblattella sayanensis

Sambyl Pass, Khakassia Republic, Russia

Storozhenko 1996

Grylloblattina djakonovi djakonovi

Petrov Island, Primorsky Krai, Russia

Bey‐Bienko 1951

Grylloblattina djakonovi kurentzovi

Kedrovaya pad National Park, Primorsky Krai, Russia

Pravdin and Storozhenko 1977

Namkungia biryongensis

Biryong Cave, Gangwon‐do, South Korea

Namkung 1974a

Namkungia magnus

Balgudeok Cave, Gangwon‐do, South Korea

Namkung 1986

Asian grylloblattids (rock crawlers) are placed in four genera (Fig. 13.3): Galloisiana (Caudell and King 1924), Grylloblattina (Bey‐Bienko 1951), Grylloblattella (Storozhenko and Oliger 1984), and Namkungia (Kim and Lee 2006). Galloisiana nipponensis was described first, from Lake Chuzenji in central Honshu, Japan, and five additional species are known from the Japanese islands of Kyushu, Shikoku, Honshu, and Hokkaido. Six additional species of Gal­loisiana are known from the Korean peninsula, as well as bordering areas in China and far‐eastern Russia. Several species of Galloisiana in South Korea and southern Japan are notable for their lack of com­ pound eyes and their affinity for cave habitats. Two taxa are placed in Gryl­loblattina, which is restricted to the Primorsky Region of far‐­ eastern Russia and was first described from Petrov Island (Bey‐Bienko 1951). The genus Grylloblattella was discovered in southern Siberia, in the Altai Mountains near Lake Teletskoye (Storozhenko and Oliger 1984), and now two other species are known from the region, including the Sayan Mountains and the Altai Mountains in northern China. Finally, the large‐bodied genus Namkungia (Kim and Lee 2006) includes two cave‐specialized taxa from

South Korea. Namkungia magnus is the largest grylloblattid, nearly twice as large as most mem­ bers of Galloisiana. Asian grylloblattids range from sea level to nearly 3000 m elevation, and are typically found in rocky habitats during spring and fall, as long as local habitat condi­ tions remain humid. Siberian Grylloblattella have been observed foraging on snow at high elevations, similar to North American species.

13.2 ­Mantophasmatodea 13.2.1  Morphology and Biology The description of the Mantophasmatodea (Klass et  al. 2002) was based on two preserved museum specimens, collected in Namibia in 1909 and Tanzania in 1950. The first live speci­ mens were found in 2002 in Namibia (Zompro et al. 2002). In the short time since, the number of described extant species has increased to 19 (Table 13.2), and heelwalkers have been noticed to be quite common in many areas and habitats of southern Africa  –  in a region quite well explored by entomologists. This story reflects a highly unexpected addition to our knowledge of

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Table 13.2  Extant and fossil species of Mantophasmatodea.

Taxon

Type locality

Austrophasmatidae

Reference including original description*

Klass et al. 2003

Austrophasma caledonense

Caledon, Western Cape, SA

Klass et al. 2003

Austrophasma gansbaaiense

Grootbos (Gansbaai), Western Cape, SA

Klass et al. 2003

Austrophasma rawsonvillense

Rawsonville, Western Cape, SA

Klass et al. 2003

Hemilobophasma montaguense

Montagu, Western Cape, SA

Klass et al. 2003

Hemilobophasma sp.n.

De Rust, Western Cape, SA

Damgaard et al. 2008, Predel et al. 2012

Karoophasma biedouwense

Biedouw Valley, Western Cape, SA

Klass et al. 2003

Karoophasma botterkloofense

Botterkloof Pass, Northern Cape, SA

Klass et al. 2003

Lobatophasma redelinghuysense

Redelinghuys, Western Cape, SA

Klass et al. 2003

Namaquaphasma ookiepense

Ookiep, Northern Cape, SA

Klass et al. 2003

Viridiphasma clanwilliamense

Clanwilliam, Western Cape, SA

Eberhard et al. 2011

Striatophasma naukluftense

Naukluft Mountains, Namibia

Wipfler et al. 2012

Austrophasmatidae sp.n. 1

Anysberg, Lemoenfontein, Hillandale, Western Cape, SA

Damgaard et al. 2008

Austrophasmatidae sp.n. 2 = Austrophasmatidae gen.n. sp.n. “VR”

Van Rhynsdorp, Northern Cape, SA

Damgaard et al. 2008, Predel et al. 2012

Austrophasmatidae gen.n. sp.n. “RV” = Austrophasmatidae sp.n.4

Richtersveld, Northern Cape, SA

Predel et al. 2012

Mantophasmatidae

Klass et al. 2002

Mantophasma zephyrum

Unknown locality, Namibia

Klass et al. 2002

Mantophasma omatakoense

Omatako‐Farm, Namibia

Zompro and Adis 2006

Mantophasma kudubergense

Kuduberg, Erongo Mountains, Namibia

Zompro and Adis 2006

Mantophasma gamsbergense

Gamsberg, Windhoek District, Namibia

Zompro and Adis 2006

Sclerophasma paresisense

Paresisberg, Otjiwarango District, Namibia

Klass et al. 2003

Pachyphasma brandbergense

Brandberg Plateau, Namibia

Wipfler et al. 2012

Praedatophasma maraisi

Karasburg Region, Namibia

Zompro and Adis 2002

Tyrannophasma gladiator

Brandberg Mountain, Namibia

Zompro et al. 2003

Praedatophasma and Tyrannophasma clade

Tanzaniophasmatidae Tanzaniophasma subsolanum

Klass et al. 2003 Ufipa “Dish” (Dist.), Tanzania

Klass et al. 2002

Raptophasma kerneggeri

Baltic amber [Eocene]

Zompro 2001

Raptophasma groehni

Baltic amber [Eocene]

Zompro 2008

Adicophasma spinosum

Baltic amber [Eocene]

Engel and Grimaldi 2004

Fossil species

13  Biodiversity of Grylloblattodea and Mantophasmatodea

Table 13.2  (Continued)

Taxon

Type locality

Reference including original description*

Adicophasma grylloblattoides

Baltic amber [Eocene]

Arillo and Engel 2006

Juramantophasma sinica

Daohugou, Ningcheng County, Inner Mongolia, Northeast China [Jiulongshan Formation, Middle Jurassic]

Huang et al. 2008

Baltic amber [Eocene]

Zompro 2005

Species with unsupported assignment to Mantophasmatodea Ensiferophasma velociraptor (fossil)

* Or references where the species is mentioned in the case of undescribed species. SA, South Africa.

insect diversity on Earth. The superficial resem­ blance of mantophasmatodeans to nymphs of praying mantises or stick insects (hence their sci­ entific name) might be one reason for these ani­ mals having been overlooked for such a long time. Among the autapomorphies of the order, those on the antennae are most striking (Drilling and Klass 2010): the flagellum is sharply divided into a basiflagellum with 14 cylindrical anten­ nomeres, the distal ones of which show a weak subdivision near mid‐length, and a distiflagel­ lum with seven spindle‐shaped antennomeres, of which the first is especially long and the sec­ ond short. The division is also evident in the antennal sensilla equipment, as, for instance, only the distiflagellum bears trichoid sensilla (appearing as fine setation). Other striking apo­ morphies are found on the tarsi (Buder and Klass 2013), where small dorsal processes are present beyond the second tarsomere (minute) and the third tarsomere (larger). The processes bear one or two campaniform sensilla and prob­ ably perceive movements of tarsomeres relative to each other. Noteworthy apomorphies of the female genitalia, which form an ovipositor, are the fusion of the reduced gonapophyses (second valves) to the claw‐shaped gonoplacs (third valves) in segment IX and a special truncated shape of the gonapophyses of segment VIII (first valves). An apomorphic median process on the subgenital plate of males is used for drumming.

Heelwalkers are about 1–4 cm long, and females are usually larger than males (Hockman et al. 2009, Roth et al. 2014). Extant species are found south of the Sahara in Africa  –  so far in Namibia, South Africa, Malawi, and Tanzania, but they might also occur in the regions in between (Fig. 13.1, Fig. 13.4). Mantophas­matodea inhabit bushes, small trees, herbs, and grasses in open landscapes, where they prey on other arthropods, catching them with their spinose fore‐ and midlegs. They possess hypognathous (in German: “orthognath”), typical biting‐chewing mouthparts (Baum et  al. 2007). The body coloration is brown, grey, green, or yellow, and is either uniform or overlain by a mottled pattern; a dorsomedian stripe of different color usually adds to this. The Mantophasmatodea are called heelwalkers, as they usually keep their large arolia and last (fifth) tarsomeres of all legs lifted up and off the substrate. The arolia are put down only when walking on smooth surfaces, during copulation, or when handling large prey – that is, when they are used for tight attachment to a surface (Eberhard et al. 2009). As currently understood, the Mantophas­ matodea are annual and univoltine (Tojo et al. 2004, Roth et al. 2014). Females oviposit on the ground, usually next to inhabited bushes or grass tussocks. By mixing eggs with secretion and sand, egg pods are produced that contain 10–12 (Tojo et  al. 2004) or 20–30 eggs (Roth et  al. 2014). The hard, resistant egg pods are

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ideal for enduring the hot and dry season; dia­ pause lasts at least 8 months (Tojo et al. 2004, Hockman et al. 2009, Roth et al. 2014). After the first rains, nymphs hatch and disperse to nearby bushes and grass tussocks. The five nymphal instars can be distinguished by the number of annuli in the basiflagellum; two annuli are added, deriving from the most basal annulus in each instar, until the adult possesses 14 basifla­ gellar annuli (Hockman et  al. 2009). The time between hatching of nymphs and adulthood is 2–4 months (Zompro et  al. 2003, Hockman et al. 2009), depending on habitat and, perhaps, weather conditions (Tojo et al. 2004). Mate localization and recognition are achieved via percussive signals generated by both sexes. Males use the median process on their subgenital plate (also called the drum­ ming‐organ) to tap on the surface, whereas females drum the whole abdomen against the substrate (Eberhard and Picker 2008). Using this behavior, heelwalkers produce substrate vibra­ tions of a defined temporal pattern, transmitted through branches or blades of grass on which they reside. Sensitive sensory organs (chordo­ tonal organs) in all legs detect the substrate vibrations (Eberhard et al. 2010). The subgenual organ located proximally in the tibia is thought to perceive most of the vibratory information. Male vibratory signals consist of repeated groups of pulses (pulse trains), and the simpler female signals comprise repeated single pulses (one pulse = one tap with the abdomen on the ground). Analysis of the vibrational signals of 13 species of heelwalkers revealed that signals of both males and females are of similar overall structure but differ among species in temporal characteristics such as pulse rate or pulse‐train duration (Eberhard and Eberhard 2013). The only detailed behavioral study on heelwalker vibrational communication to date used two austrophasmatid species that occur in sympatry at Clanwilliam, South Africa (Eberhard and Picker 2008). Here, male and female vibrational signals differed significantly between species, and males and females of one species (Karoo­ phasma biedouwense) did not react to hetero­

specific vibratory signals (from Viridiphasma clanwilliamense) (Eberhard and Picker 2008). Because mating occurs in the absence of vibra­ tional communication when males and females are put close to each other (8–10 cm apart) (Eberhard and Picker 2008, Roth et  al. 2014), vibratory signals are thought to serve for mate localization and recognition at the mid‐range, and mainly serve to bring the sexes together in the complex bushes in which they reside. When the male arrives at the female’s posi­ tion, he slowly approaches her and then quickly leaps onto her back, grabbing her with his legs. The male bends his abdomen down in an S‐shape around the right side of the female, who lifts up her abdomen. The male’s large, curved, one‐ segmented cerci facilitate the coupling, when the mostly membranous phallus is evaginated and inserted into the female vagina (Tojo et al. 2004, Eberhard and Picker 2008, Roth et  al. 2014). Copulation lasts up to three days, during which the male does not feed, while the female still successfully preys and feeds (Zompro et al. 2003, Tojo et  al. 2004, Klass 2009, Roth et  al. 2014). Multiple matings have been observed, but no critical experiment has been conducted to investigate its consequences (such as the option of sperm removal). Many details on the morphology and biology of the Mantophasmatodea remain to be investi­ gated, either altogether or in a broader selection of species or populations. According to what is currently known, however, the Mantophas­ matodea are both morphologically and ecologi­ cally fairly uniform (Fig. 13.5e–h). Morphological variation mainly concerns differences in the postabdomen and coloration, and ecological variation mainly concerns the type of vegetation inhabited. 13.2.2  Overview of Taxa

To date, 19 extant species of Mantophasmatodea have been formally described (Table 13.2) from western South Africa, Namibia, and Tanzania; they are classified in 13 genera and four family‐ level groups (Klass et al. 2002, 2003; Picker et al.

13  Biodiversity of Grylloblattodea and Mantophasmatodea

2002; Zompro et  al. 2002, 2003; Zompro and Adis 2006; Eberhard et  al. 2011; Wipfler et  al. 2012; Roth et  al. 2014). Three additional mor­ phologically and genetically distinct species from South Africa (Damgaard et al. 2008) and at least two species with distinct neuropeptide patterns (Predel et al. 2012) are currently unde­ scribed (Buder and Klass 2013), along with spe­ cies from Malawi, which are likely congeneric with those from Tanzania (Roth et al. 2014). The discovery of additional species is likely. Five or six fossil species are also known (Table 13.2) – all from the Northern Hemisphere (Zompro 2001, 2005, 2008; Engel and Grimaldi 2004; Arillo and Engel 2006; Huang et al. 2008). On a morphological basis, heelwalker taxa can be best distinguished by characteristics of  the postabdomen (including the genitalia; Fig. 13.6). Females differ in the shape and scle­ rotization of the subgenital plate (Fig. 13.6a), the shape of the internal bulb of the spermatheca (Fig. 13.6b), and the presence or absence of vagi­ nal sclerites and of setation between gonapo­ physes IX of the ovipositor. In males, diagnostic characteristics are found in several genitalic sclerites and processes (their presence and shapes, e.g., genitalic hooks; Fig. 13.6f ), in the vomeroid (a sclerotized ventral projection of abdominal segment X; Fig. 13.6c), in the shape of tergite X (especially the hind margin; Fig. 13.6d), and details of the cerci (Fig. 13.6e) (Klass et al. 2003, Damgaard et al. 2008). A determina­ tion key for the heelwalker species then known was established by Klass et al. (2003). The spa­ tial relationship of setae and epidermal pigment spots, as well as the distribution of dark pig­ mentation over the compound eye (mottled/ mosaic versus striped), have proven to be useful diagnostic features (Klass et al. 2003, Eberhard et al. 2011). By contrast, the basic body color, as well as much of the finer details of the colora­ tion pattern, show considerable intraspecific variation (Klass et al. 2003). Coloration charac­ ters can vary among conspecific populations from different habitats or within populations, but this matter needs closer study. In terms of ratios, the one between eye height and gena

height (indicating relative size of compound eyes) is relevant for species distinction, whereas other explored ratios are apparently not (Klass et al. 2003, Eberhard et al. 2011). Other charac­ teristics, such as the tibial spination of the fore­ legs, the number of subdivided distal antennal basiflagellomeres, and structural details of the processes beyond the second and third tar­ someres, are less informative owing to high intraspecific variability combined with inter­ specific overlap of ranges (Klass et  al. 2003, Buder and Klass 2013). Molecular data from mitochondrial genes (cytochrome c oxidase I (COI) and 16S riboso­ mal DNA) were obtained for a large number of specimens from South Africa (Austrophas­ matidae) and a few from Namibia (Damgaard et al. 2008). The molecular results are congru­ ent with those from morphology in delimiting species (Klass et al. 2003, Damgaard et al. 2008). Bioacoustic data also agree with the aforemen­ tioned character systems, but the resolution and coverage of taxa are a bit more limited. Analyses of neuropeptide‐sequence patterns, with much larger samples from Namibia, led to slightly dif­ ferent results on species delimitation (Predel et al. 2012). However, resolution of this charac­ ter system is limited when it comes to closely related species. In addition, the determination of species was based mainly on the collection site instead of morphological characteristics or sequencing of previously used genetic markers, and cases of syntopy are known in Mantophas­ matodea (Eberhard et al. 2011, Roth et al. 2014). Therefore, the comparability of neuropeptide‐ based results with those derived from DNA sequences and morphology is limited. This has led to considerable problems in the taxonomy of the order (Buder and Klass 2013). The data from DNA sequences (COI and 16S; Damgaard et  al. 2008) and from neuro­ peptide sequences (Predel et al. 2012) were used to establish a phylogenetic classification of the  Mantophasmatodea. The two studies agree in the distinction of three monophyletic family‐ level clades: Austrophasmatidae, Mantophas­ matidae, and the Tyrannophasma + Praedato­­-

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Insect Biodiversity: Science and Society Austrophasma caledonense (d)

(b)

(a)

(c)

(f)

(e)

Lobatophasma redelinghuysense

(d) (c)

(a) (b) (f)

(e) Karoophasma biedouwense

(d) (b)

(c)

(a)

(f) (e) Namaquaphasma ookiepense

(d) (c)

(a)

(b)

(f) (e) Sclerophasma paresisiense

(d)

(b) (a)

(c) (e)

13  Biodiversity of Grylloblattodea and Mantophasmatodea

phasma clade. The fourth family, Tanzaniophas­ matidae, is monogeneric and remains defined on a morphological basis, as no data on DNA, neu­ ropeptides, or bioacoustics are available yet. Morphological data agree with this classification but have not yet been subjected to formal analy­ sis. The relationships between these family‐level clades are poorly resolved (Damgaard et al. 2008, Predel et al. 2012). 13.2.2.1 Tanzaniophasmatidae

The description of this family is only based on the morphology of a single dried male museum specimen of Tanzaniophasma subsolanum (Klass et al. 2002, 2003; Table 13.2), collected in Tanganyika Ufipa “Dish” (probably a misspell­ ing of “Dist.”), Tanzania, in 1950. Specimens from Malawi were also assigned to the family (Hockman et  al. 2009, Roth et  al. 2014), but there is no detailed description to confirm this. Characteristics of the females are thus not known, and data from DNA, neuropeptides, and vibrational signals are also not available for confirmed members of this family. The Tanzaniophasmatidae possess bilaterally symmetrical male genitalia different from the asymmetrical ones in other Mantophasmatodea, and a much more prominent median process (drumming organ) on the subgenital plate than in all other heelwalkers. The eyes are smaller and more projecting than in the other families. 13.2.2.2 Mantophasmatidae

Three genera of Mantophasmatidae have been described: Mantophasma with four species, and Sclerophasma and Pachyphasma with one species each (Table 13.2). The species of the

Mantophasmatidae possess moderately sized eyes (less high than gena). As far as known, they differ from the other heelwalker family clades in several elements of the male and female geni­ talia (e.g., the absence of male genital hooks, Fig. 13.6f; Klass et al. 2003). The status of the taxonomy is problematic, as postabdominal characters are well known for Sclerophasma paresisense (both sexes) and Mantophasma zephyrum (only females) (Klass et  al. 2003), whereas only a few of the relevant postabdominal characters have been considered in the subsequent description of the other spe­ cies (Zompro and Adis 2006, Wipfler et al. 2012). Additionally, numerous populations from differ­ ent localities in central and northern Namibia have been assigned to Mantophasma according to their neuropeptide pattern (Predel et al. 2012). Predel et  al. (2012) suggest incorporating all these specimens, including the previously described Mantophasma and Sclero­phasma taxa (Klass et al. 2002, 2003; Zompro and Adis 2006), into one species‐level taxon (i.e., Mantophasma zephyrum), owing to limited species‐level diver­ sification of neuropeptide sequences. However, for most of these populations, investigations on morphology and DNA sequences are missing. The few recordings of vibrational signals of five Mantophasma males collected at different sites in Namibia show differences in pulse rate, pulse‐ train duration and number of pulses per pulse train (Roth et al. 2014). In addition, S. paresisense and M.  zephyrum show distinct differences in the female genitalia (Klass et al. 2003). These dif­ ferences suggest categorization as different spe­ cies rather than assuming substantial variation at the intraspecific level (Buder and Klass 2013).

Figure 13.6  Selected postabdominal elements of five exemplary species of Mantophasmatodea, showing some species‐distinguishing characters. (a) Subgenital plate of female, ventral view, posterior up; double line is a virtual cutting line. (b) Spermathecal bulb of female, with innermost part of spermathecal tube. (c) Vomeroid of male, dorsal view, posterior up; left and right lateroventral tips of tergite X (articulating with vomeroid) included at bottom. (d) Abdominal tergite X of male, dorsal view, posterior up, basal parts of cerci included on top. (e) Distal half of cercus of male, posterolateral view, dorsal side up, apex to the right. (f ) One of the two phallic hooks of the male (element absent in Sclerophasma paresisense). Scale bars 0.5 mm (a,d), 0.2 mm (b,c,e), and 0.1 mm (f ). All illustrations modified from Klass et al. (2003).

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13.2.2.3  Tyrannophasma/Praedatophasma Clade

This clade includes two species, each placed in a monotypic genus by Zompro et  al. (2002, 2003): Praedatophasma maraisi (collected in or near the Richtersveld National Park, Namibia and South Africa) and Tyrannophasma gladia­ tor (collected on Brandberg Mountain, Namibia). The only diagnostic characteristic described so far is the strong spination on several body parts (especially strong in T. gladiator; Zompro et al. 2002, 2003) and the large body size (Roth et al. 2014). No detailed studies on the postabdomen have been conducted. Inclusion of individuals in  analyses of mitochondrial DNA (Damgaard et al. 2008) and neuropeptide sequences (Predel et al. 2012) yielded the two species in a strongly supported clade. Vibrational communication signals of T. gladiator (Eberhard and Eberhard 2013; erroneously named P. maraisi therein) show distinct differences from most other heelwalker species recorded so far in terms of short  pulse‐repetition times and long pulse‐ train‐­repetition times in male signals and short pulse‐repetition times in female signals. 13.2.2.4 Austrophasmatidae

This clade includes 14 currently known species (four of them undescribed; Table 13.2). Their dis­ tribution ranges seem to be mostly mutually exclusive (Fig. 13.4). Morphology and mitochon­ drial genes (Klass et al. 2003, Damgaard et al. 2008, Eberhard et al. 2011), vibrational communication (Eberhard and Picker 2008, Eberhard and Eberhard 2013), and neuropeptides (Predel et al. 2012) have all been studied extensively in nearly all the Austrophasmatidae. Based on phylogenetic results from COI and 16S and on morphological data (Klass et al. 2003, Damgaard et al. 2008), the group was classified in six genera: Karoophasma (K. ­biedouwense and K. botterkloofense), Hemilo­ bophasma (H. montaguense and an undescribed species), Lobatophasma (L. redelinghuysense), Austro­phasma (A. rawsonvillense, A. caledonense, and A. gansbaaiense), Namaquaphasma (N. ookiepense), and Viridiphasma (V. clanwillia­

mense). A genus‐level clade comprising the unde­ scribed “Austrophasmatidae sp.n.1 and sp.n.2” adds to this. The relationships among these genera are not well‐resolved, but either Namaquaphasma or Viridiphasma is sister to the other genera. All these taxa occur in western South Africa, where they live under winter rain conditions, in contrast to all other heelwalkers (Predel et  al. 2012). In addition, two further “basal” species were detected and classified more recently (Predel et  al. 2012, Wipfler et  al. 2012): “Austrophasmatidae sp.n.4” (Richtersveld, locality S01) is sister to all afore­ mentioned taxa, and Striatophasma naukluftense (Central Namibia) is the sister taxon of all remain­ ing Austrophasmatidae  –  and the only austro­ phasmatid species found outside South Africa. A set of about 21 morphological characters altogether allows a clear distinction of the known species, the male and female postabdo­ men being the major sources of characters (examples in Fig. 13.6). Vibrational communica­ tion signals also differ considerably between species; eight characters (five for male calls, three for female calls) were established and are sufficient for the discrimination of most spe­ cies. Sequences of COI (aligned: 810 nt) and 16S (aligned: 498 nt) from 93 austrophasmatid spec­ imens that represent 12 species were used to infer phylogenetic relationships in the Austro­ phasmatidae and to outline the various species. Both sequences show sufficient divergence between and within species to be of use in spe­ cies identification and in tracing intraspecific patterns of polymorphism; divergence is slightly higher in COI. Regarding the definition of spe­ cies, the results from morphology and mito­ chondrial genes are so far fully congruent. Congruence includes vibrational communica­ tion, but the data published so far are too sparse for judging whether this is consistently true. Neuropeptide data (Predel et al. 2012) are also largely congruent with the aforementioned character systems, but species identification might be problematic. Regional patterns of intraspecific variabil­ ity  are suggested by the mitochondrial gene

13  Biodiversity of Grylloblattodea and Mantophasmatodea

sequences; for well‐sampled species, haplotypes are usually more similar (or identical) in ­specimens from the same or nearby sites (which cluster in phylogenetic trees) than they are in specimens from sites farther apart. In K. bie­ douwense, regional differentiation in morphol­ ogy has been observed; populations from the two sides of the Cederberg Mountains showed some minor differences (Eberhard et  al. 2011; specimens from the western side have not yet been included in molecular studies). However, the sample sizes are too small for definite statements on regional patterns in intraspecific variation. The winter rainfall biomes of western South Africa form a biodiversity hotspot (e.g., Myers et al. 2000) with exceptional endemism and spe­ cies richness in many groups of organisms, including insects (e.g., Picker and Samways 1996). The three included biomes differ in the  amount of annual precipitation: the mesic fynbos (> 400 mm), the drier Succulent Karoo (200–400  mm), and the dry Nama Karoo ( 50 mm). Because no Neuropterida are known from Antarctica, this continent is excluded from discussions of con­ tinental distributions (i.e., “found on all conti­ nents” = “found on all continents except Antarctica”). The best extended single‐source reviews pub­ lished to date for each of the three orders of the Neuropterida are the following: Neuroptera, New (1989); Megaloptera, New and Theischinger (1993); and Raphidioptera, H.  Aspöck et  al. (1991). The treatments of the Neuroptera (New 1991a), Megaloptera (Theischinger 1991), and Raphidioptera (H. Aspöck and U. Aspöck 1991) in The Insects of Australia remain useful shorter summaries. Oswald and Penny (1991) cataloged the genus‐group names of the Neuropterida. Comprehensive and regularly updated online catalogs (Oswald 2015) and bibliographies (Oswald 2016) are also available for taxa and literature pertinent to each of the three orders. Two additional useful works that broadly review the biodiversity of the Neuropterida, and that contain useful compendia of repre­ sentative color illustrations of numerous spe­ cies, are those of U. Aspöck and H. Aspöck (1999, 2007).

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Table 21.2  An alphabetical list of the orders and families of the extant Neuropterida of the world (after Oswald 2015), with counts of genera and species. Order

Family

Megaloptera

Corydalidae

Genera

27

Sialidae Neuroptera

Ascalaphidae

295

8

78

100

431

Berothidae

28

126

Chrysopidae

81

1,415

Coniopterygidae

23

571

4

77

Hemerobiidae

28

591

Ithonidae

10

39

Dilaridae

Mantispidae

44

395

198

1,659

Nemopteridae

36

146

Nevrorthidae

4

19

Myrmeleontidae

Raphidioptera

Species

Nymphidae

8

35

Osmylidae

30

212

Psychopsidae

5

26

Sisyridae

4

71

Inocelliidae

7

42

Raphidiidae

26

206

Megaloptera (total)

35

373

Neuroptera (total)

603

5,813

Raphidioptera (total)

33

248

Neuropterida (total)

671

6,434

Table 21.3  A higher classification (order to tribe) of the extant Neuropterida of the world (after Oswald 2015). Order

Family

Subfamily

Megaloptera

Corydalidae

Tribe

Genera

Species

Chauliodinae

18

135

Corydalinae

9

160

8

78

Sialidae Neuroptera

Ascalaphidae

Albardiinae Ascalaphinae

1

1

71

328

Acmonotini

2

2

Ascalaphini

18

55

Encyoposini

9

38

21  Biodiversity of Neuropterida

Table 21.3  (Continued) Order

Family

Subfamily

Tribe

3

12

Proctarrelabrini

4

16

Suhpalacsini

6

79

Ululodini

3

57

Incertae sedis Haplogleniinae Allocormodini Campylophlebiini Melambrotini

1 68

26

95

1

7

1

1

11

24

1

6

Tmesibasini

1

10

Incertae sedis

11

47

1

7

Berothimerobiinae Berothinae

1

1

12

89

Cyrenoberothinae

3

3

Nosybinae

4

13

Nyrminae

1

1

Protobiellinae

2

2

Rhachiberothinae

3

13

Trichomatinae

2

3

Incertae sedis

1

1

Apochrysinae

6

25

Chrysopinae

Coniopterygidae

1 25

Proctolyrini

Incertae sedis

Chrysopidae

Species

Hybrisini

Ululomyiini

Berothidae

Genera

64

1,364

Ankylopterygini

5

101

Belonopterygini

15

155

Chrysopini

36

912

Leucochrysini

7

195

Incertae Sedis

1

1

Nothochrysinae

9

24

Incertae sedis

2

2

Aleuropteryginae

12

201

Aleuropterygini

2

101

Coniocompsini

1

24

Fontenelleini

9

72 (Continued)

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Table 21.3  (Continued) Order

Family

Subfamily

Tribe

Brucheiserinae Coniopteryginae

Genera

Species

2

4

9

370

Coniopterygini

6

277

Conwentziini

3

93

3

55

Dilaridae

Dilarinae Nallachiinae

1

22

Hemerobiidae

Adelphohemerobiinae

1

2

Carobiinae

1

9

Drepanacrinae

3

9

Drepanepteryginae

3

38

Hemerobiinae

5

228

Megalominae

1

40

Microminae

5

111

Notiobiellinae

4

84

Psychobiellinae

1

2

Sympherobiinae

3

65

Incertae sedis

1

3

Ithonidae Mantispidae

10

39

Calomantispinae

2

6

Drepanicinae

4

37

Mantispinae

35

319

Symphrasinae Myrmeleontidae

3

33

174

1,509

Acanthaclisini

16

103

Brachynemurini

16

91

Myrmeleontinae

Dendroleontini

36

187

Gnopholeontini

4

10

Lemolemini

7

14

Maulini

2

2

Myrmecaelurini

16

149

Myrmeleontini

13

242

Nemoleontini

61

631

Nesoleontini

3

80

21  Biodiversity of Neuropterida

Table 21.3  (Continued) Order

Family

Subfamily

Tribe

Palparinae Dimarini Palparidiini

22

140

3

8

1

3 124

Pseudimarini

1

2

Incertae sedis

1

3

Stilbopteryginae

2

10

17

48

Crocini

7

18

Necrophylini

9

29

Crocinae

Pastranaiini

1

1

19

98

Nevrorthidae

4

19

Nymphidae

8

35

Eidoporisminae

1

1

Gumillinae

1

2

Nemopterinae

Osmylidae

Psychopsidae

Kempyninae

4

20

Osmylinae

7

38

Porisminae

1

1

Protosmylinae

3

11

Spilosmylinae

5

118

Stenosmylinae

7

20

Incertae sedis

1

1

Psychopsinae

2

18

Zygophlebiinae

3

8

4

71

7

42

5

37

Sisyridae Raphidioptera Inocelliidae

Inocelliinae Inocelliini Neghini

Raphidiidae

2

5

26

206

Agullini

1

17

Alenini

1

10

Raphidiinae

Raphidiini Totals

Species

16

Palparini

Nemopteridae

Genera

24

179

671

6,434

Suborders, superfamilies, and subtribes have been excluded. Orders are listed alphabetically and all subtaxa are listed alphabetically within each higher taxon. Counts of genera and species are for the lowest‐ranked taxon in each row.

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21.6.1  Order Megaloptera 21.6.1.1  Family Corydalidae (Fig. 21.1a)

The Corydalidae, fishflies (Chauliodinae) and dobsonflies (Corydalinae), are a moderate‐sized family (295 species in 27 genera) with aggres­ sively predaceous aquatic larvae and large to very large (forewing length up to ca. 85 mm), mostly non‐feeding adults. The family is most species‐rich in montane regions with pristine, high‐gradient, gravel‐bottomed, cold‐water streams and rivers, which are the preferred hab­ itat for the larvae of most species. The family is  represented on most continents, although absent from Europe and poorly represented in Africa. The group is particularly diverse in the Oriental Region in the mountains of southern China and northern Indochina west to the Himalayas. A smaller number of species are  adapted to lower‐gradient, warmer‐water streams and springs, which correlates with the family’s lower species diversity in the low‐lati­ tude tropics outside mountainous regions. The family is particularly well known for the extremely long male mandibles of some species of the genera Corydalus (North and South America) and Acanthacorydalis (eastern Asia). Where found, this trait is the result of strong allometric growth and is distinctly sexually dimorphic. Perhaps best known in Corydalus, recent phylogenetic work (Contreras‐Ramos 1998) has shown that full development of this trait is mostly restricted to a few species in a rel­ atively derived northern clade within the genus. Most corydalid adults are rather inconspicu­ ously colored, but some species are strikingly marked with bold black and pale color patterns (e.g., Nigronia), or are largely bright yellow in life

(Chloronia). Because of their large size and ripar­ ian habits, corydalid adults became known to entomologists early on, and the first species was described by Linnaeus (1758), even though no species are native to Europe or the adjacent areas surrounding the Mediterranean and Black Seas. The large average size of corydalids and the pref­ erence of the larvae of many species for cold mon­ tane streams are factors that contribute to the particularly long lifespans of many species (3–5 years is not uncommon), although other species are univoltine, especially in warmer waters. Overall, however, corydalid species probably have the longest average lifespan of any family in the Neuropterida. Two subfamilies  –  Chauliodinae and Corydalinae  –  are generally recognized on the basis of adult head and terminalic traits, and both seem to be monophyletic. Several interge­ neric phylogenetic works have contributed to the development of a relatively advanced and solid basis for interpreting higher‐level relationships in the family (Glorioso 1981, Penny 1993, Liu et al. 2012b). The extensive corydalid fauna of China and adjacent southeastern Asia has been treated in recent years in a large series of papers by Liu and colleagues, much of which is summarized or cited by Yang and Liu (2010) and Liu et al. (2012b, 2016). Good faunal works exist for Australia (Theischinger 1983) and southern Africa (Liu et  al. 2013), and the revisions of Corydalus by Contreras‐Ramos (1998) and Chloronia by Penny and Flint (1982) effectively cover much of the fauna of Central and South America. 21.6.1.2  Family Sialidae (Fig. 21.1b,c)

The Sialidae, alderflies, are a small family (78 species in eight genera) with predaceous aquatic

Figure 21.1  Representative adults and larvae of the orders Megaloptera and Raphidioptera. (a) Corydalus sp., adult, Brazil (Megaloptera: Corydalidae). (b) Sialis lutaria, adult, Poland (Megaloptera: Sialidae). (c) Sialis lutaria, larva, Czech Republic (Megaloptera: Sialidae). (d) Ascalaphidae sp., larva, Nicaragua (Neuroptera: Ascalaphidae). (e) Suphalomitus sp., adult, Australia (Neuroptera: Ascalaphidae). (f) Spermophorella sp., adult, Australia (Neuroptera: Berothidae). (g) Chrysopidae sp., larvae, Colombia (Neuroptera: Chrysopidae). (h) Hypochrysa elegans, adult, Belgium (Neuroptera: Chrysopidae). Photo credits: Arthur Anker (a), Łukasz Prajzne (b), Jan Hamrsky (c), Marshal Hedin (d), Craig Nieminski (e), Shaun Winterton (f), Robert Oelman (g), Gilles San Martin (h). (See color plate section for the color representation of this figure.)

21  Biodiversity of Neuropterida

(a)

(b)

(c)

(e)

(g)

(d)

(f)

(h)

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larvae and small, largely non‐feeding adults (forewing length ca. 10–15 mm). However, unlike the Corydalidae, whose larvae tend to prefer gravelly substrates in streams with at least moderate currents, sialid larva are primar­ ily burrowers in fine‐grained, muddy sediments. Thus, although their microhabitats include pools in lotic environments, they are also char­ acteristic of lentic waters. The family is repre­ sented on all continents (including Europe), although quite restricted in distribution on most of the southern continents. The majority of species are found in the cool to cold temper­ ate regions of the globe (mostly in the north, fewer in the south), and are quite uncommon in lowland tropical areas. At least in temperate North America, adults in southern populations tend to emerge early in the year, during the colder months. Alderflies are similar in overall body form and appearance worldwide and most are uniformly black (or nearly so), although some species have partially pale or reddish col­ oration, particularly on the head and prothorax. Adults are weak fliers and rarely stray far from their aquatic larval habitats. Sialid larvae seem to be mostly general predators of a variety of small aquatic organisms, mostly insects and other arthropods. The monophyly of alderflies has never been seriously questioned and has been supported by a wide‐ranging morphological phylogenetic analysis (Liu et al. 2015), which also provides a much‐needed, well‐documented hypothesis for the pattern of phylogenetic relationships among the major lineages in the family. No subfamilies or tribes are currently recognized among the extant members of the family, which is ripe for a comprehensive revisionary treatment. 21.6.2  Order Neuroptera 21.6.2.1  Family Ascalaphidae (Fig. 21.1d,e)

The Ascalaphidae, owlflies, are a moderate‐ sized family (431 species in 100 genera) with predaceous terrestrial larvae and small to very large predaceous adults (forewing length ca. 15–60 mm). The family is known from all

continents, but only the largest subfamily, Ascalaphinae, is similarly cosmopolitan. The other two subfamilies are more restricted in distribution: Albardiinae is known from a sin­ gle species, Albardia furcata, from Brazil, and Haplogleniinae is found in the Afrotropical, Neotropical (with one or two species extend­ ing  into the southern Nearctic) and Oriental Regions. The adult body form is similar to that of antlions, with abdomen and wings elon­ gated, but ascalaphid bodies are generally more robust, and the long antennae (found in almost all species) are distinctive. In many respects, owlflies are among the most highly derived of all neuropterids. This is particularly true of their flight capabilities, which are probably the most “advanced” in the superorder. Adult owl­ flies are active aerial predators, and have been likened to dragonflies in their hunting abilities and agility in the air. Most species appear to be active (only?) during the hours of twilight (particularly at dusk) and to spend the rest of the day perched, but some Old World species are distinctly diurnal. Some species perch with the abdomen flexed dorsally at a wide angle to the resting substrate. A few South America species form communal roosting aggregations (an uncommon behavior for predators), to which some of the same individuals return on multiple days (Hogue and Penny 1988, Gomes‐ Filho 2000). The biologies and ecologies of ascalaphids are poorly known. For adults, this is at least in part due to the crepuscular activity period of most species, and also to their high mobility. The diurnal species of Eurasia are the best known. Owlfly larvae are solitary, sedentary predators; known species are primarily inhabitants of the litter and soil (Badano and Pantaleoni 2014b), live on the stems and leaves of plants, or climb on other elevated objects (e.g., rocks and fence posts). Larvae are usually distinctly flattened and bear prominent lateral scoli; many lie in wait for prey with their jaws opened at extremely large angles (180–270 º). Adult females of some South American species lay abortive eggs, called repagula, which, although apparently primarily

21  Biodiversity of Neuropterida

defensive in function, may serve as a first food source for newly eclosed first‐instar larvae. The phylogeny of the family has been poorly explored. In the past, it has widely been assumed to be monophyletic, but the interrela­ tionships of putatively basal ascalaphids and antlions (particularly the Albardiinae and Stilbop­teryginae) might not be as clear cut as once thought (Winterton et  al. 2010, Michel et al. 2016), and the monophyly of both groups is currently the subject of active investigation. It is widely believed that the current intrafamilial classification of the family is highly artificial. Many of the currently recognized suprage­ neric  taxa are poorly defined and likely not monophyletic (Badano and Pantaleoni 2014b). Species‐level monographs are available for some regions – that is, Australia (New 1984b), Europe (Badano and Pantaleoni 2014b), South America (Penny 1981a, b), and southern Africa (Tjeder 1992, Tjeder and Hansson 1992) – but accurate species identification is difficult to impossible in many parts of the world. 21.6.2.2  Family Berothidae (Fig. 21.1f)

The Berothidae, beaded lacewings, are a moder­ ate‐sized family (126 species in 28 genera) with predaceous terrestrial larvae and small adults (forewing length ca. 6–15 mm). The family is widespread, with representation on all conti­ nents, but is relatively poorly represented in the New World. The faunas of Africa and southern Asia are relatively diverse, and Australia has a distinctive endemic fauna (Aspöck and Randolf 2014). Of the eight subfamilies treated here, only the largest subfamily, Berothinae, is subcos­ mopolitan. All of the other subfamilies are small and restricted in distribution: Berothi­ merobiinae from Chile, Cyrenoberothinae from Chile and Southern Africa, Nosybinae and Rhachi­berothinae from the Afrotropical Region, Nyrminae from Anatolia, Protobiellinae from Australia and New Zealand, and Trichomatinae from Australia (Aspöck and Randolf 2014, Makarkin and Ohl 2015). Adult berothids super­ ficially resemble small hemerobiids, are primar­ ily nocturnal, and exhibit a variety of dietary

preferences, including pollen, small arthropods, and fungi (Monserrat 2006). At least some spe­ cies lay stalked eggs. The biologies of berothid species are mostly unknown, and the larvae of only six genera have been described to date (Aspöck and Randolf 2014). Larvae of some Berothinae are known to be hypermetamorphic, with active, feeding first‐ and third‐instar larvae and a quiescent, non‐feeding second instar. These species live and feed inside termite nests, but it is still unknown whether this behavior and habitat is characteristic for the entire family (Wedmann et al. 2013). The limits and monophyly of the Berothidae are currently unsettled questions. Although most of the family seems to represent a good clade, discussion is ongoing about its relation­ ship with the Mantispidae, and the proper posi­ tion of the raptorial‐forelegged rhachiberothines, which have been treated in the Berothidae (Makarkin and Ohl 2015), in the Mantispidae (Willmann 1990), or as a separate family (Aspöck and Mansell 1994, Aspöck and Randolf 2014, Liu et  al. 2015). If the rhachiberothines prove to be sister to either the main body of the Berothidae or the Mantispidae, a conservative treatment that placed the rhachiberothines as a subtaxon in whichever family is appropriate would have the benefit of not artificially increasing the number of family‐ranked taxa in the Neuroptera. The composition of berothid subfamilies at the genus level is also currently under active discussion (Aspöck and Randolf 2014, Makarkin and Ohl 2015), but progress toward a more stable phylogeny and classifica­ tion of the family is being made. In any event, the classification presented here will require future modification. A useful key to all of the genera (except the Rhachiberothinae) has recently been published (Aspöck and Randolf 2014), but few species‐level keys are available (Faulkner 1992, Winterton 2010, Machado and Krolow 2016). 21.6.2.3  Family Chrysopidae (Fig. 21.1g,h)

The Chrysopidae, green lacewings, are a very large family (1415 species in 81 genera) with

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predaceous terrestrial larvae and small to large, mostly predaceous adults (forewing length ca. 3–35 mm). The family is cosmopolitan in distri­ bution, with significant faunas on all continents. Three subfamilies are commonly recognized: the Chrysopinae are cosmopolitan (and contain ca. 97% of the world species); the Apochrysinae are restricted to tropical areas in Africa, Asia,  Australia, and the Americas; and the Nothochrysinae are widespread across Europe, Australia, southern Africa, South America, and western North America (Brooks and Barnard 1990). Chrysopids are ubiquitous and promi­ nent elements of the insect faunas of most habi­ tats, where many come readily to lights. Among entomologists, green lacewings are probably the most widely recognized of all neuropterid insect groups. Although many members of the general public also recognize them, their small size, nocturnal habits, and well‐camouflaged bodies detract from their popular prominence. The adults of most species are rather uniform in appearance, with a compact body and large, transparent wings held nearly vertically along the sides of the body. As their English common name suggests, most species are bright green, but other species show a wide range of other color schemes that incorporate black, brown, yellow, red, and orange. Although many species are predaceous as adults, many other species are not. Non‐predaceous adults feed on a vari­ ety of other substances, but perhaps most prominently on pollen, nectar, and honeydew (usually dried on the surfaces of plants). The adults of some species possess particularly interesting morphologies and behaviors that have attracted special attention. Among these are the tympanal organ at the base of the radial vein in most species (this is among the smallest “ears” known in insects and is tuned to detect the frequencies of echolocating bats) (Miller 1984), and duetting courtship behaviors (mostly in Chrysoperla species, males and females duet using volleys of abdominal oscillations with vibratory signals transmitted through the sub­ strate on which the pair stands) (Henry et  al. 2013). With minor exceptions (i.e., the genus

Anomalochrysa), female chrysopids lay their eggs atop silken stalks, generally on plants. The majority of chrysopid larvae seem to be associated with the leaves and stems of plants, where their phytophagous arthropod prey commonly feed. However, the larvae of other species occupy a broader range of microhabitats and exhibit a more diverse set of feeding strate­ gies and preferences. Some species, for exam­ ple, are found in ground litter and prey on a wide variety of small organisms, even snails (Jones 1941). Others are specialized predators feeding in the nests of certain ant species (Principi 1946). Principi and Canard (1984) pre­ sent a wide‐ranging discussion of chrysopid feeding. The larvae of many plant‐inhabiting species are active and voracious predators of soft‐bodied arthropods, particularly aphids, which have made them valuable to, and widely used in, biological control programs. A promi­ nent characteristic of the larval biology of many species is their penchant for self‐decoration. Commonly described as “trash carrying” or “debris‐carrying,” the larvae of many (but dis­ tinctly not all) species actively place various kinds of debris on their dorsal surfaces – prob­ ably functioning as camouflage, a physical bar­ rier to predation, or both  –  which often bear specialized morphological structures to support and retain the debris (e.g., elongate scoli and hooked setae; Tauber et al. 2014). Much of the biological literature on the Chrysopidae is con­ cisely summarized by Canard et al. (1984). The monophyly of the Chrysopidae as currently constituted seems well established (Winter­ton and Brooks 2002, Winterton and de Freitas 2006, Winterton et al. 2010). The monophyly of its subfamilies and tribes, however, is currently under active investigation, and it is likely that the composition and classification of a number of these will require changes based on new phy­ logenetic work. Brooks and Barnard (1990) pro­ vided a checklist to the world species, summary treatments for each genus, and a key to the world genera. Other significant works on the Chrysopidae include those of Tjeder (1966), H. Aspöck et  al. (1980), New (1980), Dorokhova

21  Biodiversity of Neuropterida

(1987), Ghosh (1990), Tsukaguchi (1995), X.‐k. Yang (1997), X.‐k. Yang et  al. (2005), Brooks (1997), de Freitas and Penny (2001), and Winter­ ton and Brooks (2002). Species‐level identifica­ tion of chrysopids is relatively difficult, often requiring examination of male terminalic char­ acters. Similarly, some genera cannot be keyed without recourse to male terminalia. In many parts of the world, species‐level identification of chrysopids is difficult to impossible with exist­ ing literature. 21.6.2.4  Family Coniopterygidae (Fig. 21.2a)

The Coniopterygidae, dustywings, are a large family (571 species in 23 genera) with predaceous terrestrial larvae and very small to medium‐sized adults (forewing length 2–6 mm in most species; > 9 mm in some Brucheiserinae). The family is cosmopolitan in distribution, with significant faunas on all con­ tinents, but is particularly diverse in the Neotropical and Palearctic Regions. The two largest subfamilies  – Coniopteryginae and Aleuropteryginae – are also cosmopolitan, but the small relictual subfamily Brucheiserinae is known only from Chile and Argentina (Sziráki 2011). Dustywings are the smallest members of the order Neuroptera. Their small size, short broad wings, highly reduced venation, and habit of coating the body with a whitish waxy powder produced from special body glands render them isolated among neuropterans. Larvae and adults are predators and are generally found in trees and bushes (Meinander 1972). The larvae are active predators, often feeding heavily on scale insects, mites, and whiteflies, and for this reason are sometimes used in biological control programs. The phylogenetic position of the family in the Neuroptera is still under discus­ sion. It is considered an important group phylo­ genetically, having been placed at the base (i.e., sister to the remaining Neuroptera) or near the base of the order by almost all workers over the past century. The ultimate resolution of its position has implications for interpreting the ancestrally aquatic or terrestrial nature of stem‐ lineage neuropterans. Although treated by

Meinander (1972), on which the classification used here is based, the internal relationships and higher classification of the family are in need of a broad, modern, phylogenetic study. Meinander (1972) comprehensively revised the family, which resulted in a flurry of additional taxonomic work and precipitated the subse­ quent species catalogs of Meinander (1990) and Sziráki (2011). The latter work contains com­ prehensive keys to the world species and is the preferred starting point for entry into the litera­ ture of the group. 21.6.2.5  Family Dilaridae (Fig. 21.2b)

The Dilaridae, pleasing lacewings, are a small family (77 species in four genera) with preda­ ceous larvae and very small to small adults (forewing length ca. 4–12 mm). The family is widespread; species are known from all conti­ nents except Australia, but the family is poorly represented in Africa. Most species (> 95%) are assigned to the genera Dilar and Nallachius. Adults are distinctive in having relatively broad wings that are densely setose, males possessing pectinate antennae, and females bearing an elongate ovipositor. New World Nallachius spe­ cies tend to rest with their wings spread out to the sides of the body and resemble small moths. The biologies of dilarid species are poorly known. The larvae of only a few species ( 90%) are contained in the two genera Sisyra and Climacia, the latter of which is restricted to the New World. Of the remaining two genera, Sisyborina is Afrotropical and Sisyrina is Oriental and Australian. The last comprehen­ sive, worldwide revision of the Sisyridae was that of Navás (1935), which is now out of date. Monserrat (1977) listed the world species, and several useful regional revisions and reviews are available: Bowles (2006, North America north of Mexico), Flint (2006, Neotropical Region), Monserrat (1981, Oriental Region), Parfin and Gurney (1956, New World), Penny

The Psychopsidae, silky lacewings, are a very small family (26 species in five genera) with pre­ daceous terrestrial larvae and small to large (forewing length ca. 10–35 mm) predaceous adults. The distribution of extant species is distinctly relictual, with individual species restricted to one of three areas of endemism: southern Africa (eight species), Australia (13 species), or southeastern Asia (five species). Extinct members of the family are known from other continents. Most species are uncommon in the field (and probably locally distributed), and their biologies are poorly known. Some Australian species have been found under the bark of Eucalyptus trees, where they may aggre­ gate around sap flows and feed on other insects attracted to the same. Adults hold their wings roof‐like over the abdomen, but at a low angle, so that specimens at rest are wide and relatively flat. Most adults are inconspicuously colored, but the wings of some Australian species bear distinctive bands and colored markings. The oviposition system of silky lacewings is abso­ lutely unique within the Neuropterida. Females possess a large, membrane‐lined chamber that is invaginated from the venter into the bulbous apex of the abdomen. The female uses a pair of articulated scraping appendages to produce finely granular mineral or vegetable matter, which is packed into the chamber. The granular material is then used to coat the eggs as they emerge from the ovipore, a complex behavior that is apparently accomplished while the female is flying and immediately before in‐flight ovipo­ sition (Oswald 1993a). Two subfamilies  –  Zygophlebiinae and Psychopsinae – have been recognized. The fam­ ily was last broadly monographed by Oswald (1993a), which contains genus‐level treatments and keys, and a catalog of the species. Tjeder

21.6.2.15  Family Sisyridae (Fig. 21.3e,f)

21  Biodiversity of Neuropterida

(1981, Amazon basin), Tjeder (1957, southern Africa), and Weißmair (1999, Europe). 21.6.3  Order Raphidioptera 21.6.3.1  Family Inocelliidae (Fig. 21.3g)

The Inocelliidae, inocelliid snakeflies, are a very small family (42 species in seven genera) with predaceous larvae and small to medium‐ sized (forewing length ca. 6–21 mm) adults; it contains approximately 15% of the world raphidiopteran species. Adults of this family are distinguished, by the absence of ocelli, from the raphidiid snakeflies, which possess ocelli. The distribution of the family is essen­ tially the same as that of the Raphidiidae. What little is known about the biology of the family is mostly similar to that of the Raphidiidae, but with several distinctive features: all known inocelliid larvae are corticolous (none geo­ philous); the general feeding habit of adults, although poorly known, does not seem to be predaceous (as in raphidiids); and inocelliid mating behavior appears to involve the physi­ cal attachment of the male’s head to the ventral surface of the female’s abdomen, using a pair of unique holdfast organs that evert from the male’s antennal toruli (no such organs or head attachment is known in the raphidiids) (U. Aspöck et al. 1994). Although the monophyly of the family seems well established (Haring et al. 2011, H. Aspöck et al. 2012), additional work is needed to estab­ lish phylogenetic relationships within the fam­ ily. The family was monographed by H. Aspöck et al. (1991) as part of their comprehensive revi­ sion of the world Raphidioptera. That work contains keys for taxa recognized up to that time. Additional helpful recent works include those by H. Aspöck et al. (2012) and Liu et al. (2009, 2010b, 2012). 21.6.3.2  Family Raphidiidae (Fig. 21.3h)

The Raphidiidae, raphidiid snakeflies, are a moderate‐sized family (206 species in 26 genera) with predaceous larvae and aggressively preda­ ceous small to medium‐sized (forewing  length

ca. 6–18 mm) adults; it contains approximately 85% of the world raphidiopteran species. Adults of this family are distinguished by the presence of ocelli from the inocelliid snakeflies, which lack them. The family is entirely restricted to the Northern Hemisphere, principally in three major distributional centers (which are generally shared by the Inocelliidae): the Mediterranean (Europe, Middle East, and northern Africa), central Asia, and western North America (southwestern Canada to southern Mexico). The distribution of the family is distinctly con­ fined to areas with temperate climates, and southern records are restricted to progressively higher altitudes (H.  Aspöck et  al. 1998). The intriguing distribution of this family has been attributed, in part, to the requirement of larvae for exposure to a period of low temperature to induce pupation (H. Aspöck 2002). Under artifi­ cial conditions, larvae not subjected to low temperatures continue to molt as larviform individuals but at some point begin to display developmental anomalies that partially incor­ porate pupal traits (prothetely). It has been hypothesized (H. Aspöck 1998) that the limita­ tion of extant Raphidioptera to cool‐adapted species in the Northern Hemisphere may be an historical artefact of the Cretaceous–Tertiary impact event, eliminating a formerly more extensive, warm‐adapted snakefly fauna that existed in the Mesozoic. Adults are arboreal predators that feed broadly on a wide range of small arthropods, particularly aphids (H. Aspöck 2002). Females lay eggs under the bark of living trees, or in the leaf litter or soil, with the help of their long ovi­ positor. Larvae are predators of soft‐bodied arthropods, particularly insect eggs and larvae; many are associated with the bark of trees (cor­ ticolous), whereas others are found in ground litter (geophilous) (H. Aspöck 2002). The monophyly of the family is well established, but work is ongoing to develop a better under­ standing of its internal intergeneric relation­ ships (Haring et al. 2011, U. Aspöck et al. 2012). The family was comprehensively monographed by H. Aspöck et al. (1991), which contains keys

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for taxa recognized up to that time. The works of H. Aspöck et al. (1998, 1999) and Liu et al. (2010a) provide points of entry into the more recent work on the family.

21.7 ­Societal Importance Most of the general public do not know neurop­ terid insects by name, but many will recall hav­ ing played with larval antlions in their pits as children; having seen delicate green lacewing adults gathered around porchlights on warm summer nights; or (in North America) remem­ ber the large and fearsome‐looking (but actually harmless) mandibles of a male dobsonfly, per­ haps having seen one on a wall near a light or on a trip to a local river or lake. Others, more observant and with a more highly honed curios­ ity of the natural world, might have noted with some wonder that the small “trash packet” they discovered wandering around on a leaf or stem turned out to be a decorated green lacewing larva, or have mistaken a mantisfly, with its large grasping forelegs, for a tiny praying man­ tid. As a group, neuropterids are widespread and fairly ubiquitous insects, but most are rather inconspicuous and go unnoticed by most people, particularly as the adults of most species are active primarily or only at night. Those with more entomological knowledge will know more about the interesting behav­ iors  and biologies of the common species, as well as know that neuropterids are generally predaceous insects, and therefore broadly classed as “beneficials.” It is this predatory behavior, exploited in the service of human agriculture, which accords neuropterid insects their primary societal importance. The vora­ cious feeding capacity and actively mobile prey‐searching behavior displayed by the lar­ vae of species in several families (particularly the Chrysopidae, Hemerobiidae, and Coniop­ terygidae) make them effective biological con­ trol agents of some of the most important pests of agriculture and horticulture (Senior and McEwen 2001). Many species naturally invade

agricultural ecosystems, and these populations can be artificially augmented to provide pri­ mary or contributory control of a wide range of  phytophagous arthropod pests, including aphids, scale insects, and mites (Canard 2001). A number of species have been used as key components in the integrated pest manage­ ment (IPM) strategies deployed in a variety of crops (e.g., apple, cherries, citrus, nuts, and ornamental plants; Szentkirályi 2001a, 2001b). An important factor contributing to the effi­ cacy of neuropterans in IPM programs is that techniques have been developed for the large‐ scale rearing of several species, particularly green lacewings in the genus Chrysoperla. This has enabled the development of a commercial market for these species and facilitated their use in augmentative biological control on a range of different crops and in a variety of dif­ ferent cropping systems (Nordlund et al. 2001). An extensive body of literature related to the beneficial use of neuropterans in agricultural and horticultural systems is summarized by McEwen et  al. (2001). Paradoxically, in some specialized agricultural contexts the predatory nature of neuropterans is detrimental, rather than beneficial, such as mantispid larvae prey­ ing on managed stingless bee colonies (Maia‐ Silva et al. 2013). Even more paradoxically, in a few cases predation may be viewed as either beneficial or detrimental in essentially identi­ cal agricultural systems, depending on the desired “crop”; for example, larval hemerobiids (Sympherobius sp.) preying on cochineal scale insects (Dactylopius sp.: Hemiptera: Dacty­ lopiidae) is viewed as beneficial if the crop is the Opuntia cactus, upon which the scale is considered a pest (Pacheco‐Rueda 2011), but is viewed as detrimental if the “crop” is the scale insect itself, commercially reared on Opuntia as a source of red dye. Neuropterid insects also intersect with human activities in a variety of other more peripheral contexts. Corydalid larvae have been sold and consumed as human food (“magotaro‐ mushi”) in parts of eastern Asia (Sasaki 1915), and the harvesting of “hellgrammites” (also

21  Biodiversity of Neuropterida

larval corydalids) supports a small commercial bait fishery in the eastern United States (Nielsen and Orth 1988). Antlion larvae are used in sev­ eral traditional contexts – as oracles, as a treat­ ment against fever, and to initiate breast growth in young girls  –  by several native cultures in Africa (Kutalek and Prinz 2004). Neuropterans are also well represented in expressions of human artistry, both ancient and modern (Kevan 1992, Monserrat 2010). Although much of this usage is in visual imagery, a quick perusal of the Web will also reveal usage in the physical arts and crafts (e.g., jewelry, needlework, and pottery), literature (e.g., juvenile fiction: “Ace Lacewing: bug detec­ tive”), music (e.g., “Lacewing,” a band), and video. The word lacewing in particular, a com­ pact and euphonious compound of two conno­ tation‐rich English words, has proven to be evocative and metaphorically flexible and is widely used in a variety of contexts. Caricatures of neuropterid biology and morphology have even entered the popular imagination through the mass media  –  plucked from a terrarium filled with sand, the long‐jawed parasite aurally administered to Commander Chekov to render him susceptible to mind control in the space fantasy movie Star Trek II: The Wrath of Khan certainly seems to have been inspired by antlion biology and morphology, even if considerable artistic license was taken with the facts in the end.

21.8 ­Scientific Importance The phylogenetic position of the Neuropterida as one of the near‐basal lineages in the Holo­ metabola, and particularly its position as the presumptive sister group to the megaclade Coleoptera + Strepsiptera (the most species‐ rich clade in all of Animalia), has long grounded a deep general interest in matters pertaining to the Neuropterida among systematic entomolo­ gists. The fact that the superorder contains, within a relatively small number of species, such a large and varying array of strikingly different

biologies and life histories contributes signifi­ cantly to the fascination with the group by ento­ mologists outside the realm of systematics. The diverse and often highly specialized morpho­ logical, physiological, and behavioral systems developed in the Neuropterida have led to their use to investigate a wide variety of scientific concepts and phenomena. Over the course of nearly 50 years, Charles Henry and colleagues have investigated the obligatory duetting behavior displayed during courtship by (some) green lacewings in the genus Chrysoperla. Recently reviewed by Henry et  al. (2013), this model system involves the reciprocal exchange of substrate‐borne vibra­ tional signals produced by abdominal oscilla­ tions in duetting, conspecific, heterosexual pairs. This short‐range communication system has led to sympatric speciation within the genus at local and regional scales, and to the produc­ tion of a swarm of sibling species across the globe. This work has been influential in the the­ oretical development of sympatric speciation models, and has particular relevance to the use of chrysopids in biological control, much of which is based on Chrysoperla species. Substrate‐borne vibrations have also been investigated in the Neuropterida from the per­ spective of the morphological and physiological systems through which vibrations are sensed (e.g., Devetak 1998) and the use of sand‐borne vibrations by antlion larvae to detect and local­ ize prey (e.g., Devetak 2014). The advanced visual systems of adult asca­ laphids, many of which have eye lobes bearing differentiated ommatidia, have been the subject of studies focusing on the physiological adapta­ tions of eyes to detect different wavelengths of light (e.g., Gribakin et al. 1995). The capabilities of advanced sensory systems have also been a central theme in studies involving hearing in green lacewings (Miller and MacLeod 1966, Miller and Olesen 1979, Miller 1984), which have one of the smallest “ears” known in insects. Finally, pit‐building antlions have proven to be a remarkably interesting and flexible system for conducting manipulation experiments on a

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wide range of practical and theoretical topics, including (to name just a few) dispersion and group selection theory (Wilson 1974, Simberloff et al. 1978, Boake et al. 1984, Day and Zalucki 2000), the biomechanics of trap construction (Lucas 1982, Griffiths 1986), and the optimality and suboptimality of foraging (Bond 1980, Griffiths 1981, Lucas 1983, Scharf et al. 2011).

Acknowledgments The invitation to write this chapter placed us in the interesting and challenging position of broadly considering and synthesizing informa­ tion on many diverse aspects of this fascinating group of insects. Horst Aspöck, David Bowles, Kady Tauber, and one anonymous reviewer examined all or part of the manuscript and offered helpful suggestions. A number of other colleagues answered a late call to provide geo­ graphically broad input on the data in Table 21.1; our thanks to all for sharing their experience and perspectives. The late Norman Penny kindly provided us with a copy of the text of an earlier chapter that he had coauthored on the Neuroptera of Brazil, which helped get us started down the path that led to this review.

­References More complete bibliographical and dating infor­ mation about the references cited below can be found at http://lacewing.tamu.edu/Biblio/Main (Oswald 2016). Ábrahám, L. and Z. Mészáros. 2006. Further studies on the daily activity pattern of Neuroptera with some remarks on the diurnal activities. Acta Phytopathologica et Entomologica Hungarica 41: 275–286. Adams, P. A. 1970. A review of the New World Dilaridae. Postilla 148: i + 1–30. Ardila Camacho, A. and J. A. Noriega. 2014. First record of Osmylidae (Neuroptera) from Colombia and description of two new species

of Isostenosmylus Krüger, 1913. Zootaxa 3826: 315–328. Ardila‐Camacho, A. and A. García. 2015. Mantidflies of Colombia (Neuroptera, Mantispidae). Zootaxa 3937: 401–455. Aspöck, H. 1998. Distribution and biogeography of the order Raphidioptera: updated facts and a new hypothesis. In S. P. Panelius (ed). Neuropterology 1997. Proceedings of the Sixth International Symposium on Neuropterology (13–16 July 1997, Helsinki, Finland). Acta Zoologica Fennica 209: 33–44. Aspöck, H. 2002. The biology of Raphidioptera: a review of present knowledge. In G. Sziráki (ed). Neuropterology 2000. Proceedings of the Seventh International Symposium on Neuropterology (6–9 August 2000, Budapest, Hungary). Acta Zoologica Academiae Scientiarum Hungaricae 48 (Supplement 2): 35–50. Aspöck, H. and U. Aspöck. 1991. Raphidioptera (Snake‐flies, camelneck‐flies). Pp. 521–524. In I. D. Naumann (chief ed). The Insects of Australia. Volume 1. Second edition. Melbourne University Press, Melbourne, Australia. Aspöck, H., U. Aspöck and H. Hölzel. 1980. Die Neuropteren Europas (2 volumes). Goecke and Evers, Krefeld, West Germany. Volume 1: 495 pp., volume 2: 355 pp. Aspöck, H., U. Aspöck and H. Rausch. 1991. Die Raphidiopteren der Erde. Eine monographische Darstellung der Systematik, Taxonomie, Biologie, Ökologie und Chorologie der rezenten Raphidiopteren der Erde, mit einer zusammenfassenden Übersicht der fossilen Raphidiopteren (Insecta: Neuropteroidea) (2 volumes). Goecke & Evers, Krefeld, Germany. Volume 1: 730 pp., volume 2: 550 pp. Aspöck, H., U. Aspöck and H. Rausch. 1999. Biologische und chorologische Charakterisierung der Raphidiiden der östlichen Paläarktis und Verbreitungskarten der Kasachstan, Kirgisistan, Usbekistan, Turkmenistan und Tadschikistan nachgewiesenen Arten der Familie

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(Neuropterida: Raphidioptera: Raphidiidae). Stapfia 60: 59–84. Aspöck, H., U. Aspöck and C‐K. Yang. 1998. The Raphidiidae of Eastern Asia (Insecta, Neuropterida, Raphidioptera). Deutsche Entomologische Zeitschrift (N. F.) 45: 115–127. Aspöck, H., X‐Y. Liu and U. Aspöck. 2012. The family of Inocelliidae (Neuropterida: Raphidioptera). A review of present knowledge. Mitteilungen der Deutschen Gesellschaft für Allgemeine und Angewandte Entomologie 18: 565–573. Aspöck, U. and H. Aspöck. 1999. Kamelhälse, Schlammfliegen, Ameisenlöwen. Wer sind sie? (Insecta: Neuropterida: Raphidioptera, Megaloptera, Neuroptera). Stapfia 60: 1–34. Aspöck, U. and H. Aspöck. 2007. Verbliebene Vielfalt vergangener Blüte. Zur Evolution, Phylogenie und Biodiversität der Neuropterida (Insecta: Endopterygota). Denisia 20: 451–516. Aspöck, U. and H. Aspöck. 2008. Phylogenetic relevance of the genital sclerites of Neuropterida (Insecta: Holometabola). Systematic Entomology 33: 97–127. Aspöck, U., H. Aspöck and H. Rausch. 1994. Die Kopulation der Raphidiopteren: eine zusammenfassende Übersicht des gegenwärtigen Wissensstandes (Insecta: Neuropteroidea). Mitteilungen der Deutschen Gesellschaft für Allgemeine und Angewandte Entomologie 9: 393–402. Aspöck, U., E. Haring and H. Aspöck. 2012. Biogeographical implications of a molecular phylogeny of the Raphidiidae (Raphidioptera). Mitteilungen der Deutschen Gesellschaft für Allgemeine und Angewandte Entomologie 18: 575–582. Aspöck, U., X.‐Y. Liu and H. Aspöck. 2015. The Dilaridae of the Balkan Peninsula and of Anatolia (Insecta, Neuropterida, Neuroptera). Deutsche Entomologische Zeitschrift 62: 123–135. Aspöck, U. and M. W. Mansell. 1994. A revision of the family Rhachiberothidae Tjeder, 1959, stat. n. (Neuroptera). Systematic Entomology 19: 181–206.

Aspöck, U., J. D. Plant and H. L. Nemeschkal. 2001. Cladistic analysis of Neuroptera and their systematic position within the Neuropterida (Insecta: Holometabola: Neuropterida: Neuroptera). Systematic Entomology 26: 73–86. Aspöck, U. and S. Randolf. 2014. Beaded lacewings – a pictorial identification key to the genera, their biogeographics and a phylogenetic analysis (Insecta: Neuroptera: Berothidae). Deutsche Entomologische Zeitschrift 61: 155–172. Badano, D. and R. A. Pantaleoni. 2014a. The larvae of European Myrmeleontidae (Neuroptera). Zootaxa 3762: 1–71. Badano, D. and R. A. Pantaleoni. 2014b. The larvae of European Ascalaphidae (Neuroptera). Zootaxa 3796: 287–319. Barnard, P. C. 1981. The Rapismatidae (Neuroptera): montane lacewings of the Oriental Region. Systematic Entomology 6: 121–136. Beutel, R. G., F. Friedrich and U. Aspöck. 2010. The larval head of Nevrorthidae and the phylogeny of Neuroptera (Insecta). Zoological Journal of the Linnean Society 158: 533–562. Boake, C. R. B., D. Andow and P. K. Visscher. 1984. Spacing of ant‐lions and their pits. American Midland Naturalist 111: 192 194. Bond, A. B. 1980. Optimal foraging in a uniform habitat: the search mechanism of the green lacewing. Animal Behaviour 28: 10–19. Bowles, D. E. 2006. Spongillaflies (Neuroptera: Sisyridae) of North America with a key to the larvae and adults. Zootaxa 1357: 1–19. Brooks, S. J. 1997. An overview of the current status of Chrysopidae (Neuroptera) systematics. Deutsche Entomologische Zeitschrift (N. F.) 44: 267–275. Brooks, S. J. and P. C. Barnard. 1990. The green lacewings of the world: a generic review (Neuroptera: Chrysopidae). Bulletin of the British Museum of Natural History, Entomology 59: 117–286. Canard, M. 2001. Natural food and feeding habits of lacewings. Pp. 116–129. In P. McEwen, K., T. R. New and A. E. Whittington (eds).

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Lacewings in the Crop Environment. Cambridge University Press, Cambridge, UK. Canard, M., Y. Séméria and T. R. New (eds). 1984. Biology of Chrysopidae. W. Junk, The Hague, Netherlands. 294 pp. Carpenter, F. M. 1940. A revision of the Nearctic Hemerobiidae, Berothidae, Sisyridae, Polystoechotidae and Dilaridae (Neuroptera). Proceedings of the American Academy of Arts and Sciences 74: 193–280. Contreras‐Ramos, A. 1998. Systematics of the Dobsonfly Genus Corydalus (Megaloptera, Corydalidae). Thomas Say Publications in Entomology: Monographs. Entomological Society of America, Lanham, Maryland. ii + 360 pp. Day, M. D. and M. P. Zalucki. 2000. Effect of density on spatial distribution, pit formation and pit diameter of Myrmeleon acer Walker, (Neuroptera: Myrmeleontidae): patterns and processes. Austral Ecology 25: 58–64. de Freitas, S. and N. D. Penny. 2001. The green lacewings (Neuroptera: Chrysopidae) of Brazilian agro‐ecosystems. Proceedings of the California Academy of Sciences 52: 245–395. Dejean, A. and M. Canard. 1990. Reproductive behaviour of Trichoscelia santareni (Navas) (Neuroptera: Mantispidae) and parasitization of the colonies of Polybia diguetana R. du Buysson (Hymenoptera: Vespidae). Neuroptera International 6: 19–26. Devetak, D. 1998. Detection of substrate vibration in Neuropteroidea: a review. In S. P. Panelius (ed). Neuropterology 1997. Proceedings of the Sixth International Symposium on Neuropterology (13–16 July 1997, Helsinki, Finland). Acta Zoologica Fennica 209: 87–94. Devetak, D. 2014. Sand‐borne vibrations in prey detection and orientation of antlions. Pp. 319–330. In R. B. Cocroft, M. Gogala, P. S. M. Hill and A. Wessel (eds). Studying Vibrational Communication: Animal Signals and Communication 3. Springer, Berlin, Germany. Devetak, D., A. Špernjak and F. Janžekovič. 2005. Substrate particle size affects pit building decision and pit size in the antlion larvae

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Sekimoto, S. and K. Yoshizawa. 2011. Revision of the genus Osmylus (Neuroptera: Osmylidae: Osmylinae) of Japan. Insecta Matsumurana (New Series) 67: 1–22. Senior, L. J. and P. K. McEwen. 2001. The use of lacewings in biological control. Pp. 296–302. In P. K. McEwen, T. R. New and A. E. Whittington (eds). Lacewings in the Crop Environment. Cambridge University Press, Cambridge, UK. Scharf, I., Y. Lubin and O. Ovadia. 2011. Foraging decisions and behavioural flexibility in trap‐ building predators: a review. Biological Reviews 86: 626–639. Shi, C.‐F., S. L. Winterton and D. Ren. 2015. Phylogeny of split‐footed lacewings (Neuroptera, Nymphidae), with descriptions of new Cretaceous fossil species from China. Cladistics 31: 455–490. Simberloff, D., L. King, P. Dillon, S. Lowrie, D. Lorence and E. Schilling. 1978. Holes in the doughnut theory: the dispersion of ant‐lions. Brenesia 14/15: 13–46. Snyman, L. P., M. Ohl, M. W. Mansell and C. H. Scholtz. 2012. A revision and key to the genera of Afrotropical Mantispidae (Neuropterida, Neuroptera), with the description of a new genus. ZooKeys 184: 67–93. Sole, C. L., C. H. Scholtz, J. B. Ball and M. W. Mansell. 2013. Phylogeny and biogeography of southern African spoon‐ winged lacewings (Neuroptera: Nemopteridae: Nemopterinae). Molecular Phylogenetics and Evolution 66: 360–368. Stange, L. A. 1994. Reclassification of the New World antlion genera formerly included in the tribe Brachynemurini (Neuroptera: Myrmeleontidae). Insecta Mundi 8: 67–119. Stange, L. A. 2004. A systematic catalog, bibliography and classification of the world antlions (Insecta: Neuroptera: Myrmeleontidae). Memoirs of the American Entomological Institute 74: iv + 565. Szentkirályi, F. 2001a. Lacewings in fruit and nut crops. Pp. 172–238. In P. K. McEwen, T. R. New and A. E. Whittington (eds). Lacewings in the

Crop Environment. Cambridge University Press, Cambridge, UK. Szentkirályi, F. 2001b. Lacewings in vegetables, forests, and other crops. Pp. 239–291. In P. K. McEwen, T. R. New and A. E. Whittington (eds). Lacewings in the Crop Environment. Cambridge University Press, Cambridge, UK. Sziráki, G. 2011. Coniopterygidae of the World: Annotated Check‐list and Identification Keys for Living Species, Species Groups and Supraspecific Taxa of the Family. Lap Lambert Academic Publishing, Saarbrücken, Germany. vi + 249 pp. Tauber, C. A., M. J. Tauber and G. S. Albuquerque. 2014. Debris‐carrying in larval Chrysopidae: unraveling its evolutionary history. Annals of the Entomological Society of America 107: 295–314. Theischinger, G. 1983. The adults of the Australian Megaloptera. Aquatic Insects 5: 77–98. Theischinger, G. 1991. Megaloptera (alderflies, dobsonflies). Pp. 516–520. In I. D. Naumann (chief ed). The Insects of Australia. Second edition. Volume 1. Melbourne University Press, Melbourne, Australia. Tjeder, B. 1957. Neuroptera‐Planipennia. The Lace‐wings of Southern Africa. 1. Introduction and families Coniopterygidae, Sisyridae, and Osmylidae. Pp. 95–188 In B. Hanström, P. Brinck and G. Rudebec (eds). South African Animal Life. Volume 4. Swedish Natural Science Research Council, Stockholm, Sweden. Tjeder, B. 1960. Neuroptera‐Planipennia. The Lace‐wings of Southern Africa. 3. Family Psychopsidae. Pp. 164–209. In B. Hanström, P. Brinck and G. Rudebec (eds). South African Animal Life. Volume 7. Swedish Natural Science Research Council, Stockholm, Sweden. Tjeder, B. 1961. Neuroptera‐Planipennia. The Lace‐wings of Southern Africa. 4. Family Hemerobiidae. Pp. 296–408. In B. Hanström, P. Brinck and G. Rudebec (eds). South African Animal Life. Volume 8. Swedish Natural Science Research Council, Stockholm, Sweden.

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Tjeder, B. 1966. Neuroptera‐Planipennia. The Lace‐wings of Southern Africa. 5. Family Chrysopidae. Pp. 228–534. In B. Hanström, P. Brinck and G. Rudebec (eds). South African Animal Life. Volume 12. Swedish Natural Science Research Council, Stockholm, Sweden. Tjeder, B. 1967. Neuroptera‐Planipennia. The Lace‐wings of Southern Africa. 6. Family Nemopteridae. Pp. 290–501. In B. Hanström, P. Brinck and G. Rudebec (eds). South African Animal Life. Volume 13. Swedish Natural Science Research Council, Stockholm, Sweden. Tjeder, B. 1992. The Ascalaphidae of the Afrotropical Region (Neuroptera). 1. External morphology and bionomics of the family Ascalaphidae, and taxonomy of the subfamily Haplogleniinae including the tribes Proctolyrini n. tribe, Melambrotini n. tribe, Campylophlebini n. tribe, Tmesibasini n. tribe, Allocormodini n. tribe, and Ululomyiini n. tribe of Ascalaphidae. Entomologica Scandinavica 41 (Supplement): 3–169. Tjeder, B. and C. Hansson. 1992. The Ascalaphidae of the Afrotropical Region (Neuroptera). 2. Revision of the tribe Ascalaphini (subfam. Ascalaphinae) excluding the genus Ascalaphus Fabricius. Entomologica Scandinavica 41 (Supplement): 171–237. Tsukaguchi, S. 1995. Chrysopidae of Japan (Insecta, Neuroptera). Privately printed, Osaka, Japan. ii + 224 pp. Wedmann, S., V. N. Makarkin, T. Weiterschan and T. Hörnschemeyer. 2013. First fossil larvae of Berothidae (Neuroptera) from Baltic amber, with notes on the biology and termitophily of the family. Zootaxa 3716: 236–258. Weißmair, W. 1999. Präimaginale Stadien, Biologie und Ethologie der europäischen Sisyridae (Neuropterida: Neuroptera). Stapfia 60: 101–128. Weißmair, W. 2005. Schwammhafte (Insekta: Neuroptera: Sisyridae)—Parasiten der Moostiere (Bryozoa). Denisia 16: 299–304. Wichard, W., T. Buder and C. Caruso. 2010. Aquatic lacewings of family Nevrorthidae (Neuroptera) in Baltic amber. Denisia 29: 445–457.

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22 Biodiversity of Strepsiptera Jeyaraney Kathirithamby Department of Zoology, University of Oxford, Oxford, UK

Strepsiptera, often referred to as “twisted‐ wing” parasitoids, are bizarre insects with unusual morphology (Kinzelbach 1971a, 1971b, 1978; Kathirithamby 1989a, 1991a, 2009). They are a small order of holometabol­ ous insects, comprising 15 families, five of which are extinct (Table 22.1). All species are obligate entomophagous endoparasitoids for three of their larval stages. In the suborder Stylopidia, however, endoparasitism in the male continues through the pupal stage, and the pedomorphic female remains permanently embedded in the host up to maturity, and even up to the production of the motile first‐instar planidia. Strepsiptera also display unusual genetic characteristics. They possess one of  the smallest insect genomes (108 Mbp) (Johnston et  al. 2004), yet have one of the larger 18S ribosomal DNA sequences associ­ ated with a number of unique and unusually long insertions (Gillespie et  al. 2005, Mat­ sumoto et al. 2011). There are also two transfer RNA translocations that disrupt an otherwise ancestral insect mitochondrial genome (McMahon et  al. 2009). Both the mitochon­ drial DNA and the nuclear ribosomal DNA underwent a significant burst of molecular evolution in the early history of Strepsiptera (McMahon et al. 2011).

Strepsiptera parasitize a broad range of hosts, encompassing seven orders and 35 families of Insecta worldwide. The host insect orders are the Diptera, Blattodea, Hemiptera, Hymenop­ tera, Mantodea, Orthoptera and Zygentoma (Kinzelbach 1971a, 1971b, 1978; Kathirithamby 1989a, 1991a, 2009) (Fig. 22.1). Strepsiptera are cosmopolitan in distribution (except for one family), and are found in all habitats occu­ pied by insects, except the aquatic environ­ ment. Much remains to be discovered about the biodiversity of this insect order, primarily because of their tiny size, reclusive nature, and the extremely brief life of the free‐living adult male. Morphologically indistinguishable forms of Strepsiptera parasitizing several host species over large geographic areas have been docu­ mented, and morphological stasis is seen in both fossil and extant taxa. Molecular charac­ terization has found that cryptic species are widespread in Strepsiptera (Kathirithamby and Johnston 2004, Kathirithamby et  al. 2009a, Hayward et al. 2011, Matsumoto et  al. 2011, Isaka et  al. 2012, Nakase and Kato 2013). The wide host range in Strepsiptera might be a driver for speciation. The number of described species of Strepsiptera currently stands at around 630, but when the as‐yet‐undiscovered cryptic

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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Table 22.1  Hierarchical classification of Strepsiptera. Order Strepsiptera Kirby   Family †Protoxenidae Pohl et al.   Family †Phthanoxenidae Engel and Huang   Family †Kinzelbachillidae Pohl and Beutel   Family †Cretostylopidae Kathirithamby and Engel   Family †Mengeidae Pierce   Family Bahiaxenidae Bravo et al. Suborder Mengenillidia Kinzelbach   Family Mengenillidae Hofeneder Suborder Stylopidia Kinzelbach   Family Corioxenidae Kinzelbach (+1 fossil species) Infraorder Stylopiformia Kinzelbach   Family Myrmecolacidae Saunders (+14 fossil species)   Family Lychnocolacidae (Bohart)   Family Stylopidae Kirby (+1 fossil species)   Family Xenidae Saunders   Family Bohartillidae Kinzelbach (+3 fossil species)   Family Elenchidae Perkins (+1 fossil species)   Family Halictophagidae Perkins †Extinct.

species are revealed, this number might well prove an underestimate of their diversity. Except in brachypterous Delphacidae, host death occurs at the adult stage as follows: (i) after emergence of the last‐instar larval male and female Mengenillidae, (ii) after emergence of the free‐living adult male Stylopidia from the endoparasitic pupa, or (iii) after emergence of the motile planidia from the endoparasitic female Stylopidia (Kinzelbach 1971a, Kathiri­ thamby 2009). Strepsiptera differ in their biology and life history from the general characteristics pro­ posed by the dichotomous hypothesis regard­ ing parasitoids (Godfray 1994). Table 22.2 gives the comparisons among koinobionts (parasi­ toids that attack concealed immature insects, which continue to develop sometime after ovi­ position, Askew and Shaw 1986), idiobionts (parasitoids that consume and pupate in a host that has been stung or killed by the adult female

before oviposition, Askew and Shaw 1986), and Strepsiptera. The two most important compari­ sons are the following. First, in contrast to other insect parasitoids, a living mobile host is essen­ tial for completion of the strepsipteran life cycle, whereas koinobionts and idiobionts either develop in the host for only one or two stages, or, as a result of parasitization, the host is paralyzed and development is prevented. Second, and more importantly, a stylopized host larva or nymph continues to metamor­ phose into an adult with the exception of brachypterous Delphacidae, where the free‐liv­ ing male emerges from a fifth‐instar host nymph, which does not metamorphose into an adult and dies soon after. Other insect para­ sitoids typically kill or consume the host after  the  larval development of the parasitoid (Table 22.2). The ability of Strepsiptera to keep the host alive until completion of their life cycle might be due to the larval endoparasitic “lag phase” during the host’s larval and early pupal stages. This lag phase serves to reduce the cost of parasitism during the early developmental stages of the host life cycle (Hughes and Kathirithamby 2005). This phase also enables the host to metamorphose to the adult stage, which is essential for the completion of the strepsipteran life cycle. The term macryno­ biont (a combination of the Greek words macryno, “lengthen”, and bios, “life”) was sug­ gested  to  characterize the unique association between  most Strepsiptera and their hosts (Kathirith­amby 2009). Strepsiptera are an ancient order of the Insecta, branching from the early Holometabola, and are a sister group of the Coleoptera (Niehuis et  al. 2012, Boussau et  al. 2014). Fossil species have been described from the following: mid‐ Cretaceous amber of Myanmar (Grimaldi et al. 2005, Engel et al. 2016, Pohl and Beutel 2016), mid‐Eocene Fushun Coalfield amber from northeastern China (Wang et  al. 2016), mid‐ Eocene Baltic amber (Menge 1866; Kulika 1978, 1979, 2001; Kinzelbach and Pohl 1994; Pohl and Kinzelbach 1995, 2001; Pohl et  al. 2005, Kathirithamby and Henderickx 2008;

22  Biodiversity of Strepsiptera

Mengenillidae

Zygentoma

Corioxenidae

Hemiptera Heteroptera

Hymenoptera Myrmecolacidae

Orthoptera Mantodea

Lychnocolacidae

Stylopidae + Xenidae

Elenchidae

Unknown

Hymenoptera

Hemiptera

Delphacidae, Dictyopharidae, Eurybrachidae and Flatidae

Blattodea Blattellidae and Blatidae

Diptera Tephritidae Halictophagidae

Hemiptera Aphrophoridae, Cercopidae, Cicadellidae, Delphacidae, Derbidae, Dictyopharidae, Flatidae, Membracidae, Tettigometridae, Tropiduchidae, Psyllidae, Coreidae, Pentatomidae, Scutelleridae

Orthoptera Tridactylidae

Figure 22.1  Representations of host preferences (excluding Bahiaxenidae) mapped onto a cladogram derived from a molecular phylogenetic analysis of the major lineages (adopted from McMahon and Kathirithamby 2008, redrawn by J. Paps) (photos by M. Hrabar and J. Kathirithamby) (not to scale) (Kathirithamby 2009).

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Table 22.2  Comparison of koinobionts, idiobionts (Askew and Shaw 1986, Strand 1986, Gauld and Bolton 1988, Godfray 1994, Strand and Peach 1995, Pennacchio and Strand 2006), and Strepsiptera (not in order of importance) (after Kathirithamby 2009). Koinobiont

Idiobiont

Strepsiptera

1  Endoparasites (a few ectoparasites)

Ectoparasites or endoparasites of sessile hosts

Obligate larval endoparasites; in Stylopidia, endoparasitoids as male pupal and obligate endoparasitoids as females

2  Development of host for one or two stages after parasitization

Parasitism prevents host development

Host development continues, including metamorphosis

3  Egg, larval, or pupal parasitoids

Egg, larval, pupal, or adult parasitoids

Larval or nymphal (sometimes egg) parasitoids

4  Parasitism restricted to one or two stages of host

Parasitism restricted to one or two stages of host

Parasitism continues through all host stages, which is required for life‐ history completion

5  Parasitize exposed hosts, which normally have shorter developmental time

Parasitize concealed hosts (pupae or eggs), which normally have longer developmental time

Parasitize both concealed and exposed hosts, with longer developmental time in females and shorter in males

6  Narrow host range

Wide host range from varied habitats

Wide host range from varied habitats

7  Hosts of similar size range

Hosts of varied sizes

Hosts of a wide range (3–110 mm).

8  Host consumed after development of parasite

Hosts paralyzed and consumed soon after parasitization

Live larval‐pupal‐adult or nymphal‐ adult stages (sometimes egg) of host required for life‐cycle completion, after which the adult host dies

9  Only temporarily paralyzed after parasitization

Permanently paralyzed after parasitization

Not paralyzed after parasitization

10  Host continues to be mobile after parasitization

Host becomes immobile after parasitization due to paralysis

Host continues to be mobile until emergence of the free‐living male and the live planidia from endoparasitic female

11  Higher fecundities

Lower fecundities

Fecundity always high (production of several hundred planidia)

Henderickx et al. 2013; Kogan et al. 2015), early Miocene Dominican amber (Kinzelbach 1979, 1983; Kathirithamby and Grimaldi 1993; Kin­ zelbach and Pohl 1994; Pohl and Kinzelbach 1995), in Messel Geiseltal oil slate in Germany (Lutz 1990), Eocene brown coal in Germany (Kinzelbach and Lutz 1985), and compressed shale from the Eocene Green River formation in Colorado (Antell and Kathirithamby 2016). These fossil finds indicate that, from ancient times, the Strepsiptera have had a widespread geographical distribution.

The first molecular phylogenetic analysis of the Strepsiptera revealed that the suborder Mengenillidia is the sister group of all late‐ branching Stylopidia. The Stylopidia, which have derived characteristics, are a monophyl­ etic group comprising the family Corioxenidae and infraorder Stylopiformia, which includes all other seven extant strepsipteran families (McMahon et  al. 2011). These results corro­ borate previous studies (Kinzelbach 1971a, 1971b; Kathirithamby 1989a, 2009; Pohl and Beutel 2005).

22  Biodiversity of Strepsiptera

The most significant phenotypic modifications of the Strepsiptera are extreme sexual dimor­ phism, entomophagy, continued endoparasitism of the stylopid pupal male, and the obligate endo­ parasitic, neotenic, pedomorphic female stylopid (Kathirithamby 2009; McMahon et  al. 2011). Strepsiptera also possess a unique immune‐ avoidance mechanism that seems to circumvent the host’s innate immunity (recognition of self versus non‐self). This mechanism allows them to make effective use of a host‐derived epidermal “bag” to cloak themselves from detection by the host (Kathirithamby et al. 2003). There are only two free‐living stages in all families of Strepsiptera (except Mengenillidae and Mengeidae; a free‐living female of the latter  has been identified in Baltic amber by J. Kathirithamby and M. Engel): the first‐instar larva (“planidium” is the recommended term in reference to the motile first‐instar larva of Strepsiptera; Clausen 1972, pp.18–19), and the adult male (Fig. 22.2). In the family Menge­ nillidae, and possibly the family Mengeidae,

(a)

both sexes emerge from the host to pupate externally (Fig. 22.3), and are free‐living as adults (Fig. 22.4). In Stylopidia, the males pupate endoparasitically in the host, with an extruded cephalotheca (Fig. 22.5), and emerge as free‐­ living adults (Fig. 22.2b, Fig. 22.4c), whereas neotenic, pedomorphic females remain endo­ parasitic with only an extruded cephalothorax (Fig. 22.6, Fig. 22.7) (Kinzelbach 1971a). The pedomorphic female of the Stylopidia does not undergo metamorphosis (Erezyilmaz et  al. 2014), whereas the female of the Mengenillidae undergoes complete metamorphosis, molt­ ing   to a pupal instar and then to an adult (Fig. 22.4b). On emergence, the adult male seeks a female and is instantly attracted to her by the phero­ mone she emits (Cvačka et  al. 2012, Tolasch et al. 2012, Hrabar et al. 2015, Zhai et al. 2016). Sexual selection is by sensory exploitation (Kathirithamby et al. 2015). During this period, the neotenic female super‐extrudes the cepha­ lothorax (Hrabar et al. 2014), and mating takes

(b)

Figure 22.2  Scanning electron micrograph. (a) Planidium of Stylops sp., a parasitoid of Andrena (Hymenoptera) (scale bar = 0.1 mm). (b) Frontal view of head of male Xenos vesparum (scale bar = 0.5 mm) (Kathirithamby 2009).

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(a)

(b)

22.1 ­Family Bahiaxenidae The family Bahiaxenidae is represented by the monotypic species Bahiaxenos relictus Bravo et al. (2009) from Brazil, known only from a sin­ gle male specimen, with no female or host records. Until more material is available, the family Bahiaxenidae is classified as incertae sedis, along with the fossil, extinct families Cretostylopidae, Mengeidae and Protoxenidae (Table 22.1, Table 22.3). A cladistic analysis placed this species as sister group to all extant Strepsiptera (Bravo et  al. 2009). Bahiaxenos is the only basal strepsipteran found in South America.

(c)

(d)

22.2 ­Suborder Mengenillidia The basal suborder Mengenillidia is the sister  group to all late‐branching Stylopidia (McMahon et  al. 2011). At present, Menge­ nillidae is the only family in this suborder. 22.2.1  Family Mengenillidae

Figure 22.3  Mengenilla sp. (a,b) Empty male puparium. (a) Dorsal view. (b) Ventral view. (c,d) Empty female puparium. (c) Dorsal view. (d) Ventral view (scale bar = 3 mm). Original by author.

place either via the brood canal opening, in Stylopidia (Meinert 1896, Hughes‐Schrader 1924, Lauterbach 1954, Linsley and MacSwain 1957, Kathirithamby 2000, Hrabar et  al. 2014, Kathirithamby et  al. 2015), or by traumatic insemination, in free‐living Mengenillidae (Par­ ker and Smith 1934; Silvestri 1940, 1941, 1943). The adult male has a short lifespan (ca. 3–6 hours) and dies soon after mating. Some families and genera of Strepsiptera are better known and studied than others, because they are found fairly frequently in traps.

Kinzelbach (1971a, 1971b) recognized three subfamilies, based on the structure of the male antenna and head capsule: Mengenillinae, Iberoxerninae, and Congoxeninae. The family Mengenillidae has five genera: Congoxenos, Eoxenos, Mengenilla, Trilineatoxenos, and Yemengenilla. Congoxenos, Eoxenos, and Mengenilla branch after Bahiaxenos (Bravo et  al. 2009). The status of the genera Tri­ lineatoxenos and Yemengenilla, however, is not  known because they were not included in the analysis. The hosts for Mengenilla and Eoxenos are Lepismatidae, mainly of the genera Tricholepisma, Neoastero­lepisma, and Cteno­ lepisma (Carpentier 1939; Silvestri 1941, 1943; Delgado et al. 2014). The females and hosts of Congoxenos, Trilineatoxenos, and Yemengenilla are unknown. A total of 16 species of Mengenillidae are described from Mediterranean regions of Europe and North Africa extending south to

22  Biodiversity of Strepsiptera

Sudan and north to the Atlantic coast of Iberia; in the Middle East, Southeast Asia, East Asia; and as far south as Australia. Congoxenos, Eoxenos, Trilineatoxenos, and Yemengenilla are monotypic, although Eoxenos probably consists of more than one species (Kathirithamby unpublished). There are 12 described species of Mengenilla (Table 22.3). The distinguishing characters, plesiomorphic (ancestral) for the suborder, are the following: (i) at the fourth endoparasitic larval instar, both males and females generally emerge from the host to pupate externally (Fig. 22.3), and adult males (Fig. 22.4a) and females (which are wingless) (Fig. 22.4b) emerge from the puparium to be free‐living; (ii) a single birth opening in the female is used for the release of the planidia; (iii)  fertilization is by traumatic insemination,

with no specific area for sperm deposition (Parker and Smith 1933, 1934; Silvestri 1940, 1941, 1943); and (iv) the hindwing of the male has a single MA1 (first anterior media) vein. On emergence from the host, the last larval cuticle tans to form a hard puparium that is covered with thorn‐like structures and has a head, mouthparts, eyes, and legs (Fig. 22.4). Mengenillidae occupy mainly dry habitats, and the sclerotized puparium is presumably an adaptation to prevent water loss. In south­ ern Spain, Eoxenos and Mengenilla are found in areas covered with stones in almond and citrus plantations. Free‐living male mengenil­ lids emerge from the puparium at dusk and are nocturnal (J. Kathirithamby, J. A. Delgado, and F.  Collantes, personal observation) (Fig. 22.4a).

(a)

(b)

(c)

(d)

Figure 22.4  (a) Free‐living adult male of Eoxenos laboulbenei (dorsal view). (b) Free‐living neotenic female of Eoxenos laboulbenei (ventral view). (c) Free‐living male of Xenos vesparum (dorsal view). (d) Neotenic female of Xenos vesparum (dissected out of host) (dorsal view) (scale bar = 3 mm) (drawings by J. A. Delgado).

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(a)

(a)

(b) (b)

Figure 22.5  (a) Paper wasp, Polistes dominula, parasitized by male pupa of Xenos vesparum (top arrow) and neotenic female of Xenos vesparum (bottom arrow, cephalothorax) (scale bar = 5 mm). (b) Planthopper, Sogatella furcifera, parasitized by male pupa of Elenchus sp. (black arrow); note absence of external genitalia (scale bar = 2 mm) (Kathirithamby et al. 2015).

Only free‐living adult females of the genera Mengenilla and Eoxenos are known. Females of Mengenilla and Eoxenos possess structures char­ acteristic of adult holometabolous insects, except for the absence of wings (Fig. 22.4b). On emergence from the free‐living puparium, the female is inseminated by the male via traumatic insemination, and the planidia emerge via the birth opening (Parker and Smith 1934; Silvestri 1940, 1943). However, some females do not emerge from the puparium, but hibernate within it until the next year. In spring, the planidia hatch from the mother while she continues to remain inside the puparium (Parker and Smith, 1934; Silvestri 1941, 1943; J.  ­ Kathiri­ thamby,

Figure 22.6  (a) Cricket host parasitized by female of Caenocholax fenyesi sensu lato (arrow, cephalothorax). (b) Neotenic female of Caenocholax fenyesi sensu lato (dissected out of host). (Scale bars = 4 mm) (Kathirithamby et al. 2015).

­ ersonal observation). The release of planidia p larvae from an unemerged female is an extraor­ dinary phenomenon, and might reveal a “miss­ ing link” in the evolution of Strepsiptera: a prelude to obligate endoparasitism in female Stylopidia, the sister group to Mengenillidia. The Chalcidoidea Idiomacromerus gregarius and Hockeria mengenillarum parasitize pupae of Mengenilla (Silvestri 1943), with the former also being recorded as a pupal parasitoid of Eoxenos laboulbenei in Italy (Silvestri 1941). Nearly 70 years after this first record, E. laboulbenei parasitized by  I. gregarius (Fig. 22.8) was found in southern Spain, which is also a new country record (Delgado et al. 2014). A female puparium of Mengenillidae

22  Biodiversity of Strepsiptera (a)

(a)

(b)

(b)

Figure 22.8  (a) Female puparium of Eoxenos laboulbenei parasitized by larvae of Idiomacromerus gregarius (scale bar = 3 mm) (Delgado et al. 2014). (b) Empty female puparium of E. laboulbenei with newly emerged adult male of the chalcidoid parasitoid I. gregarius (scale bar = 3 mm).

Figure 22.7  (a). Scanning electron micrograph (SEM) of cephalothorax of female of Elenchus varleyi (Kathirithamby et al. 2015). (b) SEM of cephalothorax of female Paraxenos lugubris with planidia larvae emerging from brood canal opening (Kathirithamby et al. 2012a). (Scale bars = 0.1 mm).

parasitized by Ortochalcis mengenillarum (now Hockeria mengenillarum) in Portugal (Luna de Carvalho 1950) was later identified as I. gregarius (Luna de Carvalho 1953). Hockeria mengenil­ larum, Dibrachys microgasteri, and Merostenus sp. are listed as parasitoids of Mengenilla and Eoxenos (Noyes 2015). Owing to the superficial resem­ blance of the cocoons of D. microgasteri and puparia of Merostenus sp. to the free‐living puparia of Mengenillidae, the above two species have been mistakenly included as para­ sitoids of Mengen­ illidae. The chalcidoids H.  menge­nillarum and I.  gregarius are most probably host‐specific parasitoids, limited to parasitizing the strep­ sipteran family Mengenillidae. The host of E. laboulbenei is Lepisma aurea (Zygentoma), which is a myrmecophile that lives

as a commensal in nests of Messor barabarus (Formicidae) in St‐Jean Cap‐Ferrat, France (Carpentier 1939). Similarly, E. laboulbenei, which parasitizes myrmecophilic Thysanura, lives as a commensal in nests of Messor sp. in Mula, Murcia, Spain (J.  Kathirithamby, J.  A. Delgado, and F.  Collantes, unpublished data). The multitrophic food web consisting of ants, silverfish, strepsipterans, and the chalcidoid par­ asitoid in southern Spain, is an indication of how invertebrates have adapted to the arid conditions of this area. Perhaps a similar commensal rela­ tionship with ants led to the dual host relation­ ship of the sexes in the family Myrmecolacidae.

22.3 ­Suborder Stylopidia The suborder Stylopidia consists of the family Corioxenidae, which is a sister group to the rest  of Stylopidia, the infraorder Stylopifor­ mia  (McMahon et  al. 2011) (Fig. 22.1). The history of Strepsiptera host‐use across seven extant  families (except Bahiaxenidae, Lychno­ colacidae, and Bohartillidae, whose hosts are unknown) is summarized in Table 22.3. In an

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Table 22.3  Families and genera of Strepsiptera, with species numbers and hosts. Taxon

No. of species

Hosts, fossil deposits

Protoxenidae   Protoxenos

1

♂ Baltic amber; host/♀ unknown

1

♂ Cretaceous Burmese amber; host/♀ unknown

1

♂ Cretaceous Burmese amber; host/♀ unknown

1

♂ Cretaceous Burmese amber; host/♀ unknown

2

♂ Eocene Baltic amber; host/♀ recently identified

1

Host/♀ unknown

  Congoxenos

1

Host/♀ unknown

  Eoxenos

1

Thysanura: Lepismatidae

  Mengenilla

12

Thysanura: Lepismatidae

  Trilineatoxenos

1

Host/♀ unknown

  Yemengenilla

1

Host/♀ unknown

  Australoxenos

1

Host/♀ unknown

  Blissoxenos

1

Heteroptera: Lygaeidae

  Corioxenos

3

Heteroptera: Pentatomidae

  Dundoxenos

3

Heteroptera: Cydnidae

  Eocenoxenos

1

♂ Eocene Baltic amber; host/♀ unknown

  Floridoxenos

1

Host/♀ unknown

  Loania

2

Heteroptera: Lygaeidae

  Malagasyxenos

1

Host/♀ unknown

  Malayaxenos

3

Heteroptera: Lygaeidae

  Mufagaa

1

Host/♀ unknown

  Proceroxenos

1

Host/♀ unknown

  Triozocera

27

Hemiptera: Cydnidae

  Uniclavus

1

Host/♀ unknown

  Viridipromontoxius

1

Host/♀ unknown

Phthanoxenidae   Phthanoxenos Kinzelbachillidae   Kinzelbachilla Cretostylopidae   Cretostylops Mengeidae   Mengea Bahiaxenidae   Bahiaxenos Suborder Mengenillidia Mengenillidae

Suborder Stylopidia Corioxenidae

22  Biodiversity of Strepsiptera

Table 22.3  (Continued) Taxon

No. of species

Hosts, fossil deposits

Infraorder Stylopiformia Myrmecolacidae    Caenocholax

3

♂: Hymenoptera: Formicidae‐Dolichoderinae, Formicinae, Myrmicinae ♀: Orthoptera: Gryllidae

3

♂: Baltic and Dominican amber

2

♂: Shale, Green River Formation

  Kronomyrmecolax

1

♂: Fuschen amber

  Myrmecolax

34

♂: Hymenoptera: Formicidae‐Ecitoninae, Formicinae, Myrmeciinae, Ponerinae, Pseudomyrmecinae ♀: Mantodea: Mantidae‐Stagmatopterinae

1

♂ Dominica n amber

  Palaeomyrmecolax

6

♂ Baltic amber; host/♀ unknown

  Stichotrema

47

♂: Hymenoptera: Formicidae‐Myrmeciinae; ♀: Orthoptera: Tettigoniidae: Gryllidae; Mantodea: Mantidae

2

♂ Dominican amber

23

Host/♀ unknown

  Crawfordia

9

Hymenoptera: Andrenidae

  Eurystylops

5

Hymenoptera: Halictidae

  Halictoxenos

23

Hymenoptera: Halictidae

  Hylecthrus

3

Hymenoptera: Colletidae

Lychnocolacidae   Lychnocolax Stylopidae

  Jantarostylops

1

♂ Eocene Baltic amber; host/♀ unknown

  Kinzelbachus

1

Hymenoptera: Andrenidae; ♂ unknown

  Melittostylops

1

Hymenoptera: Melittidae; ♂ unknown

  Rozenia

3

Hymenoptera: Andrenidae

  Stylops

117

Hymenoptera: Andrenidae

Xenidae   Paragioxenos

1

Hymenoptera: Vespidae

  Paraxenos

44

Hymenoptera: Sphecidae

  Pseudoxenos

35

Hymenoptera: Vespidae

  Xenos

37

Hymenoptera: Vespidae

1

Host/♀ unknown

3

♂ Dominican amber

Bohartillidae   Bohartilla

(Continued)

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Table 22.3  (Continued) Taxon

No. of species

Hosts, fossil deposits

Elenchidae   Colacina

1

Hemiptera: Cicadellidae

  Deinelenchus

6

Hemiptera: Dictyopharidae, Eurybrachidae, Flatidae

  Elencholax

2

Host/♀ unknown

  Elenchus

19

Hemiptera: Delphacidae

  Protelenchus

1

♂ Dominican amber; host/♀ unknown

2

Blattodea: Blattidae, Blattellidae

  Callipharixenos

3

Hemiptera: Cicadellidae, Scutelleridae

  Coriophagus

13

Hemiptera: Pentatomidae, Coreidae

  Dipterophagus

1

Diptera: Tephritidae

  Halictophagus

96

Hemiptera: Aphrophoridae, Cercopidae, Cicadellidae, Delphacidae, Derbidae, Dictyopharidae, Eurybrachidae, Flatidae, Membracidae, Psyllidae, Tettigometridae, Tropiduchidae

  Stenocranophilus

5

Hemiptera: Delphacidae

  Tridactylophagus

12

Orthoptera: Mogoplistidae, Tridactylidae

Halictophagidae   Blattodeaphagus

attempt to infer the ancestral hosts, extant host preferences were mapped onto the molecular phylogeny of the major lineages of Strepsiptera (McMahon et al. 2011). This analysis suggested that (i) Callipharixenidae are a subfamily of the  Halictophagidae; (ii) Xenidae (parasitoids of Vespidae, Sphecidae, and Crabronidae), and Stylopidae (parasitoids of Apidae) are separate families; and (iii) the genus Lychnocolax, which has no host records so far, and historically was placed in Myrmecolacidae, is the sister group to the Stylopidae, Xenidae, Elenchidae, and Halictophagidae (McMahon et al. 2011). Analysis of the unusual features for the sub­ order reveals (i) endoparasitism in the males including the pupal stage, (ii) obligate endo­ parasitism in the females, and (iii) neoteny and paedomorphy in the females (Kinzelbach 1971a, 1971b; Kathirithamby 2009; McMahon et  al. 2011). The endoparasitic females do not undergo a pupal instar (Erezyilmaz et al. 2014), and remain larviform, exhibiting extreme neoteny and resemble a “bag of eggs,” with a

cephalothorax that is extruded through the host cuticle (Fig. 22.6, Fig. 22.7a). In addition, there is a reduction or loss of spiracles in the larvae and adult males, a loss of larval legs and pupal claws, and a reduction of the number of tarsomeres in the adult male Stylopidia (McMahon et  al. 2011). These modified adult male structures are due to a dramatic develop­ ment in the female stylopid, linked to obligate endoparasitism, paedomorphy, and neoteny. The result is the unique cephalothorax (a fusion of the head, thoracic and anterior abdominal segments) (Fig. 22.4d, Fig. 22.6, Fig. 22.7), and the associated structures (brood canal opening, brood canal, and genital ducts) used for receiv­ ing sperm (Fig. 22.4d, Fig. 22.7a) (Meinert 1896, Hughes‐Schrader 1924, Lauterbach 1954, Linsley and MacSwain 1957, Kathirithamby 2000, Hrabar et  al. 2014, Kathirithamby et  al. 2015). The brood canal and its associated struc­ tures also form a passage used for exit of the live planidia from the endoparasitic mother (Fig. 22.7b) (Smith and Hamm 1914; Linsley

22  Biodiversity of Strepsiptera

and MacSwain 1957; Kinzelbach 1971a, 1971b; Kathirithamby 1989a, 2009; Kathirithamby et al. 2015). The evolution of structures in the male, along with neoteny and paedomorphy in  the endoparasitic female, gave rise to the extreme sexual dimorphism in the suborder Stylopidia (Fig. 22.4d, Fig. 22.6). 22.3.1  Family Corioxenidae

The family Corioxenidae as a sister group to the Stylopiformia is well supported (McMahon et al. 2011). The members of the family are parasitoids of Heteroptera (Lygaeidae, Pentatomidae, and Cydnidae). The absence of mandibles is a unique feature of the Corioxenidae. Three subfamilies were proposed (Pohl et al. 1996), but because the subfamilies have a varying range of characters, such as the number of tarsomeres and antenno­ meres, until more material and further detailed study, all 14 genera are placed in the family Corioxenidae, as suggested by Cook and Tribull (2013). The 14 genera are Australoxenos, Blis­ soxenos, Corioxenos, Dundoxenos, Floridoxenos, Loania, Malagasyxenos Malayaxenos, Mufagaa, Proceroxenos, Triozocera, Uniclavus, Viridi­ promontoxius, and a fossil genus, Eocenoxenos, from Baltic amber (Hendrickx et  al. 2013)  – a  total of 47 described species. Twenty‐seven Triozocera species are of cosmopolitan distribu­ tion. Three Corioxenos species are from Africa, India, and Mexico; three Malayaxenos species are from Malaysia and Germany; and three Dundoxenos species are from Eastern Europe and Africa. The rest of the genera consist of monotypic species (Table 22.3). Females and hosts are known for only six gen­ era. The elongated cephalothorax of the female extrudes through the first abdominal segment dorsally, under the hemelytra, and is bent cau­ dally to 180 °. With its unusually long aedeagus, the male is able to inseminate the female through an opening in the cephalothorax. The  cephalothorax moves backwards caudally before the emergence of the planidia, which takes place either via an opening (Kirkpatrick 1937), or through the opening pierced by the male during insemination (Cooper 1938).

22.4 ­Infraorder Stylopiformia Unlike females of the Corioxenidae, females of the Stylopiformia are inseminated through the brood canal opening (Meinert 1896, Hughes‐ Schrader 1924, Lauterbach 1954, Linsley and MacSwain 1957, Kathirithamby 2000, Hrabar et  al. 2014, Kathirithamby et  al. 2015). The active planidia also emerge from their mother via the brood canal opening (Smith and Hamm 1914; Linsley and MacSwain 1957; Kinzelbach 1971a, 1971b; Kathirithamby 1989a, 2000, 2009; Kathirithamby et al. 2015) (Fig. 22.7b). 22.4.1  Family Myrmecolacidae

There are three extant genera in Myrm­ ecolacidae, Caenocholax, Myrmecolax, and Stichotrema, and two fossil genera, Palaeomyrmecolax from Baltic amber and Kronomyrmecolax from Fushun amber, with a total of 99 described species. Of these, 15 are fos­ sil species  –  the largest number of fossil Strepsiptera described thus far (Table 22.3). Extant Myrmecolacidae have a circumtropical distribution, having been found in the African, Australian, and Neotropical Regions, and in the Nearctic Region only as far north as North Carolina in the east, but fossil species have been found in Colorado to the west. They are not currently found in the Palearctic Region, although seven fossil species  have been described from Baltic amber and one  from Fuschen amber in northeastern China. An extraordinary feature of the Myrme­ colacidae is the dimorphic host relationships of the sexes (Kathirithamby and Hamilton 1992), which is further compounded by their extreme sexual dimorphism (Fig. 22.6). Identifying the neotenic, pedomorphic females, which have a total absence of typical holometabolan insect characteristics, and matching them to their conspecific males is possible only by molecular characterization (Kathirithamby and Johnston 2004; Kathirithamby et  al. 2009a; Hayward et  al. 2011). Heterotrophic heteronomy (i.e., males and females parasitize different hosts,

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sensu Walter 1983) is an unusual form of poly­ morphism, rare not only for Strepsiptera but also for insect parasitoids in general. This unique, complex, and extreme form of heter­ onomous behavior is seen in only two lineages of sexually dimorphic parasitic insects: the strepsipteran family Myrmecolacidae and the hymenopteran family Aphelinidae. The males and females develop not only in different spe­ cies of insects but also in hosts that belong to different orders (Flanders 1959; Ogloblin 1939; Walter 1983; Kathirithamby 1991b, 2005; Kathirithamby and Hamilton 1992; Hunter and Woolley 2001). Male myrmecolacids develop as primary endoparasitoids of ants (Hymenoptera: Formi­ cidae) and the females as primary endoparasi­ toids of crickets, grasshoppers, (Orthop­ tera: Gryllidae, Tettigoniidae), and mantids (Manto­ dea) (Ogloblin 1939, Kathirithamby and Hamil­ ton 1992) (Fig. 22.6a). The host relationships of  the males and females are conserved in Myrmecolacidae. In Aphelinidae, however, they are conserved in the females (primary endopar­ asitoids), but are more variable in the males (which can be primary ectoparasitoids, endo­ parasitoids, or hyperparasitoids) (Hunter and Woolley 2001). Heterotropic heteronomy has been lost in a few species of Aphelinidae (Hunter and Woolley 2001), but in Myrmecolacidae it has so far been seen to be lost only once: in  a parthenogen from Papua New Guinea, Stichotrema dallatorreanum. There might be other cases of such loss, because out of the 84 extant species, only four conspecific males and females have been matched unequivocally by molecular characterization (Kathirithamby and Johnston 2004, Kathirithamby et  al. 2009a, Hayward et al. 2011). The aphelinid mother seeks an appropriate host and, furthermore, lays an egg of an appro­ priate sex. She is efficient in so doing, as her sex‐determination haplodiploid system ena­ bles her to control the sex of the egg (Hunter and Woolley 2001). In the Myrmecolacidae, however, the host‐seeking is performed by the free‐living planidia, which is a less‐efficient

mechanism, leading to high mortality rates due to failure to locate and parasitize the appropriate host during a short lifespan. Heteronomy in the Aphelinidae is fairly well studied (reviewed by Hunter and Woolley 2001). In the strepsipteran Myrmecolacidae, however, heteronomy remains poorly under­ stood, mainly due to the cryptic nature of sty­ lopized orthopterans. Of the described species of Myrmecolacidae, the majority are known only from free‐living adult males collected from light and Malaise traps. The hosts (ants) are known for the males  of eight known species of Myrmecolax, Stichotrema, and Caenocholax, and for an addi­ tional unknown eight species of Caenocholax and four of Myrmecolax. All the ant hosts are from the major subfamilies of Formicidae: Dolichoderinae, Ecitoninae, Formicinae, Myrmi­ cinae, Ponerinae, and Pseudomyrmecinae (West­ wood 1861; Ogloblin 1939; Hofeneder 1949; Kathirithamby 1991b, 2008; Kathirithamby and Johnston 1992, 2004; Kathirithamby and Hughes 2002; Hughes et  al. 2003b; Cook et  al. 2004; Kathirithamby et  al. 2007, 2009a, 2009b; Cook 2009; Hayward et  al. 2011; Nakase et  al. 2014; Pèrez‐Lachaud and Lachaud 2014). Of the eight described species of females (which are obligate endoparasitoids), all the hosts are known, and are from the orthopteran families Gryllidae and  Tettigoniidae and the order Mantodea (Hofeneder 1910, 1920, 1949; Ogloblin 1939; Luna de Carvalho 1972; Hirashima and Kifune 1974; Kifune 1983; Kathirithamby et  al. 2001, 2009a; Kathirithamby and Johnston 2004; Kathirithamby 2008; Hayward et  al. 2011) (Fig. 22.6a). One of many aspects of the myrmecolacid host–parasitoid relationships is the host‐seeking behavior of planidia in relation to the different hosts parasitized by the sexes. The hosts are located in different habitats; the ant larvae are in the nests, whereas the ground‐dwelling orthop­ teran or mantid nymphs are dispersed in the proximity of their emergence site. The planidia, emerging from the female strepsipteran mother (endoparasitic in an orthopteran or mantid

22  Biodiversity of Strepsiptera

host), have little distance to travel, because the host populations are usually highly aggregated. The planidia would be in proximity to numer­ ous host nymphs, enabling parasitization of the abundant conspecific host species in the same vicinity. However, for the emerging planidia to parasitize the ant larvae, a mechanism is neces­ sary to enable them to move away from the emergence site to reach their future host ant lar­ vae in the nest. They might “hitchhike” to the ant’s nest via a foraging worker ant (phoresy). Owing to the brief search time and limiting resources available, once in the nest the planidia might accept and enter any species of ant larvae they encounter. Alternatively, a host (orthop­ teran or mantid) parasitized by a gravid endo­ parasitic female strepsipteran might be captured and taken back to the nest via predation by ants. Within the nest, the emerging planidia would parasitize the ant larvae in the nest. In both sce­ narios, the planidia would parasitize any species they encounter. Thus, as seen in Caenocholax, some male myrmecolacids are generalist parasi­ toids of ants (Hayward et al. 2011). The genus Caenocholax is widespread in the New World, with extant and fossil species exhibiting morphological stasis (Kathiri­ thamby and Grimaldi 1993, Kathirithamby and Hend­rickx 2008). The males and females of only four species of Caenocholax have been unequivocally matched (Kathirithamby and Johnston 2004, Hayward et al. 2011). A molec­ ular phylogeographic analysis of Caenocholax found the genus to be subdivided into several distinct lineages across the New World, con­ sistent with separate species (Hayward et  al. 2011). Some Caenocholax lineages show slight variation in the aedeagus and tenth abdominal segment, whereas in others the differences were difficult to identify and might represent true cryptic species (J. Kathirithamby, unpub­ lished data). Furthermore, the genetic diversity of Caeno­cholax is structured by host associa­ tion, and the hosts are uncoupled between the sexes, the males being generalist parasitoids in their ant hosts (parasitizing various subfami­ lies of ants), whereas the females are parasitic

in specific species of cricket hosts (Hayward et al. 2011). Two adult males of Caenocholax described from organic compression fossils of the Early Eocene (ca. 50 million years ago) Green River Formation (Colorado, United States) are not only the northernmost New World record of fossil or recent Myrmecolacidae, but are also the  oldest Myrmecolacidae reported so far (except for Kronomyrmecolax fushunicus from Fushun  amber, Wang et  al. 2016) (Antell and  Kathirithamby 2016). The occurrence of Caenocholax in Colorado illuminates the geo­ graphic history of Myrmecolacidae since the Eocene. The presence of Caenocholax in Colorado indicates the colonization and subse­ quent extinction of Myrmecolacidae in the Green River region (ca. 40° N). From the Late Paleocene to Early Eocene, the Earth experienced a rapid rise in global temperatures and carbon dioxide concentrations, termed the Paleocene– Eocene Thermal Maximum (PETM), in which megathermal climates extended to high lati­ tudes (Kennett and Stott 1991, Greenwood and Wing 1995, Zachos et al. 2003). Like members of several other warm‐loving animal clades, strepsipteran myrmecolacids lived in the Green River region during the warmer and wetter con­ ditions of the PETM, but became restricted to lower latitudes by modern times (Antell and Kathirithamby 2016). There are 35 described species in the genus Myrmecolax, one of which is a fossil species. They are found in Africa, Asia, Australia, and the New World. Males parasitize Formicidae (Ecitoninae, Formicinae, Myrmeciinae, Poneri­ nae, and Pseudomyrmecinae), and females are parasitic in Mantodea (Mantidae). Only one species, Myrmecolax incautus, has been unequivocally matched; the males parasitize ponerine ants, and females parasitize mantids (Kathirithamby et al. 2009a). There are 49 described species in the genus Stichotrema, two of which are fossils and six of which have been described solely from females that are parasitoids of Orthoptera (Gryllidae, Tettigoniidae) and Mantodea (Mantidae)

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(Hofeneder 1910, 1920; Hirashima and Kifune 1974; Kifune 1983; Kathirithamby et  al. 2001; J. Kathirithamby, unpublished data). The distri­ bution of Stichotrema is largely in Africa, Asia, Australia, the New World, and the Pacific Islands. The known hosts of the males are Formicinae and Myrmeciinae (Hymenoptera) (Luna de Carvalho 1972, Kathirithamby 1991b). The only known parthenogen in the Myrme­ colacidae is Stichotrema dallatorreanum from Papua New Guinea, which parasitizes the long‐ horned grasshopper Segestidea novaeguineae, a severe pest of oil palm. After studies on the biol­ ogy of the host–parasitoid relationship (Solulu 1996, Kathirithamby 1998, Kathiri­thamby et al. 1998, Solulu et al. 1998), S. dallatorreanum is now used as a biocontrol agent in  West New Britain Province, Papua New Guinea. Given the dual host nature of the sexes, Myrmecolacidae might not be effective biocon­ trol agents. For example, Caenocholax fenyesi texensis (Kathirithamby and Johnston 2004) is a parasitoid of the red imported fire ant Solenopsis invicta, a nuisance pest of humans and wildlife, in the southern United States (Kathirithamby and Johnston 1992). In the same area, Caeno­ cholax fenyesi sensu lato parasitizes the ant Crematogaster laeviuscula (Cook 2009). The taxonomic status of C. fenyesi sensu lato has not yet been determined, but if it is C. fenyesi texen­ sis, it would not be an effective biocontrol agent for the red imported fire ant because of its generalist nature. Moreover, two species of Caenocholax parasitize S. invicta in Argentina (Hayward et al. 2011). 22.4.2  Family Lychnocolacidae

The only genus in this family is Lychnocolax, which historically was placed in the family Myrmecolacidae (Kinzelbach 1971a, 1971b; Kathirithamby 1989a). But a more recent molec­ ular phylogenetic analysis placed it as a  sister group to the Stylopidae, Xenidae, Elenchidae, and Halictophagidae (McMahon et  al. 2011). There are no host records or females known for

this family, which has 23 species. Lynchnocolax is found in the circumtropical, Australian, and Neotropical Regions, but is absent from the Palearctic. However, one species, Lychnocolax hispanicus, which was described from southern Spain (Kathirithamby and Kifune 1991), was found in a light trap, possibly having been blown in by winds from North Africa. 22.4.3  Family Stylopidae

The Stylopidae, which are parasitoids of bees, have the largest number of described strep­ sipteran species (165) of any family. There are nine genera: Crawfordia, Eurystylops, Halicto­ xenos, Hylecthrus, Jantarostylops, Kinzelbachus, Melittostylops, Rozenia, and Stylops. The largest genus in the family is Stylops with 117 species that are restricted to the Holarctic Region, most being associated with closely related subgenera of solitary bees of the genus Andrena. There are 23 described species of Halictoxenos, which par­ asitize Halictidiae, five described species of Eurystylops, which also parasitize Halictidae, and three described species of Hylecthrus, which parasitize Colletidae. Halictoxenos is cosmopoli­ tan in distribution. The three described species (females) of Rozenia are from South America. The two monotypic genera are Kinzelbachus (from Hungary), whose host is the andrenid bee Melitturga clavicornis, and Melittostylops (from New Mexico), whose hosts are Hesperapis rho­ docerata and Hesperapis leucura. The males of both genera are unknown. Jantarostylops is a fossil genus from Baltic amber (Table 22.3). The phenomenon of early emergence of sty­ lopized Hymenoptera from cells in the spring was first reported by Smith (1850) and later by Pierce (1909). Linsley and MacSwain (1957) observed that stylopized Andrena complexa and Anadrena suavis emerge earlier than unsty­ lopized hosts, whereas Straka et al. (2011) noted that stylopized females of Andrena minutula, Andrena stromella, and Andrena vaga emerge at the same time as unstylopized male bees, unlike unstylopized females, which emerge

22  Biodiversity of Strepsiptera

later. When stylopized Andrena emerge from the cells in the spring, the male cephalothecae and female cephalothoraces of Stylops are already extruded. Soon after the stylopized host Andrena has left the cell, the free‐living male of Stylops emerges from the puparium, and fertili­ zation with an endoparasitic female Stylops occurs almost immediately thereafter. The early emergence of stylopized females of Andrena allows the quiescent, fertilized female strep­ sipteran Stylops sufficient time for development of the embryos to the motile planidia stage. The planidia are then released at the time when the next generation of larvae or eggs of the Andrena host are in the cells. Mating and development of embryos of the strepsipterans take place after the stylopized host has left the cell; the strepsipteran planidia are like­ wise released outside the cell (mainly on flowers). For infections to occur, the planidia need to be transported to cells where the host larvae or eggs are present. This means of transport, phoresy, is the usual mechanism employed by Strepsiptera parasitizing hosts that largely have nests with closed cells, such as solitary bees. The planidia “hitchhike” on a foraging host bee (usually unsty­ lopized) that is provisioning a cell (Kinzelbach 1971b, Kathirithamby et al. 2012a). But in some non‐social species, such as Andrena parasitized by Stylops (Linsley and MacSwain 1957) and Lasioglossum parasitized by Halictoxenos (Batra 1965), phoresy does not involve hitchhiking on the exterior of the bee. Instead, the planidia are ingested, together with the honey, into the crop of the host bee. On reaching the cell, the bee regur­ gitates the honey onto the pollen ball, along with the planidia (Linsley and MacSwain 1957). The planidia go on to parasitize the egg or larva of the host bee in the cell. 22.4.4  Family Xenidae

A molecular phylogenetic analysis of Strep­ siptera showed the Xenidae to be a sister group of the Stylopidae (McMahon et al. 2011), as sug­ gested by Kinzelbach (1971b). The family

Xenidae is one of the species‐rich groups of Strepsiptera, with a total of 117 described species of four genera: Xenos, Paraxenos, Para­ gio­xenos, and Pseudoxenos. Xenos (Fig.  22.2b), with 37 described species, is the dominant genus in the Neotropical and Palearctic Regions, but is poorly represented in Australia. Paraxenos has 44 described species that parasitize the Sphecidae and are of cosmopolitan distribution. There are 35 species of Pseudoxenos, also of cosmopolitan distribution, and only one species of Paragioxenos, described from Australia. Pseudoxenos and Paragioxenos parasitize the Vespidae (Table 22.3). In the tropics, there can be overlapping genera­ tions of Xenos, as observed in Mexico, where females of Xenos hamiltoni parasitizing Polistes carnifex have been found producing planidia at the same time as free‐living adult males were emerging from endoparasitic puparia. An indi­ vidual of Polistes carnifex was stylopized by nine male pupae, of which seven males emerged and two died with the host (Kathirithamby and Hughes 2006). This is a high rate of superparasitism in a single host, and also a high rate of successful emer­ gence of males from a single host, a record for Strepsiptera. Planidia generally seek out host larvae, but parasitization of host eggs has also been reported in Pseudoxenos carnifex, Pseudoxenos hookeri, Pseudoxenos iwatai, Stylops pacificus, and Xenos vesparum (Krombein 1967, Maeta et  al. 2001). However, parasitization of eggs of Polistes by X.  vesparum is unusual (Hughes et  al. 2003a), because oophagy occurs in this host genus (Karsai et al. 1996). But in this case, because the larval hosts in the nests were highly parasitized, the planidia might have avoided parasitizing the already‐stylopized larval hosts. It is not known whether those planidia that parasitize eggs com­ plete their life cycle (Hughes et al. 2003a). In social and solitary wasps, the infection of hosts by free‐living planidia does not take place in the nest. For instance, in the eusocial wasp Polistes, stylopized wasps desert the nest before the extrusion of the male cephalotheca and

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female cephalothorax (Hughes et al. 2004), with the result that the eclosion of the male, mating, and birth of the planidia take place outside the nest. When the planidia are released, two pos­ sible modes of transport enable the planidia to reach a nest where the next generation of host larvae or eggs can be found. Phoresy is presumed to be the usual mecha­ nism of transport used by Strepsiptera para­ sitizing hosts that have nests with closed cells, such as solitary wasps and bees, as reported for Anterhynchium and Rhynchium parasitized by Pseudoxenos (Maeta et al. 2001). Phoresy is also occasionally observed in wasps that have nests with open cells, such as in the vespid Polistes parasitized by Xenos (Hughes et  al. 2003a). Planidia are released onto flowers by a sty­ lopized wasp or bee and later, when a foraging wasp or bee visits the flower, the planidia attach themselves to the visitor and hitchhike back to the nest or cell (Kinzelbach 1971a, Kathirithamby 2012a). Phoresy has been recorded between the soli­ tary hunting thread‐waisted wasp Ammophila sp. and the strepsipteran Paraxenos. The elabo­ rate nest building, cleaning, provisioning behav­ ior, and final sealing of the cell by females of Ammophila prevent most parasites and parasi­ toids from entering the nest or cell. The strep­ sipteran genus Paraxenos is an exception to this rule; planidia are able to enter the cell unde­ tected via phoresy, in spite of the scrupulous nest building and cleaning habits of Ammophila. Phoresy by the planadia of Paraxenos is sexdependent; a great proportion of unstylopized females of Ammophila building and provi­ sioning nests have been observed with plan­ idia  sitting on the wings, legs, and petiole (Kathirithamby et al. 2012a). The second mode of transport involves direct release of the planidia onto or near a nest, as observed in the family Xenidae. A stylopized host with a gravid female strepsipteran periodi­ cally visits a non‐home nest (with open, multi­ ple combs). Hughes et al. (2003a) reported that stylopized wasps have been seen sitting near the nests of Polistes carnifex and Polistes gallicus,

with high parasite loads, suggesting that a direct release of planidia might have taken place. When the stylopized wasp moved from nest to nest, the planidia would emerge, enter the open cells, and parasitize the host larvae or eggs. This mechanism was suggested to be universal in eusocial wasps (Hughes et  al. 2003a). Direct release of planidia has been observed only in Xenidae that have nests with open cells. Such a method of transfer of planidia would not be suc­ cessful in nests with closed cells, because there is only a small window of opportunity for the transfer of planidia to take place – during build­ ing and the provisioning of food – when the nest or cell is open. Superparasitism has been observed in Vespa (Kifune and Maeta 1985, Makino and Yamashita 1998), Provespa (Kifune 1986), and Polistes (Hughes et al. 2003a). The low parasitism rates in Polistes stabilis (as opposed to Polistes car­ nifex) might be due to the nest‐building habits of the former, which is a widely dispersed spe­ cies in the rainforest (Hughes et al. 2003a). Xenos moutoni was thought to be a monotypic species parasitizing six species of hornets: Vespa analis, Vespa crabro, Vespa ducalis, Vespa dybowskii, Vespa mandarinia, and Vespa simil­ lima (Kifune 1992). A molecular analysis, how­ ever, suggests that there are at least two cryptic species (Isaka et  al. 2012). The results of this study were supported by Nakase and Kato (2013), who carried out a morphological and molecular analysis and found that two species of Xenos that parasitize hornets differ in their host use, with Xenos moutoni parasitic in V. crabro, V. ducalis, V. dybowskii, and V. mandarinia, and a new species, Xenos oxyodontes, parasitic only in V. analis and V. simillima. 22.4.5  Family Bohartillidae

Only one extant species, Bohartilla megalogna­ tha (Kinzelbach 1969), has been described, of which only four specimens are known: two from Honduras and one each from the Panama Canal Zone (Kathirithamby and Grimaldi 1993) and  Dominican Republic (Cook unpublished).

22  Biodiversity of Strepsiptera

Three fossil species have also been described from Dominican amber (Kathirithamby and Grimaldi 1993, Pohl and Kinzelbach 1995). Females and planidia are unknown for Bohartilla. The unique feature of this family is the first basal segment of the maxilla, which is three times longer than the short palpus and is covered with numerous microtrichia. Owing to a lack of specimens, the position of Bohartilla in the phylogeny of Strepsiptera is still unresolved. 22.4.6  Family Elenchidae

The Elenchidae include 29 described species in five genera: Colacina, Deinelenchus, Elencholax, Elenchus, and a fossil genus Protelenchus. Elen­ chidae have four antennomeres and two tar­ someres without claws. Colacina was described from a single male puparium parasitic in Epora subtilis (Hemiptera: Fulgoridae), collected by Alfred Wallace while on his adventures in Sarawak, and was placed in the family Elenchidae (Westwood 1877). The specimen is too fragile for further study and tentatively remains in the Elenchidae. Six described species of Deine­ lenchus parasitize Hemiptera (Dictyopharidae, Eurybrachidae, and Flatidae) in Australia, New  Guinea, and Africa. No hosts or females are known for the two described species of Elencholax from the Philippines. Elenchus, which is of cosmopolitan distribution, has 19 described species that parasitize the Hemiptera (Delphacidae). The fossil genus Protelenchus is from Dominican amber (Table 22.3). A molecular phylogenetic analysis by McMahon et al. (2011) indicates a sister‐group relationship between the families Elenchidae and Halictophagidae, some of the hosts of which overlap. Recent molecular analysis of Elenchus parasitizing rice delphacid planthoppers col­ lected from Asia and Japan revealed three geno­ types, of which one is associated with the small brown planthopper Laodelphax striatellus in northern Japan. Two others are found in three planthopper species throughout Asia, indicat­ ing cryptic species (Matsumoto et  al. 2011). Elenchus yasumatsui (Kifune and Hirashima

1975), described from southern Asia, was considered a synonym of Elenchus japonicus (Chandra 1978, Kathirithamby 1982), but the analysis by Matsumoto et  al. (2011) indicates that it might be a separate species. Both the external and internal genitalia of hosts can be altered as a consequence of stylopi­ zation. In no other family of Strepsiptera is this more evident than when Elenchidae parasitize Delphacidae (Heteroptera); drastic alterations of the external genitalia in Delphacidae hosts occur when parasitized by Elenchus. There has been confusion relating to this feature, as the external genitalia are extremely reduced and sometimes totally absent (Esaki and Hashimoto 1931; Hassan 1939; Lindberg 1949, 1960; Baumert‐ Behrisch 1960; Raatikeinan 1966; Kathirithamby; 1978, 1979, 1982, 1998; Machita et al. 2006) (Fig. 22.5b). Such stylopized delphacids were thought to be intersexes. The internal genitalia are also reduced or lost (as in all stylopized hosts), owing to the physical presence of the strepsipteran, which occupies almost the entire internal abdominal area of the host (Kathirithamby 1978, 1979, 1982; Yoshiyuki et al. 2006). An example of multiple parasitism in Elen­ chidae, where the host Javesella dubia (Delpha­ cidae) is parasitized simultaneously by the strepsipteran Elenchus tenuicornis and the hymenopteran Dicondylus lindbergi, has been recorded in Finland (Raatikainen 1967). When this occurred, either the strepsipteran died while the hymenopteran survived, or both the strepsipteran and hymenopteran died. Three species of the Elenchidae parasitize pests of cereal crops and might act as biocontrol agents. E. japonicus and E. yasumatsui are para­ sitoids of the rice planthoppers Nilaparvata lugens, Sogatella furcifera, and L. striatellus (Delphacidae), which are vectors of viral dis­ eases in Southeast and South Asia (Chandra 1978, 1980; Kifune and Maeta 1986). The host delphacids (some of which are stylopized) migrate in the rainy season, thus spreading seri­ ous viral diseases in rice (Otuka et  al. 2010, Otuka 2013). Elenchus tenuicornis is a parasi­ toid of J. dubia (Delphacidae), which is a serious

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pest of cereals, especially oats, in Europe (Raatikainen 1967). 22.4.7  Family Halictophagidae

This family includes six subfamilies (Blatto­ deaphaginae Callipharixenidae, Corio­phaginae, Dipterophaginae, Halictophaginae, and Tri­ dactylophaginae) and seven genera (Blat­­to­ deaphagus, Callipharixenos, Corio­phagus, Dipterophagus, Halictophagus, Steno­cra­no­ philus, and Tridactylophagus), with a total of 133 described species. The family Halicto­ phagidae has seven or eight antennomeres and a three‐segmented tarsus without claws. The Halictophagidae have the widest host range of  all strepsipterans, parasitizing at least 15 families from four orders of Insecta. Blattodeaphagus species parasitize cock­ roaches (Blattellidae, Blattidae), and only females are known of two species, from Australia and Japan. Calliphari­xenos is also known only from females of three species from Cambodia, the Philippines, and Thailand, which parasitize Heteroptera (Scutel­ leridae and Cicadellidae). There are 13 described species of Coriophagus, and these have seven antennomeres with five flabellae. They parasitize Heteroptera (Coreidae, Pentatomidae) in Australia, the Solomon Islands, and Zanzibar. Dipterophagus species parasitize Diptera (Tephritidae) and are so far known only from Australia; they have six anten­ nomeres with one flabellomere arising from the fourth antennomere. Dipterophagus daci is a parasite of fruit flies (Tephritidae) in Australia, and was placed in a separate family, Dipterophagidae (Drew and Allwood 1985, Allwood and Drew 1996). However, Kathirithamby (1989a, 1992) provided evidence of several synapomorphies shared with the Halictophagidae, confirming  that the Dipterophagidae is a subfamily (Dipterophaginae) within the Halictophagidae. Halictophagus is the largest genus in the family,  with 96 described ­species that parasitize many families of Hemiptera (Aphorophoridae, Cercopidae, Cicadellidae, Delphacidae, Derb­ idae, Dictyopharidae

Eurybrachidae, Flatidae, Membracidae, Psyllidae, Tettigo­metridae, and Tropiduchidae), all of which are cosmopolitan in distribution. Halictophagus has seven antennomeres, with five flabellae (Kathirithamby 1992) (Fig. 22.1). There are 13 described species of Tridactylophagus, which parasitize Orthoptera (Tridactylidae) in Albania, Australia, China, India, Japan, Philip­pines, and Romania. There are seven antennomeres with one flabellum arising from the third antenno­ mere in Tri­dactylophagus. Five species of Stenocranophi­lus,  which have seven antenno­ meres with two to  five flabella have been described from Europe, Java, and South America. In the Palearctic Region, a proportion of cicadellids that overwinter as adults are para­ sitized by quiescent fertilized female strep­ sipterans. The following spring, the female strepsipteran releases the motile planidia at a time when the young nymphal instars of its cicadellid host have emerged. The life cycles in the summer consist of two or three relatively short, overlapping generations per year, as seen in Halictophagus silwoodensis in England, which parasitizes the heather‐feeding leafhopper, Ulopa reticulata (Waloff 1981). Superparasitism by Strepsiptera occurs more frequently than does multiple parasitism. The reason is simple: the chance encounter of indi­ vidual strepsipterans of the same species in the same habitat and host (resulting in superpara­ sitism) is far greater than the chance encounter of two different species of strepsipterans in the same habitat and host (resulting in multiple parasitism). The latter process would have dif­ fering ecological and life‐history requirements. Furthermore, Strepsiptera are usually host spe­ cific and have a narrow host range. Multiple parasitism, therefore, rarely occurs naturally, and only two examples are known where two different families of Strepsiptera par­ asitize the same host. First, this occurs in a Platybrachus sp. (Hemiptera: Eurybrachidae) parasitized by an elenchid, Deinelenchus australensis, and by a halictophagid, Halicto­ phagus tryoni, in Australia (Kathirithamby 1989b, 1992). Second, in the mango leafhopper

22  Biodiversity of Strepsiptera

Idioscopus clypealis in the Philippines, a high percentage of multiple parasitism occurs by two species in different genera of Halictophagidae – both sexes of Halictophagus fulmeki and only females of Callipharixenos philippines (which might be a parthenogen) (Kathirithamby et  al. 2012b). In the case of multiple parasitism of I.  clypealis, it is intriguing that two species belonging to separate genera stylopize the same species of host, suggesting that a host‐switching event has occurred. This possibility is sup­ ported by the fact that hosts of Halictophagus are twelve genera of Hemiptera, including Eurybrachidae, for which there is only a single record. By contrast, Deinelenchus parasitizes Fulgoroidea (Dictyopharidae, Eurybrachidae, and Flatidae) (Kathirithamby 1992), and Callipharixenos (Halictophagidae) has been recorded as parasitizing only Scutelleridae (Kinzelbach 1971a). It is unlikely that such distantly related lineages coevolved in the same host. Given the wide host range of the Halicto­ phagidae, a large number of hosts are pests of crops: 1)  Several species of Halictophagus parasitize four leafhopper species of Nephotettix, which are serious pests of rice in South and South­ east Asia and Africa, being vectors of dele­ terious plant virus diseases (Ou et  al. 1965; Chandra 1978, 1980; Way and Heong 1994); 2)  Dipterophagus daci is a parasite of the fruit fly  Bactocera (Tephritidae) in Australia, and parasitism occurs only at the adult stage of the host. This unusual infective behavior is so far  known for Strepsiptera only in this host, and might be a parasite‐induced change. Fruits (which are usually infected by larvae of Bactocera) are eaten by birds and mammals. This could cause high mortality rates of the host Bactocera (Drew 1987). Strepsipteran infection might, therefore, have changed from the usual infection at the larval stage of the host to infection at the adult host stage; 3)  The corn leafhopper Dalbulus maidis, an important virus vector of maize, is found in

areas from the southern states of the United States to Argentina, including the Caribbean Islands (Nault 1980, 1983). Studies in Mexico have shown that it is parasitized by the strepsipteran Halictophagus naulti (Kathiri­ thamby and Moya‐Raygoza 2000), although the levels of parasitism are low. The minor leafhopper pests Dalbulus elimatus and Dalbulus gelbus of corn are also occasionally parasitized by H. naulti in Mexico (Moya‐ Rygoza et al. 2004); 4)  The leafhopper Molopoterus theae is a seri­ ous pest of an indigenous tea plant, Aspalathus linearis, from which “rooibos” tea (Afrikaans for “red tea”) is produced. It is parasitized by Halictophagus calcaratus (Kathirithamby et  al. 2010). The shrub occurs naturally and is grown in the Western Cape Province of South Africa. The prevalence of H. calcaratus among the populations of its host, M. theae, can exceed 50% (L. Hatting, personal communication); 5)  The leafhopper species Idiocerus niveospar­ sus, Idiocerus atkinsoni, and Idioscopus clypealis are minor pests of mango trees in India, and are parasitized by Halictophagus indicus. Subramanium (1922) estimated that about 30% of the commonest species, I. atkin­ soni, was parasitized by H. indicus (previ­ ously identified as Pyrilloxenos compactus), but no follow‐up studies on its potential ben­ efits seem to have been undertaken; 6)  The related mango leafhopper I. clypealis, a vector of a virus disease of mangoes in the Philippines (Serrano and Palo 1933), is para­ sitized by H. fulmeki and C. philippines (Kathirithamby et al. 2012b); 7)  Proutista moesta (Hemiptera: Derbidae) is a vector of pathogenic viruses of oil palm, coconuts, and areca nuts in southern India, and is parasitized by Halictophagus palmae (Kathirithamby and Ponnamma 2000); 8)  The white leafhopper Cofana spectra in India is parasitized by the strepsipteran Halictophagus australiensis. The strep­ sipteran inhibits the fecundity of its host (Mitra et al. 2014) and causes sterility, which

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suppresses the populations of C. spectra in West Africa (Oyediran et al. 2000).

22.5 ­Conclusions The biodiversity of Strepsiptera is largely under­ estimated. One of the main reasons for this vast,  hidden diversity is the likelihood of undis­ covered and unnamed cryptic species. Studies suggest that cryptic species are common in Strepsiptera. If so, the number of Strepsiptera, as it currently stands, is an underestimate. Certain families, such as the Myrmecolacidae, Stylopidae, Elenchidae, and Halictophagidae, are more frequently collected than are others. Even among this group there are variations in the num­ bers of males and females collected. For instance, males of the Myrmecolacidae are encountered in traps, whereas females are extremely rare. Stylopized Andrena with female Stylopidae are frequently encountered in the field, whereas free‐ living males seldom are encountered. The abun­ dance of males of Caenocholax over other genera of the Myrmecolacidae is still a mystery. One pos­ sible explanation might be that, because males of Caenocholax are generalist parasitoids of ants, they are encountered in traps, whereas other myrmecolacid males might be specialist parasi­ toids of ants, like the females that parasitize crick­ ets, and occur in populations that are patchy. The long interspecific branch lengths and the low intraspecific diversities from the molecular phylogenetic analysis of Caenocholax (Hayward et al. 2011) suggest genetic bottlenecks and high extinction rates. The dispersal of Strepsiptera might play a part in their low prevalence levels. Both the planidia and adult males would be restrictive as dispersal agents; because both are short‐lived, neither would be able to travel far. Furthermore, in a new site, the male’s ability to find a conspecific female is remote. This leaves the endoparasitic gravid females with develop­ ing embryos as the best possible source for long‐distance dispersal with their hosts. On encountering a new host species, the immune avoidance system in the form of a “bag”

(Kathirithamby et al. 2003) might help the pla­ nidia to parasitize a new host species. Again, one reason for the wide host range in Strepsiptera might be the unique immune‐ avoidance system. The condition in the suborder Stylopidia, whereby males exhibit endoparasitism until the end of the pupal stage and the neotenic, pedomorphic females are obligate endopara­ sites, is not known in other insect parasitoids. Although Strepsiptera parasitize several pest species, they have not been used as biocontrol agents, because they are difficult to rear in large numbers. The exception, which has proved to be a successful biocontrol agent, is S.  dallatorreanum. Being parthenogenetic, S.  dallatorreanum can be mass‐reared, and Sexava sp. with gravid females are released in oil palm plantations for the control of the long‐ horned grasshopper in Papua New Guinea. There is an urgent need for more information on the numbers and distribution of cryptic spe­ cies of Strepsiptera. This information will also be valuable for estimates of the biodiversity of their hosts in seven orders and 34 families of Insecta. Given the present fast depletion of vari­ ous habitats, we will probably lose a large num­ ber of strepsipterans and their hosts before they are even known to us.

Acknowledgments I thank the two reviewers who made helpful suggestions on an earlier draft of the manu­ script, M. Hrabar who provided plates for Figure 22.1, and M. Hrabar and J. A. Delgado for help in preparing the figures.

­References Allwood, A. J. and R. A. L. Drew. 1996. Seasonal abundance, distribution, hosts and taxonomic placement of Dipterophagus daci Drew & Allwood (Strepsiptera: Dipterophagidae). Australian Entomologist 23: 61–72.

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Otuka, A., M. Matsumura, S. Sanada‐Morimura, H. Takeuchi, T. R. Watanabe, R. Ohtsu and H. Inoue. 2010. The 2008 overseas migration of the small brown planthopper, Laodelphax striatellus, and subsequent outbreak of rice stripe disease in western Japan. Applied Entomology and Zoology 45: 259–266. Ou, S. H., C. T. Rivear, S. J. Navaretnam and K. G. Goh. 1965. The virus nature of ‘penyakiyt merah’ disease of rice in Malaysia. Plant Disease Report 49: 778–782. Oyediran, O., A. Ndongidila and E. A. Heinrichs. 2000. Strepsipteran parasitism of white leafhoppers, Cofana spp. (Hemiptera: Cicadellidae) in lowland rice in Côte d’Ivoire. Journal of Pest Management 46: 141–147. Parker H. L. and H. D. Smith. 1933. Additional notes on the strepsipteron Exenos laboulbenei Peyerimhoff. Annals of the Entomological Society of America 26: 217–233. Parker, H. L. and H. D. Smith. 1934. Further notes on Eoxenos laboulbenei Peyerimhoff with a description of the male. Annals of the Entomological Society of America 27: 468–479. Pennacchio, F. and M. R. Strand. 2006. Evolution of developmental strategies in parasitic Hymenoptera. Annual Review of Entomology 51: 233–258. Pérez‐Lachaud, G. and J. P. Lachaud. 2014. Arboreal ant colonies as ‘hot‐points’ of cryptic diversity of myrmecophiles: the weaver ant Camponotus sp. aff. textor and its interaction network with its associates. PLoS ONE 9: e100155. Pierce, W. D. 1909. A monographic revision of the twisted winged insects comprising the order Strepsiptera Kirby. Bulletin of the United States National Museum 66: 1–232. Pohl, H. and R. G. Beutel. 2005. The phylogeny of Strepsiptera (Hexapoda). Cladistics 21: 328–374. Pohl, H. and R. G. Beutel. 2016. Kinzelbachilla ellenbergeri—a new ancestral species, genus and family of Strepsiptera (Insecta). Systematic Entomology 41: 287–297. Pohl, H. and R. K. Kinzelbach. 1995. Neufunde von Fächerflüglern aus dem Baltischen und

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Dominkanischen Bernstein (Strepsiptera: Bohartillidae & Myrmecolacidae). Mitteilungen aus dem Geologisch‐Paläontologischen Institut der Universität, Hamburg 78: 97–209. Pohl, H. and R. K. Kinzelbach. 2001. First record of a female stylopid (Strepsiptera: ?Myrmecolacidae) parasite of a prionomyrmecine ant (Hymenoptera: Formicidae) in Baltic amber. Insect Systematics and Evoltuion 32: 143–146. Pohl, H., A. Katheh‐Bader and W. Schneider. 1996. Description of a new genus and two new species of Corioxenidae from Jordan (Insecta: Strepsiptera). Zoology of the Middle East 13: 107–119. Pohl, H., R. G. Beutel and R. K. Kinzelbach. 2005. Protoxenidae fam. nov. (Insecta, Strepsiptera) from Baltic amber—a ‘missing link’ in strepsipteran phylogeny. Zoologica Scripta 34: 67–69. Raatikainen, M. 1966. The effect of different sexes of the parasite Elenchus tenuicornis (Kirby) on the morphology of the adult Javesella pellucida (F.) (Hom., Delphacidae) Suomen Hyönteistieteellinen Aikakauskirja [Annales Entomologici Fennici] 32: 138–146. Raatikainen, M. 1967. Bionomics, enemies and population dynamics of Javesella pellucida (F.) (Hom., Delphacidae). Annales Agriculturae Fenniae 6: 1–49. Serrano, F. B. and M. A. Palo. 1933. Blossom blight of mangoes in the Philippines. Philippine Journal of Science 1: 211–277. Silvestri, F. 1940. Descrizione preliminare di una specie nuova di “Mengenilla” (M. spinulosa, Insecta, Strepsiptera) della Sicilia e notizie sul suo ciclo e sul particolare modo di fecondazione. Atti dell’Accademia d’Italia Rendiconti delle Classe di Scienze Fisiche, Matematiche e Naturali 7: 614–618. Silvestri, F. 1941. Studi sugli “Strepsiptera” (Insecta) I. Redescrizione e ciclo dell’ Eoxenos laboulbenei Peyerimhoff. Bollettino del Laboratorio di Zoologia Generale e della Agraria della Facoltà Agraria in Portici 31: 311–341.

Silvestri, F. 1943. Studi sugli ‘Strepsiptera’ (Insecta). III. Descrizione e biologia di 6 specie italiane di Mengenilla. Bollettino del Laboratorio di Zoologia Generale e della Agraria della Facoltà Agraria in Portici 32: 197–282. Smith, F. 1850. Observations on the Stylopites and their affinites. Zoologist 8: 2826–2829. Smith, G. and A. H. Hamm. 1914. Studies in the experimental analysis of sex. Part II. On Stylops and stylopisation. Quarterly Journal of Microscopical Science 60: 435–461. Solulu, T. M. 1996. Influence of Stichotrema dallatorreanum Hofeneder (Strepsiptera: Myrmecolacidae) on the performance of Segestidea novaeguineae (Brancsik) (Orthoptera: Tettigoniidae) in Papua New Guinea. M.Sc. thesis, University of Oxford, Oxford, UK. 75 pp. Solulu, T., S. J. Simpson and J. Kathirithamby. 1998. The effect of strepsipteran parasitism on a tettigoniid pest of oil palm in Papua New Guinea. Physiological Entomology 23: 388–398. Strand, M. R. 1986. The physiological basis of parasitoids with their hosts and their influence on reproductive strategies. Pp. 97–136. In G. Wagge and D. Greathead (eds). Insect Parasitoids. Academic Press, London, UK. Strand, M. R. and L. L. Peach. 1995. Immunological basis for compatability in parasitoid‐host relationships. Annual Review of Entomology 40: 31–56. Straka, J., K. Rezkova, J. Batelka and L. Kratochvíl. 2011. Early nest emergence of females parasitised by Strepsiptera in protandrous bees (Hymenoptera: Andrenidae). Ethology, Ecology & Evolution 23: 97–109. Subramanium, T. V. 1922. Some natural enemies of mango leafhoppers (Indiocerus spp.) in India. Bulletin of Entomological Research 12: 465–467. Tolasch, T., S. Kehl and S. Dötterl. 2012. First sex pheromone of the order Strepsiptera: (3R,5R,9R)‐3,5,9‐trimethyldodecanal in Stylops melittae Kirby, 1802. Journal of Chemical Ecology 38: 1493–1503.

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Waloff, N. 1981. The life history and descriptions of Halictophagus silwoodensis sp. n. (Strepsiptera) and its host Ulopa reticulata (Cicadellidae) in Britain. Systematic Entomology 6: 103–113. Walter, G. H. 1983. ‘Divergent host ontogenies’ in Aphelinidae (Hymenoptera: Chalcidoidea): a simplified classification and a suggested evolutionary sequence. Biological Journal of the Linnean Society 19: 63–82. Wang, B., J. Kathirithamby and M. S. Engel. 2016. The first twisted‐wing parasitoid in Eocene amber from north‐eastern China (Strepsiptera: Myrmecolacidae). Journal of Natural History 50: 1305–1313. Way, M. J. and K. L. Heong. 1994. The role of biodiversity in the dynamics and management of insect pests of tropical irrigated rice—a review. Bulletin of Entomological Research 84: 567–587. Westwood, J. O. 1861. Notice on the occurrence of a strepsipterous insect parasitic in ants discovered in Ceylon by Herr Nietner.

Transactions of the Entomological Society of London 5: 418–420. Westwood, J. O. 1877. Notes upon a Strepsipterous insect parasitic in an exotic species of Homoptera. Transactions of the Royal Entomological Society of London 1877: 185–187. Yoshiyuki, M., Y. Maeta and K. Kitamura. 2006. Effect of the stylopization by Elenchus japonicas (Easki et Hashimoto) (Strepsiptera, Elenchidae) on its host, Sogatella furcifera (Horváth) (Homoptera, Delphacidae). Chugoku Kontyû 20: 19–27. Zachos, J. C., M. W. Wara, S. Bohaty, M. L. Delaney, M. R. Petrizzo, A. Brill, T. J. Bralower and I. Premoli‐Silva. 2003. A transient rise in tropical sea surface temperature during the Palaeocene‐Eocene Thermal Maximum. Science 302: 1551–1554. Zhai, H., M. Hrabar, R. Gries, G. Gries and, R. Britton. 2016. Total synthesis, stereochemical assignment, and field‐testing of the sex pheromone of the strepsipteran Xenos peckii. Chemistry–a European Journal 22: 6190–6193.

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23 Biodiversity of Mecoptera Wesley J. Bicha Oliver Springs, Tennessee, USA

Mecoptera are a small, relict order of holome­ tabolous insects with likely origins in the Lower Permian (Novokshonov 2004). Mecoptera inhabited almost all the continents during the Permian and continued to be one of the most abundant groups of insects until the Cretaceous, when they declined in abundance to the recent level (Bashkuev 2010). The order is peculiar for this reason, with seven of its nine extant families having one to a score of species, with the remaining two families – one with essentially a Southern Hemisphere distribution (and possi­ ble Gondwanan origin) and the other with mostly a Northern Hemisphere distribution (and possible Laurasian origin)  –  containing 91% of the species. Extinct, long‐proboscid Mecoptera feeding on gymnosperm pollen might have served as early plant pollinators during the Late Jurassic to Early Cretaceous (Ollerton and Coulthard 2009, Ren et al. 2009, Labandeira 2010). Changes in climate, accompanied by basic changes in plants and the development and radiation of modern insects, with which they had to com­ pete, likely accounted for the decline of the order. Fossil mecopterans were three times more diverse at the genus level, with at least 98 genera in 34 extinct families, and were richer in comparison with recent forms than were any other insect order, with the exception of the

Blattodea (Ren and Shih 2005). Today, there are  only approximately 737 extant species of Mecoptera recognized by regional experts, dis­ tributed among 39 genera and nine families, some with ancient ties (Table 23.1). Mecoptera are generally called “scorpionflies” because the males of some families have bul­ bous genitalia on an upturned abdominal tip, which resembles a scorpion’s stinger. Scorpion­ flies are moderate‐sized insects that can be fairly abundant in moist forest ecotone, but usually are not noticed by the general public. However, social networks recently were causing alarm among people with stories of “flying scor­ pions” in Sonora, Mexico. Five attacks by flying scorpions were reported on social networks, and some media outlets were talking about pos­ sible apocalyptic signs. After several people were driven to panic, a spokesman of the Red Cross had to explain that this insect posed no danger to humans (Hugo 2014). Scorpionflies are characterized by the adults having the labrum and clypeus combined to pro­ duce a rostrum with chewing mouthparts at the apex, filiform antennae, and three ocelli (except in Apteropanorpidae and Meropeidae). The mecopteran thorax has a transverse prothorax; large and similar mesothorax and metathorax; a meron between the coxae of the middle and hind legs; three similar pairs of thin, long legs; and

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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Table 23.1  Number of extant genera and species of Mecoptera by family. Family

Genera

Species

Apteropanorpidae

1

4

Bittacidae

18

214

Boreidae

3

30

Choristidae

3

8

Eomeropidae

1

1

Meropeidae

2

3

Nannochoristidae

1

7

Panorpidae

7

457

Panorpodidae

2

13

Total

38

737

fore‐ and hindwings of similar size with simple venation and a frenulate coupling device (except in the Bittacidae). Mecopteran male genitalia are of two different structural forms, the bittacid form, similar to certain primitive Diptera, and the form of the remaining families, with the gen­ italia consolidated into a genitalic bulb. The mecopteran digestive system has a muscular pump and well‐developed salivary glands. This condition is especially true in the Panorpidae, which exhibit considerable sexual dimorphism, with the salivary glands sometimes extending the length of the male abdomen (Ma et al. 2011). Most mecopteran larvae have compound eyes like those of hemimetabolous larvae. Boreid lar­ vae appear to have stemmata, and eyes are absent in panorpodid larvae (Chen et  al. 2012). The sperm pumps of the Antliophora and Strepsiptera are  not homologous (the sperm pump of Strepsiptera lacks any sclerotized parts), sug­ gesting that they evolved independently (Hünefeld and Beutel 2005). The larvae retain other primitive characters, e.g., abdominal legs on every segment and four caudal adhesive lobes. The caudal lobes are modified into a pair of anal hooks in Nannochoristidae and absent in boreines. Mecopteran pupae are exarate and decticous. The mecopteran diet is diverse:

adult  bittacids and nannochoristid l­arvae are predaceous; panorpids, choristids, meropeids, ­ and nannothamids are sapro­ phagous; and boreids and panorpodids are phytophagous.

23.1 ­Suborder Nannomecoptera The suborder Nannomecoptera lacks a sperm pump. Among the mecopteran families, this primitive state is only retained in the Nanno­choristidae. 23.1.1  Family Nannochoristidae

The Nannochoristidae are a small, relict family with only seven extant species (Byers 1989, Penny 2016) in one genus, yet the Nanno­choristidae fossil record is well documented by  hundreds of specimens and 16 species from  numerous Jurassic and Lower Cretaceous deposits in Australia, China, Kazakhstan, Mongolia, and Siberia, suggesting a mid‐Mesozoic zenith (Cao et al. 2015). The Nannochoristidae are the most archaic extant mecopteroid family and likely separated from the remainder of the Mecoptera in the Upper Permian (Tillyard 1935). The Mecoptera in general, but particularly nan­ nochoristids, are regarded as having great phylo­ genetic importance. The exact systematic position of the Nannochoristidae in the Antliophora is uncertain. Willmann (1987), com­ paring external and internal morphological details, proposed that all extant Mecoptera fami­ lies with males possessing a sperm pump be placed in the suborder Pistillifera, leaving the Nannochoristidae alone in the suborder Nannomecoptera. The larval legs of nanno­ choristids were considered to be the most primi­ tive among Mecoptera, retaining a two‐segmented tarsus and a claw (Pilgrim 1972), but this was shown to be incorrect by Kluge (2003), and that the leg was not so primitive. Whiting (2002), ana­ lyzing four genes, suggested that Mecoptera are paraphyletic, having two major lineages, with the  Nannochoristidae plus Boreidae and

23  Biodiversity of Mecoptera

living on the South Island of New Zealand was assigned to Microchorista (Riek 1954; Byers 1973, 1989), but the genus was synonymized with Nannochorista by Kristensen (1989).

23.2 ­Suborder Pistillifera

Figure 23.1  Nannochorista andica (image by W. Bicha).

Siphonaptera being basal to the remainder of the extant Mecoptera. Alter­ natively, Beutel and Baum (2008) suggested that the Nannochoristidae are a sister group to the Diptera. Nannochoristids are small insects with four equal‐sized wings held roof‐like over the abdo­ men when at rest (Fig. 23.1). The male genitalia are unique among the Mecoptera, with the basi­ styles broadly fused dorsally and ventrally. The males lack a distinct sperm pump (Willmann 1981). The labial structure of nannochoristids is unique among Mecoptera and remarkably like that of some nematocerous Diptera, particularly the expanded apical segments of the labial palps (Byers 1989). The extant species of Nannochoristidae have  a Gondwanaland, Southern Hemisphere distribution, being found thus far in Argentina, Chile, New Zealand, Tasmania, and southeast Australia, making the insects of particular inter­ est to biogeographers. Nannochoristid larvae live in springs and small, flowing streams, and are predaceous on aquatic Diptera (Pilgrim 1972, Byers 1989). Adults are found low on lush vegetation not far from the stream where the larvae live. Nannochoristid adults first appear in the late spring, and there is a second peak of abundance in the fall. The insects are unique among Mecoptera in having elateriform larvae (Pilgrim 1972). There are three extant, described species of Nannochorista living in Argentina and Chile and three species living in southeast­ ern Australia and Tasmania. A seventh species

The Pistillifera are a sister group to the Nannomecoptera, containing the remainder of the extant Mecoptera families, and are charac­ terized by the male possessing a sperm pump consisting of a pistillum and a pumping chamber. 23.2.1  Infraorder Raptipedia

The Raptipedia are characterized by having only one tarsal claw, which is used to capture prey. Today, the infraorder contains only one family, the Bittacidae. 23.2.1.1  Family Bittacidae

The Bittacidae are a truly cosmopolitan family with representatives in North America, South America, Africa, Australia, and from Europe across North Asia to Southeast and South Asia. The family seems to have a Gondwanan origin, being particularly diverse in Australia and South America, and comprises the entire mecopteran fauna of Africa (Grimaldi and Engel 2005). The insects are commonly called hangingflies because of their habit of hanging from the undersides of vegetation; they superficially resemble crane flies. Bittacids were placed in the infraorder Raptipedia in the suborder Pistillifera (Willmann 1987). Bittacids diverged early from the rest of the order, likely having arisen in the Jurassic (Willmann 1987). Wang et  al. (2012) hypothesized that the Middle Jurassic hangingfly Juracimbrophlebia ginkgofolia of the extinct family Cimbrophlebi­ idae had an extraordinary association with the Ginko‐like tree Yimaia capituliformis (Yimai­ aceae). They suggested the hangingfly wings resembled the leaves (leaf mimesis) of the tree to avoid predators (crypsis), or perhaps that the hangingfly provided an antiherbivore function for its plant host (mutualism).

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Bittacids are unique among the Mecoptera by having a single claw at the end of each tarsus. Bittacid adults typically hang from the under­ sides of ground vegetation by their front legs and are predaceous, using their raptorial tarsi with one tarsal claw of their hind legs to capture prey. Male bittacids are unique among the Mecoptera in possessing a penisfilum, and have genitalia structurally more similar to those of lower Diptera than do the remaining Mecoptera families, which have the genitalic structure con­ solidated into a genitalic bulb. The wings of bit­ tacids are long, narrowed basally, and generally equal‐sized. Bittacid adults are predaceous and typically live among lush, herbaceous vegetation growing in the shade of mesic forests, often along slow‐ moving streams. Yet other species prefer to live among grasses in open, apparently less mesic habitats, although often under or near shrubs or trees that provide protection from direct sun. Bittacids often coexist in the same mesic, shady habitat with other species of bittacids and one or several species of panorpids, although in mountainous areas, bittacids generally seem to prefer lower elevations than do panorpids. Bittacid females drop their eggs to the ground, where they might hatch in several weeks or dia­ pause before hatching, depending on the spe­ cies (Setty 1940, Penny 2006). The insects have one generation per year, as far as is known. The larvae are eruciform and typically develop through four instars, with a head featuring com­ pound eyes and a median ocellus, a thorax with three pairs of legs, and an abdomen with eight pairs of prolegs on the first eight segments. The larvae are similar to panorpid larvae, with each body segment having several setae that are thickened at the base. The larvae develop in the soil, likely feeding on dead insects and detritus. The fourth‐instar larva constructs a pupal cell in the soil. Bittacids have exarate, decticous pupae (Tan and Hua 2009). Bittacids are of interest to behavioral and evo­ lutionary biologists. The males provide the females with a nuptial gift of prey (Fig. 23.2) during courtship and copulation (Setty 1940,

Thornhill 1978). Males initiate olfactory calling while hanging by their forelegs from the under­ sides of vegetation. Pheromone‐dispersing glands between the sixth and seventh, and sev­ enth and eighth abdominal segments are everted from the male abdomen while calling females. After pair formation, the male and female hang from the vegetation by their front legs, facing one another. Prey is offered to the female by the male, which grasps one or more of the female’s legs while maintaining a grip on the prey. The female tastes the prey while it is held by both insects. The male then attempts to copulate with the female. During copulation, the male abdomen twists 180 degrees to accommodate the bittacid’s face‐to‐face mating position. The dististyles of male bittacids are useless to hold the female during copulation in the 180‐degree twisted mating position, and so the male epian­ drial lobes are used instead to clasp the female genitalia. This face‐to‐face mating position, which is unique among the Mecoptera, might have been adopted to maintain control of the nuptial gift. It is in the male’s interest to reuse the nuptial gift, while the female benefits by consuming the entire nuptial gift (Thornhill 1978, Gao and Hua 2013). The Bittacidae have 214 recognized extant species distributed among 18 genera, although there are several undescribed species known from Japan, and other species likely remain to be discovered in South America, Indochina, and possibly China. Bittacus is certainly paraphyl­ etic and has likely become a catch all for a num­ ber of hangingflies of unknown generic relationships. Much work remains to be done with bittacid systematics, especially with those from South America and Japan. Bittacus, as cur­ rently defined, has representatives in Europe (two species); North America (seven species in the United States and Canada, of which two spe­ cies are shared with Mexico, plus seven species endemic to Mexico (Byers 2011) and one species shared by Mexico and Central America); Africa (51 species, Londt 1994); Asia (one species endemic to Far East Russia (Plutenko 1985), 27 species endemic to China, two species shared by

23  Biodiversity of Mecoptera

Figure 23.2  A mating pair of Harpobittacus similis. The female is feeding on the arthropod nuptial gift given to her by the mating male. (Image by W. Bicha). (See color plate section for the color representation of this figure.)

China and Korea, one species shared by China, Korea, and Japan, nine species endemic to Japan, four species in Taiwan, five species endemic to Indochina (Bicha 2015), and six species in South Asia (Bicha 2011)); Central and South America (27 species, Machado et al. 2009); and Australia (one species, Lambkin 1993). Apterobittacus and Orobittacus are each monotypic and restricted to central California. Orobittacus has features in common with Anabittacus, which is also monotypic, but is endemic to Chile. Hylobittacus, also monotypic, is widespread across eastern North America and has characteristics in common with Kalobittacus from Mexico and Central America, currently with eight described species (Machado et al. 2009). Eremobittacus, with two species, is seemingly limited to xeric areas of central Mexico. Issikiella currently contains five spe­ cies  and ranges from Columbia to Bolivia. Nannobittacus, with five species, ranges from Panama to Brazil. Pazius, with eight species, ranges from Costa Rica to Brazil. Central and South America is considered home to 27 species of Bittacus (Byers 1996,

2004; Machado et al. 2009), but previous authors split the fauna between 21 species of Thyridates (Willmann 1983; Collucci and Amorim 2000, 2001) and two species of Neobittacus (Penny 1977, Penny and Byers 1979), with the remain­ der indistinctive bittacids left as Bittacus. However, recent molecular analysis suggests that South American Bittacus may be only distantly related to its North American cousin (W. J. Bicha and N. Schiff, unpublished data). Bicaubittacus, with six species, occurs in China, Taiwan, and Indochina. Terrobittacus, with six species, is endemic to China. Har­ po­bittacus is endemic to Australia and has 11  species (Lambkin 1994). Austrobittacus, Edrio­bittacus, Symbittacus, and Tytthobittacus are each endemic to Australia and monotypic. Afrobittacus, endemic to central Africa, and Anomalobittacus, endemic to South Africa, are each monotypic. 23.2.2  Infraorder Opisthogonopora

The Opisthogonopora are characterized by the female genital chamber being located

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toward the end of the ninth abdominal seg­ ment (Willmann 1987). 23.2.2.1  Group Boreomorpha

The Boreomorpha are characterized by reduced wings, sperm transfer by means of  a spermato­ phore (Willmann 1987), and scarabaeiform larvae, with stammata rather than compound eyes. The Boreomorpha contain one extant family, the Boreidae. Family Boreidae  Russell (1979) split the family Boreidae into two subfamilies, the Caurininae Russell, 1979 and the Boreinae Russell, 1979, based on the primitive characters of the Caurininae. Boreines (Palaepboreus) are known from Asia in the late Jurassic and early Cretaceous; so the family is likely to date to at least to the early Jurassic. Boreids (Fig. 23.3) are small insects with reduced or no wings, and are unique among the Mecoptera in their ability to hop on snow (it is unknown whether apteropanorpids hop). The Boreidae are one of the two mecopteran families that independently adapted to cold climates. The resilin that enables boreids to hop is secreted in a manner similar to that of the Siphonaptera. Female boreines are unique by having panoistic ovaries, another morphologi­ cal feature shared with Siphonaptera, rather

than polytrophic ovaries, as in other Mecoptera. Cerci of female boreids are modified to an elon­ gated ovipositor. The boreid head is arranged slightly differently from that of other Mecoptera. The antennae are located on the head between the compound eyes. The lateral ocelli are close to the margins of the compound eyes, and the median ocellus, if present, is just dorsal to the antennae, rather than in a triangular arrange­ ment as in other Mecoptera. Caurinines have the distinction of lacking the developed rostrum of the rest of the Mecoptera (with the exception of the nannochoristids) and the key characters of the wingless adult insect orders, and thus may not key to any particular order, using most ordinal keys. Boreid larvae are scarabaeiform and lack abdominal prolegs (Penny 1977, Russell 1979). There are currently 30 recognized species of Boreidae, although there is at least one unde­ scribed species of boreid from the northwestern United States and likely others waiting to be dis­ covered in inaccessible regions of Central Asia and Siberia. The Caurininae contain one genus, Caurinus, which currently contains two species (Sikes and Stockbridge 2013). The Boreinae contain two genera, Hesperoboreus and Boreus. Hesperoboreus is endemic to the Coast Moun­ tains of Oregon and Washington and the San  Jacinto Mountains of southern California. Figure 23.3  Male Boreus californicus on snow (image by W. Bicha).

23  Biodiversity of Mecoptera

Boreus is distributed at the higher latitudes of the Holarctic, with representatives in North America (12 species), Western and Eastern Europe (six species) (Penny 1977), Central Asia to the Russian Far East (seven species) (Plutenko 1984, 1985), and Japan (one species) (Hori and Morimoto 1996). 23.2.2.2  Group Meropomorpha

The Meropomorpha are characterized by hav­ ing broad wings with numerous cross veins, elongated basistyles and dististyles, and the male sperm‐pump piston depressors inserting dorsally rather than ventrally (Willmann 1987). The Meropomorpha contain one extant family, the Meropeidae. Family Meropeidae  The Meropeidae are cock­ roach‐like insects with broad, flattened wings that cover the body and have numerous cross veins and a pronotum that nearly covers the head. The insects have strange reniform com­ pound eyes and lack ocelli. Merope retains a six‐ branched media in the forewing. The insects are commonly called earwigflies because the males have elongated forcipate terminalia. The Meropeidae contain two extant genera, Merope, which is monotypic, and Austromerope, which contains two species. These three species of Meropeidae have an extremely disjunct distri­ bution, with Merope inhabiting the tem­perate hardwood forests of eastern North America (Byers 1973), and Austromerope living in a vari­ ety of “Mediterranean” habitats of southwestern Australia (Faithfull et  al. 1985, Abbott et  al. 2007) and the Atlantic forest of southeastern Brazil (Machado et. al. 2013). A mid‐Cretaceous species has been discovered from Himalayan Myanmar (Grimaldi and Engel 2013) and a mid‐ Jurassic species from Siberian Russia (Novokshonov 1995). This distribution suggests geographic extinction over a range that was widespread since at least the Oligocene, when Australia was still connected to South America via a vegetated Antarctica (Grimaldi and Engel 2013).

Meropeids are possibly phytophagous, but they have been trapped on carrion, suggesting that they are more likely saprophagous (Pechal et  al. 2011). The larvae of meropeids remain unknown. Although considered rare for many years, the documented range of these insects is broadening with the increased use of Malaise and pitfall traps (Byers 1973). As with notio­ thaumids, meropeids are often trapped in mature forests with trees growing on slopes with deep crevices in the soil at the base of trees. Given the highly flattened nature of the mero­ peids, it is possible that the reason the insects are trapped, but not seen alive, and that the lar­ vae remain undiscovered, is that they live up in the inaccessible crevices among the roots of large mature trees. Such a habitat would explain the ability of Austromerope to survive the dry southwestern Australian summer. 23.2.2.3  Group Panorpomorpha

Tergite IX of the males of the Panorpomorpha is elongated and, with segments X and XI, forms a genitalic bulb. Family Eomeropidae  The family Eomeropidae

contains one monotypic genus, Notiothauma, with a distribution limited to the southern beech forests of southern Chile, although the family was more widely distributed in the Cenozoic, with fossils recorded from northeast­ ern Asia and North America (Archi­bald et  al. 2005). Notiothauma was initially placed in the family Notiothaumidae until it was later deter­ mined to be closely related to the extinct Eomerope, of the family Eomeropidae, which predated Notiothaumidae (Ponomarenko and Rasnitsyn 1974). The insect is likely sapropha­ gous (Jara‐Soto et  al. 2007). Areas where Notiothauma have been collected contain old‐ growth forest with steep, crevice‐laden hill­ sides. The greatly flattened form suggests that Notiothauma might live in the inaccessible crevices among the roots of large mature trees. Unlike the remainder of the Chilean Mecoptera, the insect is active in the cool, rainy season.

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As for most Mecoptera families, although the Eomeropidae play a role in the intricate web of life, they do not seem to be noticeably beneficial or detrimental to the environment, nor do they appear to be sufficiently abundant to have noticeable societal impact on humans. Further discovery and study of fossil Eomeropidae might result in scientists having a better understand­ ing of Gondwanan biodiversity and the develop­ ment of early holometabolous insects. Family Choristidae  The Choristidae contain three

genera: Chorista with two species, Taeniochorista, with four species, and Neocho­rista with two species. All are endemic to the wetter areas of southeastern Australia (Riek 1973). Adults are moderate‐sized, con­spicuous insects observed on the top surfaces of vege­tation growing in moist areas, often near streams, and emerge in late summer to fall (Riek 1973). Choristids retain a five‐branched media in the forewing.

Family Apteropanorpidae  This family is endemic to the high elevations of Tasmania (Australia) and contains one genus (Aptero­panorpa) with only four species. These small, completely apterous insects are active as adults in winter, as are boreids, and yet there are no close morphological similarities. Molecular analysis suggests that the  two groups developed independently. The adults have a rostrum as large as that of panorpids, small compound eyes, and no ocelli. The female  abdomen is bulbous, whereas the male  abdomen bears genitalia consolidated in a bulb, as in panorpids. The insects are primarily saprophagous. The larvae of aptero­panorpids are eruciform, with 16 ommatidia per compound eye. The first nine abdominal segments bear a pair of small dorsal processes, and the tenth abdominal segment bears ever­sible papillae forming a suction organ (Byers and Yeates 1999). Family Panorpodidae  The earliest panorpodids

belonged to the “Orthophlebiidae,” which appeared in the Late Triassic and were last seen in the Cretaceous (Willmann 1987), to later be replaced in the Eocene. The superfamily

Panorpoidea radiated to its greatest family‐level diversity of six families in the Eocene, but four of the families were gone by the Oligocene, leaving only the Panorpidae and Panorpodidae to persist to today (Archibald et al. 2013). The Panorpoidea have a Laurasian distribution, suggesting that that they are not closely related to the Bittacidae (Grimaldi and Engel 2005). The Panorpodidae and Panorpidae possess a four‐branched media in  both the fore‐ and hindwings, as did the Austropanorpidae (extinct), whereas the Orthophlebiidae, from which they likely arose, possessed a five‐branched media in the forewing and a four‐branched media in the hindwing. The remaining three extinct panorpodid families had a five‐branched media in both the fore‐ and hind­ wings (Archibald et al. 2013). The oldest panor­ podid is from Baltic amber (Carpenter 1954). Panorpodids are restricted to the temperate climate, mid‐latitude Holarctic Region, extend­ ing into Indonesia. The Panorpodidae cur­ rently have 13 recognized, extant species in two genera, Panorpodes, with representatives in northern Asia and the United States (central California), and Brachypanorpa, with repre­ sentatives in the mountains of North America. Typical of seven of the nine mecopteran families,  the Panorpodidae have few extant species. Seven currently recognized species of Panorpodes occur in China, Japan, and Korea, and one species of Panorpodes occurs in California. Three species of Brachypanorpa are described from the Coast, Cascade, and Rocky Mountains, and two species are described from the southern Appalachians. Several panorpo­ dids in Japan are waiting to be described, and possibly several others are yet to be discovered in China. The family is in need of a worldwide revision, although new taxonomic tools will need to be developed, or extensive biological studies will need to be performed. There are relatively minor and possibly unreliable mor­ phological differences between some of the described species of Panorpodes and Brachy­ panorpa. Females of Panorpodes paradoxa have a bewildering variety of wing patterns and both long‐ and short‐winged forms, which warrants

23  Biodiversity of Mecoptera

Figure 23.4  A mating pair of Panorpodes paradoxa. The male is uppermost in the picture. (Image by W. Bicha.)

further study. A molecular phylogenetic study of the Panorpodidae concluded that the genus Panorpa and the family Panorpodidae both form monophyletic clades, that the Asian and California Panorpodes species are likely mono­ phyletic, and that the Brachypanorpa form a monophyletic group (Pollmann et al. 2008). Panorpodids are typically observed with their wings held in a “V” while resting horizontally on the top surfaces of vegetation in the broken shade of moist forest and can be quite abundant in the optimum location and season (Fig. 23.4). Females of Brachypanorpa and at least one spe­ cies of Panorpodes have wings of reduced size. These brachypterous females rest vertically on plant stems down in the vegetation and are not often seen. In the field, the insects resemble panorpids, except for having a shorter ros­ trum  and feeding on herbaceous vegetation. The male genitalia are consolidated in an up‐ curved genitalic bulb, as are the genitalia of male panorpids, giving each a scorpion‐like appearance. Panorpodids mate with their bod­ ies at a 30‐degree angle from each other, as do

panorpids. The larvae of Brachypanorpa are scarabaeiform (Byers and Thornhill 1983) and possibly feed on decaying logs. Family Panorpidae  The fossil record suggests

that panorpids were rare, compared with other Mecoptera (Ding et al. 2014). Two species of the extinct genus Jurassipanorpa are known from the middle Jurassic, one species of the extinct genus Baltipanorpa Krzeminski, 2012 is known from Eocene Baltic amber, and seven species of Panorpa are known from the Eocene and Oligocene. By contrast, today the Panorpidae are the most species‐rich mecop­teran family, with six genera and approximately 448 extant species. Panorpids are typically observed sitting on the upper surfaces of herbaceous vegetation growing in the broken shade of moist forest ecotone (Fig. 23.5). The insects are commonly called “scorpionflies” because the males have characteristic up‐curved abdominal termi­ nalia. Panorpids are similar to panorpodids except they are saprophagous (panorpodids are

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Figure 23.5  Male Panorpa coreana (image by Michael J. Plaster).

phytophagous), have eruciform larvae rather than scarabaeiform larvae, and are much more species‐rich. Panorpids feed on soft‐bodied arthropods that fall to the tops of ground veg­ etation or the forest floor, but are also klep­ toparasites of the prey of web‐building spiders, entering webs and feeding on trapped prey (Thornhill 1975). Although panorpids are not known to be pollinators, they have been reported to feed on pollen and have been col­ lected from flowers in treetops in Taiwan (N. D. Penny, personal communication). Panorpids have a longer rostrum than do panorpodids, toothed tarsal claws, and a longer, thinner abdomen. Panorpa, with approximately 250 currently recognized species, is restricted to the temper­ ate climate, mid‐latitudes of the Holarctic Region, extending south into Mexico at the higher elevations (Bicha 2006). Molecular anal­ ysis has revealed that Panorpa is paraphyletic (Whiting 2002) and is likely a catch‐all genus.

The genus is in need of a world‐view revision, and conceivably the genus might be further split into a number of additional genera. Panorpa occurs in temperate North America, with 55 species from southeastern Canada and the eastern United States, and 33 species from the mountainous regions of Mexico (Byers 2011, 2013). The genus also occurs from the United Kingdom to central Siberia and south to the Caucasus, with 24 species (L. Dvorak, personal communication), which seem to be unrelated to North American or Asian species. Northeast Asia is well represented with Panorpa, with one species endemic to Far East Russia, 68 species endemic to China, one species shared by China and India, one species shared by China and Korea, one species shared by China, Korea, and the Russian Far East, five species endemic to Korea, three species shared by the Russian Far East and Korea, 27 species endemic to Japan, one species shared by Japan and China, 29 spe­ cies endemic to Taiwan, one species in northern

23  Biodiversity of Mecoptera

Burma, and one species in India. Panorpa has been well collected, with the possible exception of China, and the number of species probably will be slightly reduced on revision. The contin­ ued existence of many Mexican species might be in peril because of extensive cultivation and land development. Neopanorpa has 161 currently recognized spe­ cies, although a few species probably remain to be discovered in China and Indochina. This Oriental genus is found across Bhutan, India, and Nepal (21 species; Rust and Byers 1976), Indochina (40 species; Bicha 2015), southern China (74 species; B. Hua, personal communication), and Indonesia (Sumatra has five species, Borneo has three spe­ cies, and Java has six species; Chau and Byers 1978). Neopanorpa, and Mecoptera of any sort, are seemingly absent from the Philippines, but are well represented on Taiwan with 10 species (Byers 1994, 2002, 2008) and the Japanese Ryukyu Islands with one species (T. Nakamura, personal communication). One species is shared by China and Taiwan. Cerapanorpa with 22 species, has 16 species in China, 4 species in Japan, one species in Taiwan, and one species shared by Russian and Korea. Leptopanorpa, with 12 species, is restricted to the higher elevations of Sumatra and Java (Chau and Byers 1978), and in Java this genus is under serious threat due to extensive cultivation and development. Furcatopanorpa is monotypic, Sinopanorpa contains three species, and Dicerapanorpa has eight species; all occur in China. Male panorpids exhibit any of three mating tactics. A male may locate a dead insect or other soft‐bodied arthropod, defend it from other males, and offer it to a female while copulating. Alternatively, a male may exude a salivary mass or pellet on a leaf and stand by it as he disperses a pheromone (Thornhill 1973, 1981; Kock et  al. 2007) and vibrates his abdomen and wings to attract a female. The female may feed on the sali­ vary secretion while copulating with the male (Byers and Thornhill 1983). Male panorpids have extraordinary salivary glands of varying sizes among species, which extend into the abdomen (Liu and Hua 2010, Ma and Hua 2011). Third, a

male may grasp a female with its abdominal for­ ceps (dististyles) and use a ­clasping organ on the third and fourth tergum to clasp the leading edge of the female wings to secure the female during copulation, with the two at an approximate 30‐ degree angle to each other. Courtship and mating of most species occurs at dusk, dawn, and night. During courtship, males disperse a sex phero­ mone attractive to females of their species from a gland in an eversible pouch between their basi­ styles (Thornhill 1973). The specific chemical identity of the sex pheromone of one European species has recently been identified and is a mix­ ture of two aldehydes (Kock et al. 2007).

23.3 ­Societal Value of Mecoptera Apteropanorpids, boreids, eomeropids, mero­ peids, and nannochoristids are somewhat rare insects, living in a restricted habitat and limited in distribution, and so have little potential to be of obvious economic or societal value. As Penny (2016) has written, “Mecoptera are most often defined by the characters they do not possess.” The most remarkable thing about Mecoptera is that there is nothing remarkable about them. Mecoptera have a generalized holometabolous life history without specialized intermedi­ ate  forms such as in Strepsiptera or parasitic Hymenoptera. The insects seem to have rela­ tively few morphological adaptations that would allow them to exploit the many niches of modern plant or animal life as do Diptera, Hymenoptera, Lepidoptera, or Siphonaptera. No Mecoptera developed social behavior beyond that associ­ ated with mating. The insects are not crop pests, and they do not bite or carry disease agents of humans or livestock. They do not pollinate crops or provide a foodstuff. They do not parasitize or hold in check undesirable organisms. They do, however, play an essential role in the ecosystem as recyclers in the intricate web of life. In contrast to other families of Mecoptera, hangingflies are cosmopolitan, more tolerant of diverse habitat, and often reach high abundance. Because the insects are predaceous on Diptera,

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small lepidopterous larvae and adults, Hemip­ tera, and Hymenoptera (Thornhill 1978), they are the one group of Mecoptera, more than any other, that are likely to make an unnoticed posi­ tive societal contribution by keeping potentially destructive insect populations in check. Panorpids can be quite numerous in their peak flight season and likely play a significant role in recycling carrion. In an interesting twist, panorpids might be of value to forensic ento­ mologists. Panorpids have arrived at a human cadaver within less than 20 minutes and out­ numbered all other insects, including blow flies, for the first 24 hours; they continued to feed until the cadaver desiccated a week later (Lindgren et al. 2015).

23.4 ­Scientific Value of Mecoptera Studies of fossil Mecoptera have been of great value in developing an understanding of the ori­ gins and development of the holometabolous insect orders. Eomeropids, meropeids, and nan­ nochoristids appear little changed from antiq­ uity and serve as living subjects for further scientific investigation into the origin and bio­ geography of the holometabolous insect orders (Kristensen 1999). Likewise, the close relation­ ship of the Mecoptera with the Siphonaptera is of particular evolutionary interest to scientists (Whiting 2002, Grimaldi and Engel 2005, Trautwein et al. 2012). Nannochoristids would be excellent candi­ dates as key indicators of small‐stream health in countries where they exist. Boreids and aptero­ panorpids, with their specific requirements for stable boreal climate and limited ability to dis­ perse, would likely be excellent candidate “canaries” of climate change. The complex and interesting courtship and mating behavior of hangingflies and scorpion­ flies, involving male attractant pheromones, abdominal vibrations, nuptial gift offerings of food items including salivary secretions, and forced copulation have been the subject of numerous studies on mate choice, mating

strategy, male resource investment, rape, and other topics (Thornhill 1980a, 1980b, 1981, 1984; Thornhill and Alcock 1983).

23.5 ­Conclusion The small, evolutionarily ancient order Mecop­ tera has much to teach scientists about the early development and dispersion of the Holo­ metabola, insect mating systems, and environ­ mental change. But because of its relative scarcity, it is unlikely to have noticeable, signifi­ cant societal impact compared with most other insect orders.

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Ollerton, J. and E. Coulthard. 2009. Evolution of animal pollination. Science 326: 808–809. Pechal, J. L., M. E. Benbow and J. K. Tomberlin. 2011. Merope tuber Newman (Mecoptera: Meropeidae) collected in association with carrion in Greene County, Ohio, USA: an infrequent collection of an elusive species. American Midland Naturalist 166:453–457. Penny, N. D. 1977. Two new species of Bittacidae (Mecoptera) from Amazon Basin. Acta Amazonica 7: 423–427. Penny, N. D. 2006. A review of our knowledge of California Mecoptera. Proceedings of the California Academy of Sciences 57: 365–372. Penny, N. D. 2016. World Checklist of Extant Mecoptera Species. http://researcharchive. calacademy.org/research/entomology/ Entomology_Resources/mecoptera/ [Accessed 2 March 2016]. Penny, N. D. and G. W. Byers. 1979. Chave para as familias e generos da Mecoptera (Insecta) da America, do sul dos Estados Unidos. Acta Amazonica 9: 363–364. Pilgrim, R. L. C. 1972. The aquatic larva and pupa of Choristella philpotti Tillyard, 1917 (Mecoptera: Nannochoristidae). Pacific Insects 14: 151–168. Plutenko, A. V. 1984. A new species of the genus Boreus (Mecoptera, Boreidae) from the Soviet Far East. Zoologicheskii Zhurnal 63: 778–781. Plutenko, A. V. 1985. New and little known species of Mecoptera from the Soviet Far East. Entomologicheskoe Obozrenie 64: 171–176. Pollmann, C., B. Misof and K. P. Sauer. 2008. Molecular phylogeny of panorpodid scorpionflies: an enigmatic, species‐poor family of Mecoptera (Insecta). Organisms Diversity and Evolution 8: 77–83. Ponomarenko, A. G. and A. P. Rasnitsyn. 1974. New Mesozoic and Cenozoic Protomecoptera. Paleontology Journal 4: 493–507. Ren, D., C. C. Labandeira, J. A. Santiago‐Blay, A. Rasnitsyn, C. K. Shih, A. Bashkuev, M. A. Logan, C. L. Hotton and D. Dilcher. 2009. A probable pollination mode before Angiosperms: Eurasian, long‐proboscid scorpionflies. Science 326: 840–847.

Ren, D. and C. K. Shih. 2005. The first discovery of fossil eomeropids from China (Insecta, Mecoptera). Acta Zootaxonomica Sinica 30: 275–280. Riek, E. F. 1954. The Australian Mecoptera or scorpionflies. Australian Journal of Zoology 2: 143–168. Riek, E. F. 1973. A revision of Australian scorpionflies of the family Choristidae (Mecoptera). Journal of the Australian Entomological Society 12: 103–112. Russell, L. K. 1979. A new genus and a new species of Boreidae from Oregon (Mecoptera). Proceedings of the Entomological Society of Washington 81: 22–31. Rust, M. K. and G. W. Byers. 1976. The Mecoptera of India and adjacent regions. University of Kansas Science Bulletin 51: 19–90. Setty, L. R. 1940. Biology and morphology of North American Bittacidae (order Mecoptera). American Midland Naturalist 23: 257–353. Sikes, D. and J. Stockbridge. 2013. Description of Caurinus tlagu, new species, from Prince of Wales Island, Alaska (Mecoptera, Boreidae, Caurininae). ZooKeys 316: 35–53. Tan, J. L. and B. Z. Hua. 2009. Description of the immature stages of Bittacus planus Cheng (Mecoptera: Bittacidae) with notes on its biology. Proceedings of the Entomological Society of Washington 111: 111–121. Thornhill, R. 1973. The morphology and histology of new sex pheromone glands in male scorpionflies, Panorpa and Brachypanorpa (Mecoptera: Panorpidae and Panorpodidae). Great Lakes Entomologist 6: 47–55. Thornhill, R. 1975. Scorpionflies as kleptoparasites of web‐building spiders. Nature 258: 709–711. Thornhill, R. 1978. Sexually selected predatory and mating behavior of the hangingfly, Bittacus stigmaterus (Mecoptera: Bittacidae). Annals of the Entomological Society of America 71: 597–601. Thornhill, R. 1980a. Rape in Panorpa scorpionflies and a general rape hypothesis. Animal Behavior 28: 52–59.

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Thornhill, R. 1980b. Mate choice in Hylobittacus apicalis (Insecta: Mecoptera) and its relation to some models of female choice. Evolution 34: 519–538. Thornhill, R. 1981. Panorpa (Mecoptera: Panorpidae) scorpionflies: systems for understanding resource‐defense polygyny and alternate male reproduction efforts. Annual Review of Ecological Systems 12: 355–386. Thornhill, R. 1984. Alternative female choice tactics in the scorpionfly Hylobittacus apicalis (Mecoptera) and their implications. American Zoologist 24: 367–383. Thornhill, R. and J. Alcock. 1983. The Evolution of Insect Mating Systems. Harvard University Press, Cambridge, Massachusetts. 576 pp. Tillyard, R. J. 1935. The evolution of the scorpion‐ flies and their derivatives (Order Mecoptera). Annals of the Entomological Society of America 28: 1–45. Trautwein M. D., B. M. Wiegmann, R. Beute, K. M. Kjer and D. K. Yeates. 2012. Advances in insect phylogeny at the dawn of the

postgenomic era. Annual Review of Entomology 57: 449–468. Wang Y., C. C. Labandeira, C. Shih, Q. Ding, C. Wang, Y. Zhao and D. Ren. 2012. Jurassic mimicry between a hangingfly and a ginkgo from China. Proceedings of the National Academy of Sciences USA 109: 20514–20519. Whiting, M. F. 2002. Mecoptera is paraphyletic: multiple genes and phylogeny of Mecoptera and Siphonaptera. Zoological Scripta 31: 93–104. Willmann, R. 1981. Phylogenie und Verbreitungsgeschichte der Eomeropidae (Insecta: Mecoptera) Ein Beispiel für die Anwendung der phylogenetischen Systematik in der Palaontologie. Paläontologische Zeitschrift 55: 31–49. Willmann, R. 1983. Die phylogenetischen Beziehungen unter den sudamerikanischen Bittacidae (Insecta: Mecoptera). Zoolischer Beiträge 28: 47–65. Willmann, R. 1987. The phylogenetic system of the Mecoptera. Systematic Entomology 12: 519–524.

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Part III Perspectives

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24 The Fossil History of Insect Diversity Conrad C. Labandeira1,2,3 1

Department of Paleobiology, Smithsonian Institution, National Museum of Natural History, Washington, DC, USA Department of Entomology, University of Maryland, College Park, Maryland 20742, USA 3 College of Life Sciences, Capital Normal University, Beijing, P.R. China 2

The overwhelming feature of macroscopic life on Earth is the abundance and dominance of arthropods. Although arthropods occupy virtu­ ally every nook and cranny of an overwhelm­ ingly marine planet, their diversity has become intensely magnified over the past 420 million years, from an inauspicious beginning through an expanding presence as macroscopic life invaded the continental realm of terrestrial and  freshwater ecosystems. Of the continental arthropodan groups that have undergone major radiations, such as arachnids, myriapods, and crustaceans, it is the insects (including closely related groups within the broader Hexapoda clade) that have contributed to the overwhelm­ ing bulk of species richness (Wheeler 1990). This process commenced in a significant way sometime during the Middle Silurian to Early Devonian about 420 to 405 million years ago  (mya), and by the Late Carboniferous (Pennsylvanian) 323 to 299 mya, insects became the dominant, most speciose animal group on land. Perhaps the best way to chart this expand­ ing cone of insect diversification is to observe their development through the historical lens of biodiversity, in all of its major taxic, taxonomic, ecological, morphological, functional, and behavioral manifestations.

The insect fossil record documents long‐term trends and sudden episodes that influence the development of biodiversity. Long‐term trends include environmental and biological shifts of lineages expressed over tens of thousands to hundreds of millions of years. Environmental changes include the sudden, 100,000‐year‐long Paleocene–Eocene Thermal Maximum (PETM) at 56 mya that initiated a major transformation of global climate change. The PETM was a tran­ sient spike in elevated atmospheric tempera­ tures and carbon dioxide levels that reset the clock for insect‐herbivore diversity for the remaining Cenozoic Era. This geologically tran­ sient interval was followed by a several million‐ year‐long gradual warming during the Eocene Epoch, which climaxed in the Early Eocene Climatic Optimum about 50 mya. Similarly, a major biological transformation included the approximately 35 million‐year‐ long initial angiosperm diversification inter­ val  spanning the late Barremian (125 Ma) to Turonian (90 Ma) geological stages of the mid‐ Cretaceous. The diversification of angiosperms is one example of a geologically important, pro­ tracted process that eventually affected all levels of insect diversity. This contrasts with episodic, geologically short‐term perturbations, such as

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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the earlier Permian–Triassic (P–Tr) ecological crisis and attendant mass extinction that trans­ formed insect diversity and its ecologically related associations 252 mya. The P–Tr catas­ trophe was overwhelmingly the most significant event or process to affect insect diversity over its 420‐million‐year presence on the planet. These three processes and events represent a sample of the types of influences that have determined the trajectory of insect diversity in geological deep time. In this contribution, “deep time” is distinguished from “shallow time,” with the latter referring to mostly biogeographical consequences of the Pleistocene glaciation and inter‐glaciation phases of the Pleistocene during the past 2.6 million years, ending in the mega­ faunal extinctions that occurred around 11,000 years ago. Human civilization was launched and expanded during the brief, post‐extinction interval of the Holocene Epoch. I focus on the five major types of diversity documented in the insect fossil record. Consequently, the broader aspects of past insect biodiversity are discussed, avoiding a focus on taxic and taxonomic diversity that historically has been the overwhelming context for most studies of past insect diversity (Labandeira and Sepkoski 1993, Jarzembowski and Ross 1996, Dmitriev and Ponomarenko 2002). A broad spectrum of diversity from the fossil record is discussed, most of which spotlights the work of the Labandeira Lab at the National Museum of Natural History, and among other collabora­ tions, research with colleagues at the Wilf Lab (Pennsylvania State University), the Wappler Lab (University of Bonn, Germany), and the Ren Lab (Capital Normal University, Beijing, China). These research partnerships have expanded alternative approaches toward understanding past insect diversity in the fossil record. The integration of other, related fossil records, such as the fossil plant, sedimentological, and food‐ web records, provide a broader context for understanding insect diversity. A focus of this contribution is plant–insect interactions in the fossil record, specifically her­ bivory, pollination, and mimesis. Other topics

include the historical role that parasitoid insects have had in consumption of other arthropods, the importance that insects in lake ecosystems have had in collectively linking past with recent world biotas, and evidence for major environ­ mental and biotal changes that have determined the course of insect diversity through time. One aspect of past insect diversity that I do not dwell on extensively is phylogenetic approaches, which are adequately covered in other chapters of this volume. The concluding discussion incorporates an understanding of why fossil insect diversity is relevant for the current condi­ tion of terrestrial and freshwater ecosystems of the planet.

24.1 ­Importance of the Insect Fossil Record Insects have a distinctive preservation potential that has contributed to their unique fossil record. Like other arthropods, insect exoskele­ tons have a modest ability to be incorporated into the fossil record  –  somewhere between soft‐bodied groups such as comb jellies and flat­ worms, which essentially lack a fossil record, and highly skeletonized groups such as brachio­ pods, mollusks, and vertebrates, which have a comparably excellent fossil record (Gaston and Spicer 2005). Insect exoskeletons are leathery to brittle, occasionally mineralized, but structur­ ally durable, and consist of typically flexible cuticle that often is impregnated with organic, crosslinked molecules that provide resistance to environmental and biological destruction (Duncan 1997). Nevertheless, insect exoskele­ tons also are considerably more amenable to decay than are mineralized shells composed of inorganic molecules such as calcium carbonate (mollusks) or silica (some sponges). This inter­ mediate level of preservation is compensated by broader‐scale features of insects in the fossil record, such as their elevated taxonomic diver­ sity, high abundance levels, and considerable stratigraphic throughput in lacustrine (lake) deposits and amber (Labandeira 1999, 2014a).

24  The Fossil History of Insect Diversity

Consequently, when considered with other freshwater and terrestrial organisms, insects are only surpassed in fossil abundance (numbers of individuals) by microorganisms and the leaves of vascular plants, and typically far surpass the numbers of vertebrates, fungi, nematodes, and other arthropods in local deposits.

24.2 ­Types of Insect Diversity Past and Present Simply put, biodiversity is a measure of the vari­ ety of organisms occurring within a particular unit of interest (Wilson 2000). Historically, the predominant type of biodiversity recorded in the fossil record has been taxonomic and taxic diversity. Diversity in the insect fossil record typically is provided either at the locality level, with an enumeration of all resident species (tax­ onomic diversity), or more generally has been counts at the genus or, more commonly, the family level, expressed globally through time (taxic diversity). Taxonomic and taxic diversity are but one type of diversity assessed in the fos­ sil record. Five major types of biodiversity have been used or mentioned, or at least are applica­ ble to the fossil insect record: (i) taxonomic and taxic diversity, (ii) ecological diversity, (iii) mor­ phological diversity (also referred to as dispar­ ity), (iv) functional diversity, and (v) behavioral diversity. These five primary types of biodiver­ sity are linked in complex ways. For example, morphological diversity, typically expressed as structural disparity in the fossil record, is inti­ mately tied to functional diversity, or can have an ecological basis, such as feeding type (Labandeira 1997). Behavioral diversity can be part of morphological and functional diversity that is present, for example, in a particular group of fossil social insects. Occasionally these five primary types of biodiversity have diffuse boundaries. Because these five types of biodiversity share elements in common, there have been other proposals for classifying biodiversity. Gaston and Spicer (2005) list the three primary types of

biodiversity as organismal diversity, genetic diversity, and ecological diversity, whereas Erwin (2008) discusses seven such types: (i) taxic diversity, (ii) phylogenetic diversity, (iii) morphological disparity (or diversity), (iv) func­ tional or ecospace diversity, (v) architectural diversity and ecosystem engineering, (vi) behav­ ioral diversity and social complexity, and (vii) developmental diversity. Clearly, Erwin’s (2008) divisions intergrade. Could developmental diversity, reflecting the diversity of genes and their immediate gene products, be integrated with morphological disparity that expresses the observable phenotypes of those same genes? How do the trophic roles of ecological diversity, for example, become functional diversity when those same trophic roles are expressed by how organisms function within a broader ecological community? Overlap issues might become evi­ dent in the discussion below, but the five types of biodiversity are treated as separate entities. 24.2.1  Taxonomic and Taxic Diversity

Taxonomic and taxic diversity consists of the variety of insects that are categorized by the taxonomic hierarchy and includes species, gen­ era, families, and orders measured at a fossil site (taxonomic diversity), or globally through time (taxic diversity). As traditionally practiced, tax­ onomic diversity, also termed richness, is a tally of taxa at a particular local site or larger speci­ fied region. Geographically based taxonomic diversity is measured within a site (alpha diver­ sity), between sites within a region of interest (beta diversity), or among more broadly defined regions (gamma diversity). These measures of diversity, or richness, have a long and distin­ guished history in the marine and terrestrial realms in biology (Whittaker 1972, Purvis and Hector 2000), as well as in the marine and con­ tinental (terrestrial and freshwater) realms in paleobiology (Alroy et al. 2008). Taxic diversity consists of taxa, typically gen­ era or families that are measured for a particular time unit. Taxic diversity in the insect fossil record usually involves worldwide or regional

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tallies of families for a specified interval of time and typically is global in geographic scope. Examples include families of Insecta over a 420‐ million‐year interval from the Early Devonian to the Recent (Labandeira and Sepkoski 1993) (Fig. 24.1); families of Coleoptera, Diptera, Hymenoptera, and Lepidoptera since their ear­ liest fossil occurrences sometime during the early Permian to Early Jurassic (Sohn et  al. 2015, Smith and Marcot 2015) to the present (Fig. 24.2); or genera of Coleoptera over the past 20,000 or so years since the Late Pleistocene and through the Holocene (Elias 1996). Although originally fossil insect diversity was evaluated at the conventional family level for all Insecta with fossil records, recent examinations have con­ centrated on family‐level analyses, but within orders such as the Lepidoptera (Sohn et  al. 2015) or Coleoptera (Smith and Marcot 2015). Recently, there have been efforts to comprehen­ sively examine the taxic fossil insect record at the genus level (Clapham and Karr 2013, Nicholson et  al. 2015). Another approach has been the study of Pleistocene and Holocene tax­ onomic and taxic diversity with an emphasis on the regional diversities of beetle genera and spe­ cies during the waxing and waning of major gla­ ciation events (Matthews 1970, Coope 1990, Elias 1996). The two components of taxonomic diversity are origination and extinction (Labandeira 2005b). Origination is the number of first appear­ ances of a particular taxon of interest, expressed as a rate (Labandeira 2005b). Extinction is the converse of origination, and is the number of last appearances of the relevant taxon, also expressed as a rate. Throughout a given time interval, the initial standing diversity, which is the beginning number of evaluated taxa plus the subsequent additions from originations and subtractions from extinctions, provides the diversity history of a taxon. Subtraction of an extinction rate from an origination rate yields the turnover rate, which can be positive or negative. The family‐level diversity curve of insects is almost flat for the 87‐million‐year period from the Early Devonian to the boundary between the

Mississippian and Pennsylvanian subperiods (Fig. 24.1a); this range represents the earliest three known insect families (Whalley and Jarzembowski 1981, Labandeira et  al. 1988, Grimaldi and Engel 2004, Garrouste et al. 2012). After this interval of depauperate diversity, there is a distinct semilogarithmic increase of insect families that commenced at the beginning of the Pennsylvanian Subperiod and continued through the Permian Period, followed by a dramatic drop at the end of the Permian Period (Labandeira and Sepkoski 1993), coinciding with the ecological and evolutionary crisis that defines the P–Tr boundary event. A distinct recovery interval ensues throughout the Triassic, as diversity levels increased through the succeeding Triassic Period, although the peak of Paleozoic diversity at the mid‐Permian was not reached until the Early to Middle Jurassic boundary, some 90 million years later. Family‐level insect diversity continued to rise during the Late Jurassic, but remained essen­ tially flat, with minor upturns and downturns, throughout the Late Jurassic to Late Cretaceous. Insect diversity seems to have been unaffected by the Cretaceous–Paleogene (K–Pg) event that caused the extinction of much marine and con­ tinental life, including ammonite mollusks, rud­ istid corals, dinosaurs, and seed plants (Labandeira et al. 2016b). The subsequent trend of insect diversity during the Paleogene and Neogene periods is best described as a sustained increase to the present day, with a capture rate (as of 1993) of 63% of the approximately 980 modern insect families  –  a rate that has not changed appreciably with more recent analyses. However, family‐level diversity tapered off dur­ ing the Neogene Period over the past 23 million years. One explanation for this is that the mod­ ern level of family‐level diversity represents a saturation limit that has dampened the Neogene rate of family‐level increase. An alternative explanation is the pull‐of‐the‐recent phenome­ non (Raup 1979), in which the more modern part of the insect‐family diversity record is bet­ ter sampled than are older deposits, thereby increasing the potential recent recovery rate relative to the older part of the fossil record.

24  The Fossil History of Insect Diversity (a)

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Figure 24.1  Taxic and taxonomic diversity 1. (a) Family‐level insect diversity for the Phanerozoic, using the range‐ through method (Labandeira and Sepkoski 1993, Labandeira 2005b), updated from Labandeira (1994). (b) Raw data on originations, based on Labandeira (1994). (c) Raw data on extinctions. All data for (a–c) are based on Labandeira’s (1994) compendium of fossil insect families. For geochronologically short stages from the latter part of the Permian Period, the Bajocian–Bathonian of the Jurassic Period, and much of the Paleogene and Neogene Periods, data representing two or sometimes three stages are combined into a single point. Numbered arrows represent major diversification and extinction events discussed in the text. Abbreviations: Cis, Cisuralian; E, early; Eo, Eocene; G, Guadalupian; L, late; Lo, Lopingian; M, middle; Mio, Miocene; Neog, Neogene; Ol, Oligocene; P, Pleistocene; Pa, Paleocene; Pen, Pennsylvanian. The Carboniferous Period is formally divided into an earlier Mississippian Subperiod and a later Pennsylvanian Subperiod. Adapted from Labandeira (2005b), used with permission of the Entomological Society of America.

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Figure 24.2  Taxic and taxonomic diversity 2. Family‐level diversity of four major, holometabolous orders, from fig. 4 of Sohn et al. (2015). All data were sourced from Labandeira (1994), with updates. Range‐through data were plotted at stage‐interval midpoints. The age of Baltic amber is taken as Middle Eocene (Labandeira 2014a); the overall geochronology is from Gradstein et al. (2012). Abbreviations as in Figure 24.1. Adapted from Sohn et al. (2015), used under terms of the Creative Commons Attribution 4.0 international license (http://creativecommons.org/ publicdomain/zero/1.0/).

A more dynamic, time‐specific view of fos­ sil  insect diversity data involves origination (Fig. 24.1b) and extinction (Fig. 24.1c). For orig­ inations, there are five major, distinctive peaks. In the Paleozoic Era, the initial 87 million years started with the earliest insect occurrences of the Collembola, Archaeognatha, and a probable stem‐group pterygotan lineage from the earlier Devonian, with an origination rate remaining at zero. After several tens of millions of years, and starting at the Mississippian–Pennsylvanian boundary, there is a dramatic increase in origi­ nation that lasts through the first 15 or so mil­ lion years of the Early to Middle Pennsylvanian Period, at which time the family‐level diver­ sification of the Insecta was dramatically launched (Labandeira 2005b). This near geo­ logically  instantaneous event represents the initial increase of 12 major, overwhelmingly

pterygote (winged insect) lineages that approxi­ mately are  equivalent to taxonomic orders (Event 1 of Fig. 24.1b). The diversification event is followed by a major increase in the extinction rate at the Middle Pennsylvanian to Late Pennsylvanian boundary in equatorially cen­ tered Euramerica (most of North America joined to Western Europe) and other paleocon­ tinents (Event 1 of Fig. 24.1c). This extinction is associated with an ecological transformation of coal‐swamp floras dominated by extinct line­ ages of lycopods, horsetails, ferns, and a variety of seed plants, such as cordaite coniferophytes and medullosan seedferns, to a successor flora dominated overwhelmingly by marattialean tree ferns. Insect origination dramatically spikes through­out the succeeding early Permian, which represents the initial diversification of already

24  The Fossil History of Insect Diversity

established orthopteroid, hemipteroid, and hol­ ometabolous insect groups (Event 2, Fig. 24.1b). Simultaneously, the extinction of many Late Pennsylvanian taxa occurred (Event 2, Fig. 24.1c), especially many lineages of paleodictyop­ teroid “beaked dragonflies” with piercing‐and‐ sucking mouthparts (Labandeira and Phillips 1996a), some archaic odonatopteran dragonfly taxa with raptorial mandibulate mouthparts, and early orthopteroid lineages that were inhab­ itants of coal‐swamp biotas. This event evidently was global in scope and is documented in the paleocontinents of Euramerica, Angara (north‐ central Asia), and various terranes that consti­ tuted Cathaysia (portions of eastern Asia). For both origination and extinction rates, there was a severe drop‐off toward the P–Tr boundary that is complicated because of the Signor–Lipps effect (Fig. 24.1c). The Signor–Lipps effect (Signor and Lipps 1982) states that the last occurrences of taxa poorly represented in the fossil record will be recorded as a major extinc­ tion event in a stepwise fashion, and smeared back onto earlier time intervals even if their true extinction is at the major event. Consequently, the Signor–Lipps effect explains that a stair‐ stepped pattern of clade extinction (in this case, insect families) indicates that a poor fossil record is present, rather than providing evidence for any intrinsic series of biological events, such as closely spaced, minor, later Permian extinctions preceding the major P–Tr mass extinction. Nevertheless, the four orders of the Paleodi­ ctyopteroidea became extinct, and there is no doubt that the P–Tr event was ecologically cata­ strophic and the “mother of all extinctions” (Erwin 2006). For the Mesozoic Era, the Triassic Period and the first two‐thirds of the Jurassic Period indi­ cate that insect origination and extinction rates increased modestly but stayed at a plateau through several cycles until the mid‐Late Jurassic. During the mid‐Late Jurassic, origina­ tion rates increased significantly (Event 3, Fig.  24.1b) but extinction rates much less so (Event 3, Fig. 24.1c). Following these spikes, originations declined during the first half of the

Early Cretaceous, but a rise in both origination and extinction commenced during the last half of the Early Cretaceous, peaking in the Aptian– Albian of the Early Cretaceous (Event 4, Fig.  24.1b,c), contemporaneous with the initial phase of the angiosperm radiation. Notably, the effect is approximately the same for originations and extinctions, indicating that angiosperm diversification had no net effect on insect ­diversity at the family level, although there was ­significant turnover (Labandeira and Sepkoski 1993). One explanation for this pattern is that, at least for 280 plant-associated insect families, extinctions of earlier gymnosperm‐associated families are approximately balanced by subse­ quent originations of angiosperm‐associated families (Labandeira 2014b; Fig. 24.2). This pro­ cess would explain the intervening diversity gap of the Aptian–Albian stage interval, responsible for the lag between extinctions of earlier gym­ nosperm‐affiliated taxa and the yet‐to‐occur originations of subsequent angiosperm‐associ­ ated taxa (Labandeira 2014b; Fig. 24.2). For the remainder of the Late Cretaceous, origination and extinction rates decreased to a basal level, with a small spike in the Campanian Stage of the Late Cretaceous. At the end of the Cretaceous, family‐level diversity began to rise, and remained unaffected by the K–Pg ecological cri­ sis that caused the demise of the dinosaurs and many other terrestrial groups (Labandeira et al. 2016a). For the Cenozoic Era, there are two major peaks of originations in family‐level diver­ sity  –  a modest rise that occurred in the early Eocene, the most dramatic of which is a spike in the late Eocene (Event 5, Fig. 24.1b). These orig­ ination events have been attributable to excep­ tionally well‐preserved and taxonomically diverse deposits such as Baltic amber and the Florissant Formation of Colorado, United States (Labandeira and Sepkoski 1993, Sohn et  al. 2015). Notably, extinction declined markedly throughout the Paleogene and Neogene p ­ eriods, approaching levels of zero. Much of this improvement of the fossil record toward the Recent has to do with the pull‐of‐the‐recent

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phenomenon, which causes past biodiversity estimates to be more aligned with modern bio­ diversity (Raup, 1979). Repeated family‐level analyses of the insect fossil record have verified the general pattern of fossil‐insect diversity shown by Labandeira and Sepkoski (1993) (Fig. 24.1a). This pattern has been borne out by (i) subsequent analyses using largely independent data sets (Jazembowski and Ross 1996, Dmitriev and Ponomarenko 2002, Nicholson et  al. 2015), (ii) a more complete account of the fossil‐insect record incorporat­ ing new discoveries of insect families previously unrecognized in the fossil record, and (iii) inclu­ sion of recent taxonomic revisions (Sohn et al. 2014, Nicholson et al. 2015, Smith and Marcot 2015). The most notable change is a recent anal­ ysis (Nicholson et  al. 2015) that involves the movement of some peaks of origination and extinction to an adjacent geological stage, attributable to more modern techniques for re‐ dating of several fossiliferous deposits with abundant and diverse insect occurrences. Phylogenetic diversity can be expressed directly from associated chronograms providing the ­relationships among closely knit lineages such as planthoppers and certain parasitoid fungi (Fig. 24.3) (Song and Liang 2013). In another case, hypocrealean ascomycotan fungi are a clade that has substantial phylogenetic diversity and con­ sists of pathogens, parasites, and parasitoids on a variety of terrestrial arthropods, especially insects (Fig. 24.4a) (Sung et  al. 2008). Ants infected with hypocrealean fungi (Fig. 24.4c), which become “zombie ants,” have a fossil record extending to the earliest middle Eocene (Fig. 24.4b) (Hughes et  al. 2011). Such phylogenetic data could be considered an independent source of taxonomic or taxic diversity. Accordingly, a recent contribution to taxonomic diversity of

modern and fossil‐insect taxa involves lineages‐ through‐time (LTT) that provide past diversity estimates from phylogenetic chronograms (e.g., Moreau et al. 2006, McKenna and Farrell 2006). LTT data are retrieved from a phylogenetic chronogram and are a plot of the number of line­ ages present as a function of geological time. The number of lineages for a particular time interval is derived partly from the proportion of terminal taxa. Consequently, diversity trends are generated from phylogenetic analyses and may bear consid­ erable divergence from source fossil occurrences, some of which, however, may be used as calibra­ tions of the phylogenetic chronogram from which an LLT is derived. In many cases, LTT data, together with detailed phylogenetic analy­ ses of hyperdiverse groups, such as beetles, can reveal intervals of major increases in diversity, such as evolutionary radiations (McKenna and Farrell 2006, Hunt et al. 2007). 24.2.2  Ecological Diversity

Ecological diversity expresses the differences among insects that pertain to their trophic interactions or to other community‐level pro­ cesses at scales ranging from local populations, through habitats and into biomes. Ecological diversity can be partitioned into community‐ level or even broader ecosystem‐wide diversity that includes all the multifarious inter‐relation­ ships among organisms, such as the role of insect development and ontogeny on feeding mechanisms, membership in dietary guilds, nestedness within food webs, local herbivore communities on source‐plant species, habitat structuring of broader communities, and even­ tually ecological structure in entire ecosys­ tems  (Hochuli 2001; Bascompte et  al. 2003; Bascompte 2009; Novotny et al. 2010, 2012). In

Figure 24.3  Taxic and taxonomic diversity 3. Chronogram of the Fulgoroidea (Hemiptera) that was estimated with BEAST, from Bayesian phylogenetic software. Shaded rectangles group major clades. The estimated divergence times are given near the nodes; white bars represent 95% confidence intervals. Time units are provided in millions of years. N, Neogene Period. Additional details are provided by Song and Liang (2013). Adapted from figure 3 in Song and Lang (2013) from PLoS ONE, used under Attribution 4 international license through a Creative Commons agreement.

24  The Fossil History of Insect Diversity Heteroptera Cicadomorpha Sternorrhyncha 195

Cixiidae

221 129

Delphacidae

255 120

Kinaridae 24

Caliscelidae

65

Tettigometridae Achilixiidae Nogodinidae

79

Derbidae 10

Acanaloniidae

141

Issidae

Lophopidae

90

Eurybrachidae Flatidae

18 128

98

Ricaniidae Achilidae

46 133

Nogodinidae 16

Tropiduchidae

60

Meenoplidae

78

Fulgoridae

123 90

Carboniferous Permian Triassic

350

300

250

200

Jurassic

Dictyopharidae

Cretaceous Paleogene N

150

Millions of years

100

50

0

731

732

Insect Biodiversity: Science and Society (a) Stachybotrys clade Nectriaceae Boinectriaceae Hypocreaceae

Hypocreales Cordycipitaceae

Clavicipitaceae

(b)

Animal Plant Fungi Soil Ambiguous Missing

Ophiocordycipitaceae

Ophiocordyceps

Triassic

200

Jurassic

Cretaceous

150

100

(c)

Paleogene Neog

50

0 Ma

Figure 24.4  Taxic and taxonomic diversity 4, ecological diversity 1. The diversity of fungal parasitoid clades during the mid‐Mesozoic to Recent, and an example of an ant zombification association with a parasitoid fungus revealed by distinctive plant damage from the Middle Eocene Messel deposit in central Germany. (a) Divergence age estimates of major lineages of the fungal lineage Hypocreales. The chronogram was constructed based on a tree from maximum likelihood analyses. The calibrations of the crown node of Ophiocordyceps were based on an identified fungal parasite of a scale insect from the Upper Albian of the Early Cretaceous. Other details are provided by Sung et al. (2008). (b) A nearly complete fossil leaf from Messel, Middle Eocene of Germany, with 29 ant death‐grip scars centered on eleven secondary veins. (c) A modern ant specimen (Anderson et al. 2009) showing a mature Ophiocordyceps unilateralis stroma emerging from the head capsule of a dead ant, Camponotus leonardi, whose mandibles are attached to the undersurface of a major vein (Hughes et al. 2011). Part (a) adapted from Sung et al. (2008), under an open‐access agreement with Elsevier. Parts (b,c) reproduced by unrestricted use of Hughes et al. (2010), provided by terms of the Creative Commons Attribution, under a license of The Royal Society.

24  The Fossil History of Insect Diversity

practice, there are two frequently discussed ways that ecological diversity has been exam­ ined in the fossil record. First is ecological char­ acterization of entire biotas, such as well‐studied lacustrine shale and terrestrial amber deposits that are highly diverse assemblages from par­ ticular habitats containing plants, arthropods, fungi, vertebrates, and their inter‐relationships (Behrensmeyer and Hook 1992, Dunne et  al. 2014, Buatois et  al. 2016). Such an approach constitutes an examination of biotal diversity. Second is the more particular study of multi­ trophic plant–insect interactions. Most ecologi­ cal approaches to fossil‐insect diversity involve examinations in space or time of trophic diver­ sity or its biological aspects that pertain to feed­ ing or the biology of food consumption. The examination of plant–insect interactions in deep time has been a major, expanding research program during the past two decades and increasingly has been linked to evolutionary processes in modern (Schoener 2011) and fossil (Labandeira 2002a) ecosystems. 24.2.3  Biotal Diversity

Ecological diversity has increased since the ear­ liest greening of coastlines somewhat more than 400 mya, as morphologically simple vas­ cular plants colonized transitional marine– freshwater and marine–terrestrial habitats during the Silurian and Devonian (Labandeira 2005a). This was soon followed in the Early Devonian by further terrestrialization of organ­ ismic groups and establishment of mutualistic and antagonistic associations of fungi, plants, and early terrestrial arthropods, the latter of which consisted of crustaceans, early myria­ pod‐like forms, scorpions, arachnids, mites, collembolans, bristletails, and possible ptery­ gotes. Although the earliest, significant biota of continental organisms is the Rhynie chert deposit in southern Scotland, of Early Devonian age (Fayers and Trewin 2004), by the Middle Devonian at about 385 mya a significantly more complex biota was established. This biota con­ tained more mutualistic and antagonistic f­ ungal

taxa and advanced groups of plants such as the earliest seed‐bearing trees, but still lacked any significant diversity increase in the basic types of arthropods exemplified by mites, myriapods, and crustaceans (Stein et  al. 2012). Insects other than bristletails were conspicuously absent (Shear and Kukalová‐Peck 1990). By this time, a limited number of herbivore associa­ tions with terrestrial microarthropods had been present (Labandeira 2007a, Laban­ deira et  al. 2013). Notably, vertebrates had not entered continental ecosystems, restricting ani­ mal diversity primarily to arthropods. Verteb­ rate absence might have been attributable to low atmospheric oxygen levels when compared to present‐day concentrations, allowing arthro­ pods the ability to efficiently respire by tracheae and book lungs under conditions that were suffocating for vertebrates, which possessed more inefficient mechanisms of respiration under marginal to lethally low O2 levels (Ward et al. 2006). Not much is known about terrestrial biotal diversity throughout the rest of the Devonian and into the Mississippian, although freshwater biotas are known from deposits such as East Kirkton in Scotland (Minter et  al. 2016). Subsequently, during the Pennsylvanian Period, biotal diversity substantially increased among local terrestrial communities within the great, regional coal‐swamp forests. Perhaps the most notable of these biotas is the terrestrial aspect of  the Mazon Creek Biota of the Middle Pennsylvanian time, dated to approximately 311 million years in age (Shear and Kukalová‐Peck 1990, Shabica and Hay 1997, Selden and Nudds 2005). This deposit, of north‐central Illinois in the United States, consisted of an extensive delta and associated estuary surrounded by a forest of pole‐like lycopods and horsetails, along with less statuesque medullosan seed ferns, marattialean tree ferns, and often shrubby cord­ aites (Wittry 2006). Arthropods were diverse in this environment, represented by varied crusta­ ceans, medium‐sized to gigantic myriapods, scorpions, arachnids, horseshoe crabs, and myriad insects such as large odonatopterans

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and paleodictyopteroids, difficult to distinguish orthopteroids, ancestors to modern stoneflies, and the earliest known holometabolan, pre­ served as an externally feeding caterpillar (Shear and Kukalová‐Peck 1990, Labandeira 1999, Grimaldi and Engel 2005, Nel et al. 2013, Haug et al. 2015). This ecosystem constituted the Wet Biome, which had its origin during the Middle Pennsylvanian interval, and it coexisted with the Dry Biome throughout the later Pennsyl­ vanian. The Dry Biome gradually supplanted the Wet Biome later in the Permian Period. The early Permian Elmo Biota of Kansas is probably the best known expression of the Dry Biome (Beckemeyer 2000), consisting of plants, arthro­ pods, and vertebrates rivaling those of the Mazon Creek Biota in diversity. After the major ecological crisis at the P–Tr boundary 252 mya, biotal diversity collapsed at local, regional, and global geographical scales (Looy et al. 2001, Erwin 2006). The recovery of biotal diversity was deeply constrained by an Early Triassic greenhouse interval that caused lethally hot temperatures in continental interi­ ors, especially in the Southern Hemisphere on the supercontinent of Gondwana (Sun et  al. 2012). Nevertheless, new lineages emerged dur­ ing the Middle Triassic, which included ferns and seed plants (Hochuli et al. 2010), along with new groups of insects that evolved from the pruned‐down lineages surviving the P–Tr cull­ ing event (Labandeira 2006b). A similar pattern held for the emergence of vertebrates (Botha and Smith 2006) that became ancestors to the dominant groups later in the Mesozoic, promi­ nently including dinosaurs, mammals, and birds. Insect diversity did not approach the lev­ els seen during the Permian until the late Middle Jurassic; plant–insect associational diversity took from 20 to 25 million years to recover to pre‐P–Tr‐crisis levels (Labandeira 2006b, Labandeira et al. 2016a). The best documenta­ tion for this recovery of ecological diversity is the Molteno Biome from the Karoo Basin of South Africa, which preserves numerous assem­ blages of terrestrial plants, insects and plant– insect interactions, and limited occurrences of

fish and other elements from associated lacus­ trine deposits (Anderson and Anderson 1989, 1998; Labandeira 2006b). Notably, terrestrial vertebrates are represented only as sedimentary trace fossils. By the mid‐Mesozoic, particularly during the mid‐Jurassic of Eurasia, a series of diverse biomes arose in which a substantial number of insect herbivore, pollinator, and mimicry asso­ ciations codiversified with gymnosperms (Labandeira 2006a, 2010, 2014b; Ren et al. 2009; Ding et  al. 2014, 2015). These ecosystems  – particularly the eastern Asian biotas of the Jiulongshan, Karabastau, and Yixian formations – collectively lasted for about 50 million years and preceded the diversification of angiosperms launched during the mid‐Cretaceous. The types of plant–insect associations in these ecologi­ cally diverse communities of gymnosperms during the Middle Jurassic to Early Cretaceous were analogous to those of subsequent angio­ sperm‐dominated communities of the later Cretaceous (Labandeira et al. 1994; Labandeira 2010, 2014b; Doorenweerd et al. 2015). One way of examining insect‐herbivore diversity on gym­ nosperms is through long‐term tracking of component community structure formed by insect herbivores on a specified lineage or related lineages of plant hosts that share the same morphological attributes. (A component community consists of all trophically dependent species on a plant‐host species, as initially pro­ posed by Root in 1973.) In one study, long‐term changes in herbivory were examined on the component community of three taxa of broad­ leaved conifers with structurally identical leaves (Ding et al. 2015). This study documents major shifts in the style of feeding damage on broad­ leaved conifer hosts throughout an 80‐million‐ year interval during the mid‐Mesozoic of Northeastern China, indicating shifting eco­ logical links among plant hosts, their insect her­ bivores, and their insect consumers (Ding et al. 2015). The diversification rate of the earliest angio­ sperms seems to have had less impact on insect diversity than previously indicated by Crepet and

24  The Fossil History of Insect Diversity

Niklas 2009). The angiosperm diversity pattern seems to be one of slow and steady expansion during the first 20 million years of its Early Cretaceous fossil record, from 135 to 115 mya, followed by ecological dominance in some regions through accompanying major changes in reproductive biology and leaf form. Taxonomic diversity, much of it cryptic, increased during the succeeding 15 million years from 115 to 100 mya and into the rest of the Cretaceous (Friis et al. 2011). As for insect herbivores, pollinators, and plant mimics, there was a fourfold pattern of host‐plant‐related diversity shifts during this gymnosperm‐to‐ angiosperm transition (Labandeira 2014b). These four patterns were: (i) earlier, gymno­ sperm‐feeding lineages continuing across the Cretaceous, retaining their ancestral host rela­ tionships with gymnosperms; (ii) other gymno­ sperm‐adapted lineages becoming extinct when confronted with adapt‐or‐perish consequences in an angiosperm‐dominated world; (iii) still other gymnosperm‐adapted lineages shifting hosts and targeting newly emergent angio­ sperms; and (iv) the origin of new, angiosperm‐ dependent herbivore lineages lacking any earlier history of gymnosperm‐host consumption (Fig.  24.5). The teasing apart of these various modes of extinction, host‐shifts, and origina­ tion requires an adequate fossil history of plant hosts and their insect herbivores. The forma­ tion of these insect‐herbivore diversity patterns likely were contingent on the diversities of the major host‐plant groups – gymnosperms, angi­ osperms and possibly ferns – during this crucial interval of time. Biotal diversities were low, and ecological diversity was depauperate in many plant com­ munities immediately following the K–Pg eco­ logical crisis (Labandeira et al. 2016b), although not nearly to the degree as the considerably more devastating and much earlier P–Tr event (Twichett 2006). There is evidence that biodi­ versity suffered a major and sustained decrease throughout western North America immedi­ ately following the K–Pg event (Labandeira et  al. 2002a, 2002b), whereas its effects in

Western Europe might have been considerably ameliorated (Wappler et al. 2009). Local, com­ munity‐level biodiversity remained disharmo­ nious, with unusual patterns of herbivory occurring up to 1.5 million years after the event (Wilf et  al. 2006), but by the middle Eocene, about 17 million years after the K–Pg crisis, ecological diversity seems to have equilibrated to a modern structure, as evidenced by a biodi­ verse Baltic amber biota (Labandeira 2014a) and a food‐web structure at Messel in Germany that slightly predates deposition of Baltic amber (Dunne et al. 2014) (Fig. 24.6). Evidence for the modernity of later Cenozoic ecosystems is pro­ vided by Dominican amber from the early Miocene, at about 21 million years in age. The Dominican amber biota (Labandeira 2014a) represents one of the most biodiverse biotas of the fossil record. 24.2.4  Plant–Insect Interactional Diversity

Historically, plant–insect interaction studies from the fossil record were examinations of a single association, in which a distinctive type of insect‐mediated interaction occurred on a par­ ticular plant host for a moment in time during the deep past (e.g., Labandeira and Phillips 1996b, 2002). Examples of such extinct, bilateral associations include the following: from the Paleozoic, the occurrence of stereotyped dam­ age of punch and sucking by a thrips or related lineage on pollen of a noeggerathialean seed plant from the late Permian of north‐central China (Wang et  al. 2009); from the Mesozoic, the presence of distinctive ovipositional odona­ tan damage on an herbaceous lycopod from the Middle Triassic of Kyrgyzstan (Moisan et  al. 2012); and from the Cenozoic, cambium engrav­ ings by a scolytine bark beetle on its coniferalean larch host from the Middle Eocene of the Canadian High Arctic (Labandeira et  al. 2001). On rare occasions, an exceptional tritrophic association is discovered (Fig. 24.4b,c), such as a leaf with distinctive death‐grip scars astride major veins, which was made by a particular type of ant that, in turn, was zombified by a p ­ arasitoid

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Angiosperm radiation 35 Ma

Plateau 2 Plateau 1

Stage/ Epoch

Period Ma

Figure 24.5  Ecological diversity 2. A plot showing the diversity of major plant hosts consisting of ferns (in dark gray), gymnosperms (in medium gray), and angiosperms (in light gray), associated with plant‐associated fossil insect families (vertical axis) as a function of geological time (horizontal axis). Plateau 1 of 95 gymnosperm‐associated families (and a few fern families) is replaced by plateau 2 of 110 angiosperm‐associated families (and fewer fern families), on either side of the Aptian–Albian gap. The plateaus are separated by a downturn in plant‐associated insect diversity during the angiosperm radiation. Host-plant affiliations of insect lineages: A, cryptogam and fern hosts only; B, cryptogam to angiosperm transitions; C, gymnosperm hosts only; D, gymnosperm to angiosperm host transitions; E, angiosperm hosts only. The data are derived from figure 13.3 of Labandeira (2014b) used with permission, © (2014), Springer Cham Heidelberg–New York–Dordrecht–London.

fungus (Hughes et al. 2010). Such examinations are important in a broader evolutionary con­ text, but by themselves do not integrate a par­ ticular interaction into an ecological network, and thus cannot be deemed a study of biodiver­ sity. If ecological diversity is the goal of a study, all preserved interactions or associations need to be examined within a biota. Examples from

the modern world include studies of herbivory along altitudinal (Schiedel et al. 2003) and lati­ tudinal (Adams and Zhang 2009) gradients. For fossil data, plant–insect interactional diversity is studied in three basic ways. The first approach to the study of past asso­ ciations involves sets of multiple, individual associations that form a herbivore‐component

24  The Fossil History of Insect Diversity

(a)

(b)

Lake web

Forest web

Figure 24.6  Ecological diversity 3. Ecological diversity involving plant‐ and insect‐dominated ecosystems, as depicted in the lake (a) and forest (b) food webs from 47‐million‐year‐old Eocene Lake Messel in central Germany. Spheres represent taxa, lines are feeding links, and links that loop indicate cannibalism. The vertical axis is the short‐weighted trophic level, with autotrophs and detritus at the lowest level and predatory taxa at the upper levels. Node gray‐scale intensity designates the broad, taxonomic affiliation of trophic species. The bottom‐most ring in black represents primary‐producer plants, including algae, diatoms, and land plants. The light, gray‐hued taxa in the ring immediately above are invertebrates; both the bottom and superjacent rings contain several interspersed, dark gray to black taxa that represent bacteria, fungi, and detritus. More loosely arranged, medium gray taxa at the central to upper part of the web represent vertebrates. Sources are Dunne et al. (2014) and Labandeira and Dunne (2014). Unrestricted use of Dunne et al. (2014) is provided by terms of the Creative Commons Attribution, under a license of the Royal Society.

community (Root 1973) on a particular host‐ plant species. An example of a herbivore‐com­ ponent community is Psaronius chasei, an extinct marattialean tree fern from the Late Pennsylvanian (302 mya) of the Illinois Basin. There is documentation for a variety of interact­ ing arthropods on this tree fern (Labandeira 1998a), including detritivorous oribatid mites on the outer root mantle (Labandeira et  al. 1997), insects engaged in pith boring in trunk

tissues (Labandeira and Phillips 2002), external feeding on frond pinnules (Labandeira 2001), piercing and sucking and galling in and along frond rachises (Labandeira and Phillips 1996a, 1996b), and feeding on sporangia on pinnule undersides (Labandeira 1998a, 1998b, 1999, 2001). Psaronius chasei is the earliest known plant host for which a diverse, insect‐herbivore‐ component community has been documented. The community of Psaronius herbivores was

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extinguished in the Euramerican lowlands at the end of the Pennsylvanian, but survived mini­ mally changed (Futuyma and Mitter 1996) in Cathaysia until the late Permian (D’Rozario et al. 2011). Another example is the herbivores on the extinct conifer Liaoningocladus boii, from the mid‐Early Cretaceous of Liaoning Province in northeastern China. This source‐plant host har­ bored distinctive types of external foliage feed­ ing, piercing‐and‐sucking lesions, a distinctive pattern of leaf mining, and several types of gall­ ing (Ding et al. 2014, 2015; Wong et al. 2015). In addition, there was a stereotyped pattern of ovi­ position that might be linked to the leaf mining (Ding et  al. 2014). The herbivore diversities of these two deep‐time component communi­ ties  –  Late Pennsylvanian P. chasei and Early Cretaceous L. boii  –  are comparable to those occurring today on bracken (Lawton and MacGarvin 1986), a globally distributed fern species that has been studied extensively for the development of geographically based compo­ nent communities. These component commu­ nities have implications for the long‐term biology of the plant host, the evolution of their insect‐herbivore diversities, and more remotely, development of a third trophic level of parasi­ toids and predators (Labandeira 2002b). However, a direct trophic link between insect herbivores and their consumers typically is indi­ rect and inferred in the fossil record. A second approach is to measure standing diversity of plant–insect associations at a par­ ticular locality (Table 24.1, Studies 1–8). Almost always this is done as a measure of alpha diver­ sity; occasionally, individual sites along the same bed or stratum are sampled, which are suf­ ficiently distant from each other that beta diver­ sity can be evaluated. Although diversities of plant–insect interactions have been established for most major intervals of geological time, only two prominent clusters of studies with data from adjacent multiple sites in time and space have been quantitatively analyzed (Labandeira 2013a). The first are studies from four early  Permian localities in Texas, which have

e­ stablished the floral and plant–insect interac­ tional diversities from varying habitats associ­ ated with river drainage systems (Table 24.1, Studies 2–5). The published studies (Beck and Labandeira 1998; Labandeira and Allen 2007; Schachat et  al. 2014, 2015) indicate that each flora had a distinctive signature of insect her­ bivory when compared to other Early Permian bulk floras. This distinctiveness is based on the abundance and diversity of interactions, as well as the amount of herbivorized plant‐surface area. In addition, some plant lineages in each of the four sites exhibit different, often specialized, patterns of targeting plant hosts by particular insect herbivores and feeding types. A third approach is the examination of fossil plant–arthropod associational diversity or rich­ ness, and related indices. This involves assess­ ments of sampled single or multiple floras across a broad spectrum of geological time intervals, spatial scales, habitats, and external conditions, such as mass extinction events or periods of dramatic to gradual environmental change. The narrowest time scales are short‐ term intervals of single deposits representing 102 to 104 and perhaps to 105 years, depending on the depositional environment of the particu­ lar examined deposit. Fluvial (watercourse‐ associated) deposits represent shorter time intervals and greater overall sedimentary thick­ nesses than lacustrine (lake) deposits, which account for greater time intervals and less over­ all thicknesses of strata (Behrensmeyer and Hook 1992). Plant–arthropod interaction stud­ ies at medium‐term time scales are from 106 to 107 years and for long‐term time scales range from 107 to 108 years, accompanied by similar spatial scales. Studies at very‐long‐term time scales can range up to the past 420 million years, representing the time interval for which macro­ scopic land biotas have existed on the planet. Spatial, or geographical, scales range from a single outcrop of 1 m‐deep quarried sites of 1–5 m2 in area to much of the global land sur­ face, the latter in the case of synoptic studies that include a variety of time, habitats, latitudi­ nal zones, and regions with extensive basin

Table 24.1 Biological and geological scale of 30 biodiversity studies using the plant–insect fossil record from the Labandeira, Wilf and Wappler Laboratories*.

Study†

Floras Ages: period studied and stage‡

Plants examined§

Short term (< 106 yr): single locality studies††

Herbivory data recorded¶

Deposit(s) time Deposit(s) spatial duration# extent**

References

DT frequency, 103–104 yr DT richness

1–5 m2

Labandeira et al. 2013

Sphenopsids, ferns, cordaites, conifers, pteridosperms‡‡

DT frequency, 102–104 yr DT richness, herbivorized foliage area

1–5 m2

Labandeira and Allen 2007

Early Permian (Artinskian): ca. 285 Ma

Sphenopsids, conifers, cycadophytes, gigantopterids, peltasperms, incertae sedis

DT frequency, 102–104 yr DT richness, herbivorized foliage area

1–5 m2

Beck and Labandeira 1998

Early Permian (Artinskian): ca. 285 Ma

Sphenopsids, conifers, gigantopterids, cycadophytes, medullosans, noeggerathialeans, peltasperms

DT frequency, 102–104 yr DT richness, herbivorized foliage area

1–5 m2

Schachat et al. 2015

1 Middle Devonian liverwort herbivory and antiherbivore defense

1

Middle Devonian Metzgeriothallus (late Givetian): 384 Ma sharonae, a liverwort

2 Minimal insect herbivory for the Lower Permian Coprolite Bone Bed site of north‐central Texas, USA and comparison to other late Paleozoic floras

1

Early Permian (Sakmarian): ca. 292 Ma

3 Permian insect folivory on a gigantopterid‐ dominated riparian flora from north‐ central Texas

1

4 Insect herbivory from early Permian Mitchell Creek Flats of north‐central Texas: Opportunism in a balanced component community (single locality substudy

1

(Continued)

Table 24.1 (Continued)

Study†

Floras Ages: period studied and stage‡

Plants examined§

Herbivory data recorded¶

Deposit(s) time Deposit(s) spatial duration# extent**

References

5 Plant–insect interactions from early Permian (Kungurian) Colwell Creek Pond, north‐central Texas: The early spread of herbivory in riparian environments (single locality substudy)

1

Early Permian (Kungurian): ca. 278 Ma

Sphenopsids, conifers, cycadophytes, medullosans, peltasperms, gigantopterids

DT frequency, 102–104 yr DT richness, herbivorized foliage area

1–5 m2

Schachat et al. 2014

6 Portrait of a Gondwanan Ecosystem: A new Late Permian fossil locality from KwaZulu‐Natal, South Africa

1

Late Permian (Wuchiapingian): 257 Ma

Lycopods?, sphenopsids, ferns, glossopterids

DT frequency, 102–104 yr DT richness

1–5 m2

Prevec et al. 2009

16¶¶

Late Paleocene (Thanetian): 58 Ma

Ferns, conifers, monocot DT frequency, 104–105 yr and dicot angiosperms DT richness

1–10 km2?

Wing et al. 2009

1

Early Eocene (Ypresian)–middle Eocene (Lutetian) boundary: 47.8 Ma

All vascular plants

0.7 km2

Dunne et al. 2014, Labandeira and Dunne 2014

7 Late Paleocene fossils from the Cerrejón Formation, Colombia, are the earliest record of Neotropical forest 8 Highly resolved early Eocene food webs show development of modern trophic structure after the end‐Cretaceous extinction

Food‐web 6 × 105 yr indices## based on DT data

Intermediate term (106–107 yr): multiple locality studies†† 9 Impact of the terminal Cretaceous event on plant– insect associations

80

Late Cretaceous (late Maastrichtian)–early Paleocene (early Danian)

Bryophytes, sphenopsids, DT frequency, 2.2 million yr DT richness (66.3– ferns, conifers, cycads, 64.1 Ma) Ginkgo, angiosperms

Narrowly regional: (70 km × 25 km)

Labandeira et al. 2002a, 2002b

10 Novel insect leaf mining after the end‐Cretaceous extinction and the demise of Cretaceous leaf mines, Great Plains, USA

228

Late Cretaceous (late Maastrichtian)–late Paleocene (mid Thanetian)

Ferns, angiosperms

Leaf mining DT frequency and DT richness

8.8 million yr (66.3– 57.5 Ma)

Broadly regional: (630 km × 560 km)

Donovan et al. 2014

11 No post‐ Cretaceous ecosystem depression in European forests? Rich, insect‐feeding damage on diverse, middle Palaeocene plants, Menat, France

15

Late Cretaceous (late Maastrichtian)– Paleocene–Eocene boundary (Thanetian–Ypresian boundary)

Dicotyledonous angiosperms

DT frequency, 10.5 million yr 10 km2 DT richness (66.5– 56.0 Ma)

12 Sharply increased insect herbivory during the Paleocene– Eocene Thermal Maximum

5

Late Paleocene (mid Thanetian)–early Eocene (mid Ypresian)

Dicotyledonous angiosperms

DT frequency, 3.7 million yr DT richness (58.9– 55.2 Ma)

Narrowly regional: (150 km × 70 km)

Currano et al. 2008

13 Fossil insect folivory tracks paleotemperature for six million years

9

Late Paleocene (mid Thanetian)–early Eocene (mid Ypresian)

Dicotyledonous angiosperms

DT frequency, 6.7 million yr DT richness (58.9– 52.7 Ma)

Global: (South America, North America, western Europe)

Currano et al. 2010

14 Response of plant–insect associations to Paleocene–Eocene warming

13

Paleocene–Eocene Thermal Maximum– Early Eocene (mid Ypresian)

Dicotyledonous angiosperms

DT frequency, 5.5 million yr DT richness (56.0– 50.5 Ma)

Narrowly regional (100 km2)

Wilf and Labandeira 1999

Wappler et al. 2009

(Continued)

Table 24.1 (Continued)

Study†

Floras Ages: period studied and stage‡

Plants examined§

Herbivory data recorded¶

Deposit(s) time Deposit(s) spatial duration# extent** 2

2

References

15 Testing for the effects and consequences of mid Paleogene climate change on insect herbivory

2

Early Eocene (latest Ypresian)–middle Eocene (mid Lutetian)

Angiosperms

DT frequency, 3.5 million yr DT richness (47.8– 44.3 Ma)

0.4 km –0.7 km for each of the two sites

Wappler et al. 2012

16 Richness of plant–insect associations in Eocene Patagonia: A legacy for South American biodiversity

4

Early Eocene (mid Ypresian)–middle Eocene (early Lutetian)

Dicotyledonous angiosperms

DT frequency, 4.0 million yr DT richness (52.0– 48.0 Ma)

Global: (South America, North (America)

Wilf et al. 2005

Long term (107–108 yr): multiple locality studies†† 17 Insect herbivory from early Permian Mitchell Creek Flats of north‐central Texas: Opportunism in a balanced component community (multilocality substudy)

4

Early Permian (Sakmarian)–Early Permian (Kungurian)

Sphenopsids, conifers, cycadophytes, peltasperms, medullosans, gigantopterids

Narrowly regional DT frequency, 16 million yr (293–277 Ma) (ca. 60 km2) DT richness, herbivorized foliage area

Schachat et al. 2015

18 Plant–insect interactions from the Early Permian (Kungurian) Colwell Creek Pond, north‐ central Texas: The early spread of herbivory in riparian environments (multilocality substudy)

4

Early Permian (Sakmarian)–Early Permian (Kungurian)

Sphenopsids, a conifer, gigantopterids, cycadophytes, medullosans, peltasperms, noeggeranthialeans

Narrowly regional DT frequency, 16 million yr (293–277 Ma) (ca. 60 km2) DT richness, herbivorized foliage area

Schachat et al. 2014

19 Patterns of herbivory across the Permian–Triassic sequence in the Dolomites Region, Southern Alps, Italy

6

Early Permian (Kungurian)–Middle Triassic (Anisian)

Lycopods, sphenopsids, DT frequency, 36 million yr ferns, conifers, cordaites, DT richness (275–239 Ma) cycadophytes, pteridosperms‡‡, ginkgophytes, incertae sedis

20 Patterns of insect herbivory, plant‐host specialization and tissue partitioning on broad‐leaved conifers from the mid Mesozoic of northeastern China

3

Late Triassic (Rhaetian)–Early Cretaceous (late Barremian)

Conifers

DT frequency, 80 million yr Broadly regional: DT richness (205–125 Ma) extensive lake basins in NE China

21 Decoupled plant and insect diversity after the end‐ Cretaceous extinction

14

Late Cretaceous (late Maastrichtian)–early Eocene (mid Ypresian)

Dicotyledonous angiosperms

DT frequency, 13 million yr DT richness (66.5– 53.5 Ma)

Wilf et al. 2006 Broadly regional: Williston, Great Divide, Greater Green River, Denver, Washakie, Powder River, and Bighorn Basins

22 The fossil record of plant– insect dynamics

31

Late Cretaceous (late Maastrichtian)–early Eocene (mid Ypresian)

Dicotyledonous angiosperms

DT frequency, 23 million yr (66.5– DT richness; 43.5 Ma) leaf‐mining DT frequency, and leaf mining DT richness

Global: South America, North America, Western Europe

Labandeira and Currano 2013

23 Distinguishing Agromyzidae (Diptera) leaf mines in the fossil record: New taxa from the Paleogene of North America and Germany and their evolutionary implications

2

Early Paleocene (Danian, 64.4 Ma); Paleocene–Eocene boundary (Thanetian–Ypresian, 47.8 Ma)

Platanus raynoldsi (Platanaceae); Toddalia schaarschmidti (Rutaceae)

DT frequency

Intercontinental: Montana, USA; Germany

Winkler et al. 2010

16 million yr separation between localities (64.4; 47.8 Ma)

Narrowly regional: (NE Dolomites Region of NE Italy)

Wappler et al. 2015; Labandeira et al, 2016

Ding et al. 2015

(Continued)

Table 24.1 (Continued)

Study†

24 Insect herbivory, plant defense, and early Cenozoic climate change

Floras Ages: period studied and stage‡

6

Plants examined§

Dicotyledenous Paleocene–Eocene angiosperms boundary (Thanetian–Ypresian) to middle Eocene (Lutetian)

Herbivory data recorded¶

Deposit(s) time Deposit(s) spatial duration# extent**

DT frequency, 13 million yr DT richness (56–43 Ma)

Broadly regional: (200 km × 60 km)

References

Wilf et al. 2001

Very long term (>109 yr): multiple locality studies†† 25 Silurian to Triassic plant and hexapod clades and their associations: New data, a review, and interpretations

24

Late Silurian (Ludlow)–Late Triassic (Norian)

Protracheophytes, “rhyniophytes,” zosterophyllophytes, “trimerophytes,” pteridiphytes, pteridosperms‡‡, cycadophytes, coniferophytes, ginkgophytes, bennettitaleans

FFG richness, DT richness

205 million yr Global (425–220 Ma)

Labandeira 2006b

26 Deep‐time patterns of tissue consumption by terrestrial arthropod herbivores

59

Late Silurian (Ludlow)–Late Neogene (Pliocene)

“Polysporangiophytes,” “rhyniophytes,” “zosterophyllophytes,” lycopsids, pteridiphytes, stem spermatophytes§§, gymnosperms, angiosperms

FFG richness, DT richness (by plant tissue)

425 million yr (425–0 Ma)

Global

Labandeira 2013b

27 The four phases of plant–insect associations in deep time

180

Late Silurian (Prídolí)–Late Neogene (late Pleistocene)

Protracheophytes, “trimerophytes,” Rhyniopsida, Lycopsida, Zosterophyllopsida, pteridiphytes, stem spermatophytes§§, gymnosperms, angiosperms

FFG richness, 417 million yr (417–0 Ma) DT richness (by plant taxa)

Global

Labandeira 2006a

28 The origin of herbivory on land: Initial patterns of plant tissue consumption by arthropods

26

Late Silurian (Prídolí)–Late Pennsylvanian (late Kasimovian)

29 Evidence for outbreaks from the fossil record of insect herbivory

4

Gigantopterid, voltzian Early Permian conifer, dicotyledonous (Sakmarian), Late angiosperms Triassic (Carnian), Late Cretaceous (Maastrichtian), early Paleocene (Danian)

Herbivorized 221 million yr (285–64 Ma) surface area, leaf‐mine DT frequency, DT frequency

Global: Texas, North Dakota, Montana, South Africa

Labandeira 2012

30 The insect herbivores of Ginkgo and its fossil antecedents: 230 million years in the evolution of component communities

12

Late Triassic (Carnian)–Recent

230 million yr FFG (230–0 Ma) frequency, FFG richness, DT frequency, DT richness

Global: United States, Canada, South Africa, China, United Kingdom

Labandeira et al., unpublished data

Presence/ “Polysporangiophytes,” absence of “zosterophyllophytes,” lycopsids, pteridophytes, herbivory progymnosperms

Ginkgo biloba and its fossil ginkgogalean predecessors

114 million yr Global (417–303 Ma)

Labandeira 2007a

* These studies are based on research originating from the Labandeira Lab at the National Museum of Natural History in Washington DC; the Wilf Lab at Pennsylvania State University in University Park, Pennsylvania; and the Wappler Lab at the University of Bonn in Bonn, Germany. † The study is provided as the title of the relevant article. ‡ The geochronology is from Gradstein et al. (2012). § Plant taxa designations, both formal and informal, are from the original publication, without updating. ¶ DT, damage type; FFG, functional feeding group; from Labandeira et al. (2007). Wilf and Labandeira (1999) give the first use of this method. # Time durations represented by deposits are estimates based on sediment‐accumulation rates in analogous modern deposits and the thicknesses of the deposits examined. ** For regional sites, the spatial extent of multiple localities is taken as the best fit of a rectangle that encompasses all studied localities, expressed in kilometers for the length (longer dimension) and width (shorter dimension). †† Discussions related to short‐term, medium‐term, long‐term, and very‐long term intervals of plant–insect interactions are in the main text. Deposits are ordinated by age within each time‐interval category. ‡‡ Pteridosperms are commonly known as seed ferns, and collectively include informally the gymnospermous medullosans, peltasperms, corystosperms, caytonialeans and possibly gigantopterids and glossopterids. §§ Spermatophytes are commonly known as seed plants, and consist of gymnosperms and angiosperms. ¶¶ These 16 localities constitute a single megasite, for which all data were pooled. ## Dunne et al. (2014) provide an explanation of food‐web indices.

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Insect Biodiversity: Science and Society

deposits. Typically, spatial scales in such studies are more or less congruent with analogous tem­ poral scales mentioned above. Table 24.1 shows relevant data of 30 studies affiliated with the Labandeira (National Museum of Natural History), Wilf (Pennsylvania State University), Wappler (Bonn University), and Ren (Capital Norman University) laboratories that have quantitatively analyzed plant–arthropod inter­ action studies at short‐term, medium‐term, long‐term, and very‐long‐term time scales. 24.2.4.1  Short‐Term Studies

Short‐term plant–insect associational studies sample single localities that represent less than 106 years and frequently are in the range of 102– 104 years to occasionally 105 years for fluvial and deltaic (major river distributary) environ­ ments. However, finely stratified lacustrine deposits are more highly resolved, typically rep­ resenting 100–103 years (Behrensmeyer and Hook 1992). For typical fluvial environments, the most common accessibility to the fossil record are excavations that range from “holes in the ground” to more methodically dug quarries using quadrat methods. Examples of such stud­ ies include the early Permian (Cisuralian) of north‐central Texas (Table 24.1, Studies 2–5), in which the damage types (DTs) (Labandeira et al. 2007), are used to evaluate the frequency and richness of insect damage and foliage surface area removed by insect herbivory (Beck and Labandeira 1998; Labandeira and Allen 2007; Schachat et  al. 2014, 2015). The Late Permian (Lopingian) of South Africa (Table 24.1, Study 6; Prevec et al. 2009) is another context where plant–arthropod interactions at individual sites are being evaluated to eventually determine from multilocality data the ecological effects of the massive biodiversity crisis at the end of the Permian (Labandeira 2006b, Prevec et al. 2009, Labandeira et  al. 2016a). For some localities, short‐term plant–insect associations are sam­ pled but are not further included in longer‐ term, multilocality studies. Examples of such studies include the variety of arthropod herbi­ vores on a particularly  well‐preserved, early,

Middle Devonian liverwort species from New York state (Table 24.1, Study 1; Labandeira et al. 2013); a late Paleocene megalocality demon­ strating the extent of herbivory on an early Neotropical forest (Table  24.1, Study 7; Wing et  al., 2009); and a food‐web reconstruction resulting principally from abundant DT data that recreated well‐preserved subtropical forest and lake food webs from the early Eocene of Messel, in central Germany (Table 24.1, Study 8) (Dunne et al. 2014). The Messel study shows that the scaling up of individual plant– insect associational data can be used to detect competition and multitrophic relationships in modern, local food webs (Bascompte et al. 2003, Morris et al. 2004). 24.2.4.2  Intermediate‐Term Studies

Intermediate‐term studies range from 106 to 107 years and involve multiple sites that sample temporal shifts and spatial changes in patterns of herbivory, typically as a consequence of geo­ logically short‐lived events. Such short‐lived events notably include extinctions with short rebound intervals, such as the fates of plant– insect associations subjected to mass extinc­ tion, or alternatively, transient, major periods of climate change with significant effects on eco­ logical relationships. Intermediate‐term studies of plant–arthropod associations have been important in several important ways (Table 24.1, Studies 9–16). One example is the impact that the K–Pg ecological crisis and mass extinction had on the types and levels of insect herbivory in the Williston Basin of North Dakota in the United States (Table 24.1, Studies 9, 10). That evaluation included generalized and specialized DTs (Labandeira et  al. 2002a, 2002b), and the negative consequences of the extinction on plant–insect relationships and ecological diver­ sity immediately after the boundary. An approximately similar study was made in Western Europe, but failed to explicitly demon­ strate any effect of a major extinction event on ecological diversity, unlike the pattern in western North America (Table 24.1, Study 11) (Wappler et  al. 2009). Another series of assessments

24  The Fossil History of Insect Diversity

(Table  24.1, Studies 12–14) explored the effect that the PETM, a geochronologically instanta­ neous spike of increased atmospheric tempera­ tures and CO2 content at 56 mya, had on the type and level of insect herbivory during and after the event (Fig. 24.7) (Wilf and Labandeira 1999; Currano et al. 2008, 2010). The response of plants to these dramatic levels of herbivory was partitioned into poorly defended and “hyperherbivorized” deciduous hosts, poorly defended and less‐herbivorized hosts, and ever­ green hosts that were physically and presumably chemically defended (Fig. 24.8) (Labandeira 2007b). Another investigation (Table 24.1, Study 15) probed the effect that the much more grad­ ual but equally intense multiple maxima of early to middle Eocene global temperature and CO2 increases had on insect‐herbivore patterns (Wilf and Labandeira 1999, Wappler et  al. 2012). A biogeographical comparison (Table 24.1, Study 16) of insect herbivory levels between plant hosts in approximately contemporaneous early to middle Eocene floras from South America and North America, occupying the same habi­ tat, indicated that the Neotropics was a signifi­ cantly greater source of biodiversity than was the Palearctic (Wilf et  al. 2005). As a conse­ quence of these studies involving durations of approximately 1–10 million years, there now is evidence that insect herbivores respond to phys­ ical and biological perturbations at time inter­ vals that are sufficiently long to allow robust ecological feedback between cause and effect. One biological process that has been insuffi­ ciently evaluated involves assessments of insect‐ herbivore diversity before, during, and after the major angiosperm ecological expansion of the mid‐Cretaceous (Labandeira 2014b) (Fig. 24.5). The origin of the grassland biome during the mid‐Cenozoic (Strömberg 2011) can provide another, currently unstudied, example of the response of insect diversity to a major biological event that occurred variously in time and space. 24.2.4.3  Long‐Term Studies

Long‐term studies range from 107 to 108 years and document broad evolutionary patterns of

plant–insect interactions. These studies involve multiple localities that are well separated in time and space. The duration of analysis usually is toward the shorter time ranges, involving 10–35 million years, and often somewhat shorter intervals. Assessments such as Studies 17 and 18 document a 16‐million‐year‐long, early Permian interval involving the expansion of herbivory in fluvially dominated habitats of north‐central Texas (Labandeira 2013a; Schachat et  al. 2014, 2015; Table 24.1). Other examples, such as Studies 21 and 22, as well as the intermediate‐term Study 11, document the ecological imbalance of food webs surviving the K–Pg crisis and the heavy toll that the event had on specialized plant–insect interactions, such as the demise of most leaf‐mining taxa (Wilf et al. 2006, Labandeira and Currano 2013, Donovan et  al. 2014). This event affected associational diversity well into the late Paleocene 9 million years later (Table 24.1). The most effective use of long‐term studies records the effects on insect herbivores of major changes in the origin, immigration, or transformation of regional flo­ ras as a result of major, environment‐altering global events. Other long‐term studies have been conducted on phenomena such as the con­ sequences of the P–Tr ecological crisis (Erwin 2006), which have been inferred from patterns of insect‐herbivore damage in a 45‐million‐year interval before, during, and after the event (Table 24.1, Study 19) (Labandeira et al. 2016a). 24.2.4.4  Very Long‐Term Studies

Intervals of very‐long‐term studies greater than 108 years effectively encompass the past 100 to ca. 420 million years, and are used to evaluate the broadest ambit of sampling for terrestrial plant–arthropod associations. The context for such studies is the increase in biodiversity through the colonization of the continental realm by microorganisms and multicellular plant, fungal, and animal life, particularly insects. The two most comprehensive examina­ tions are (i) analyses of the feeding diversity of terrestrial herbivorous arthropod (overwhelm­ ingly insect) lineages as they invaded, colonized,

747

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Insect Biodiversity: Science and Society HF MF S SF G PS M E5 Allophylus flexifolia E1 dicot sp. WW006 E1 Fabaceae sp. WW002

4

E1 dicot sp. WW005 E5 Alnus sp. E5 Lauraceae sp. WW061 E1 dicot sp. WW004 E1 Populus wyomingiana E5 Aleurites fremontensis P4 Betulaceae sp. FU744 P3 Averrhoites affinis E1 Fabaceae sp. WW007 E1 Fabaceae sp. WW001 E5 “Dombeya” novi-mundi E5 Platycarya castaneopsis

2

B

P3 Betulaceae sp. FU741 E4 Macginitiea gracilis E1 Populus wyomingiana P3 Cercidiphyllum genetrix P3 Zizyphoides flabella E3 “Ampelopsis” acerifolia P3 Juglandaceae sp. FU740 E5 dicot sp. WW052 P4 Fabaceae sp. FU750 E3 Lauraceae sp. WW036 P4 Cercidiphyllum genetrix E4 Fabaceae sp. WW040 E4 Fabaceae sp. WW007 E3 Averrhoites affinis

3

A

E3 Averrhoites affinis E3 “Dombeya” novi-mundi P4 Macginitiea gracilis P2 Browniea serrata E3 dicot sp. WW037 P3 Davidia antiqua P4 dicot sp. FU745 P4 dicot sp. FU749 E3 dicot sp. WW034 P3 Platanus raynoldsi P3 Macginitiea gracilis E2 Alnus sp. P1 dicot sp. SC1 P4 Platanus raynoldsi P1 Cercidiphyllum genetrix P2 Zizyphoides flabella P2 Cercidiphyllum genetrix P2 “Ampelopsis” acerifolia

1

P2 Platanus raynoldsi P1 Platanus raynoldsi P1 Browniea serrata P2 Davidia antiqua P2 Persites argutus P2 “Ficus” artocarpoides P2 “Celtis” peracuminata

24  The Fossil History of Insect Diversity

became established, and accessed food resources from vascular plant‐host lineages (Table 24.1, Study 27) (Labandeira 2006a) (Fig. 24.9); and (ii) the same herbivore lineages acquiring food from the 14 principal tissue types of vascular plant lineages (Table 24.1, Study 26) (Labandeira 2013a). Both studies represent the same 420‐ million‐year interval of time, of which the last 115 million years (27%) documents the expan­ sion of angiosperm hosts and their arthropod herbivores, indicating a geologically rapid pro­ cess of diversification at multiple trophic levels (Fig. 24.10). Four other important assessments at very‐long‐term time scales are (i) lineages of colonizing arthropod herbivores displaying lag times of up to 98 million years for the eventual herbivorization of particular, mid‐Paleozoic plant organs and tissues (Table 24.1, Study 28) (Labandeira 2007a); (ii) the early expansion of arthropod herbivory on a variety of emerging Permian plant lineages as they contended with the P–Tr ecological crisis and became re‐ established during the Triassic (Table 24.1, Study 25) (Labandeira 2006b); (iii) the four best‐ case examples for pest outbreaks in the fossil record, ranging in age from the Early Permian to Early Paleocene (Table 24.1, Study 29) (Labandeira 2012); and (iv) the tracking through 12 slices of time from the Late Triassic to the Recent of the herbivore‐component communi­ ties of a ginkgoalean species belonging to a well‐ documented, 230‐million‐year‐long fossil record (Table 24.1, Study 30). Very‐long‐term studies  such as these provide the coarsest‐

grained analyses of the manifold ways that arthropod herbivores have partitioned their food‐resource world and, in the process, have generated an astonishing level of interactional and associational diversity. 24.2.5  Morphological Diversity

Morphological diversity is best expressed as disparity, which is the range of anatomical, molecular or other structurally based form of a particular group or evolutionary lineage of insects. A wide range of techniques, morpho­ logical attributes, and modes of presentation document the morphological disparity of fossil (and modern) insects. One approach is to measure or express quantitatively single struc­ tures, such as wings, to express size and shape dis­parity by techniques such as landmark or Fourier analyses. Complex multi‐element structures such as mouthparts, however, invar­ iably require categorization by phenetic char­ acters and character‐states for adequate description (Labandeira 1990, 1997), prin­ cipally because of the difficulty of achieving robust descriptions based on analyses of standard measurements. Mouthparts, legs, and genitalia are ideal multi‐element features for revealing structural disparity. Another way of expressing morphological disparity is to exam­ ine the fundamental genotypic roots of dispar­ ity rather than concentrate on its observable phenotypic results. Developmental disparity hints at the sources of morphological disparity,

Figure 24.7  Ecological diversity 4. Differences in insect herbivore diversity before (gray) and after (white) the Paleocene–Eocene Thermal Maximum (PETM) in the Bighorn Basin of Wyoming, United States. The diversity shift is shown by a cluster analysis of insect damage on species‐site pairs, based on the relative abundances of the seven functional feeding groups and 71 damage types (DTs) occurring on those pairs. Each plant host has a minimum of 20 leaf specimens included in the analysis. Leaf morphotypes that have not been formally named have been tagged with Fort Union (FU) or Wildwood (WW) Formation prefixes. Black circles are scaled based on their relative abundances for each functional feeding category on each plant‐host species. The clusters 1, 2, 3A, and 3B are explained by a general pattern of galling, piercing and sucking, and mining DTs having significantly greater abundance on host plants from the warmer Eocene sites. Feeding categories: HF, hole feeding; MF, margin feeding; S, skeletonization; SF, surface feeding; G, galling; PS, piercing and sucking; M, leaf mining. More details are given by Currano et al. (2010). Adapted from Currano et al. (2010), copyright by the Ecological Society of America.

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Insect Biodiversity: Science and Society

(a)

(b) Number of damage types (bootstrapped)

750

(c)

(d)

14 Macginitiea Bulk

Populus wilmattae

12 10

Cedrelospermunm

Allophylus

8 “Caesalpinia”

6

Rhus 4

Parvileguminophyllum

Cardiospermum 2 0

0

20

40

60

80

100

Number of leaves

Figure 24.8  Ecological diversity 5. Plant–insect interactions are used for documenting the diversity of antiherbivore defense strategies in insect lineages during the Early Cenozoic Thermal Maximum. (a) Populus wilmattae (Salicaceae), a member of the hyperherbivorized accommodationist cohort. (b) Bootstrapped insect damage diversity for host‐plant species with 25 or more specimens from the early Middle Eocene (43 Ma) Green River Formation of northeastern Utah, the earliest of three time slices assessed by Wilf et al. (2001). (c) Cedrelospermum nervosum (Ulmaceae), a member of the herbivorized accommodationist cohort. (d) Parvileguminophyllum coloradoensis, a member of the highly defended cohort. Additional details are given by Wilf et al. (2001) and Labandeira (2007b). From Labandeira (2007b), copyright SEPM (Society for Sedimentary Geology), Tulsa, Oklahoma, USA; used with permission.

24  The Fossil History of Insect Diversity Functional feeding groups

ch Ep o

M

a Pe rio d

External foliage feeding Leaf mining Borings Rhizophagy Piercing & sucking Galling Palynophagy (generalized) Oviposition Seed predation Fluid feeding Mixed–four or more

Figure 24.9  Ecological diversity 6. The fossil history and diversity of terrestrial arthropod feeding on plant tissues. With the exception of early occurrences during the latest Silurian to Early Pennsylvanian, the documentation for hexapod consumption of tissue types is not exhaustive. A legend of hexapod dietary modes is shown at the upper left, and five basic tissue systems in land plants at top. Reference sources and descriptions of the type of data used for this chart are provided by Labandeira (2013b). Used with permission, © Springer‐Verlag, Berlin, Heidelberg.

751

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Insect Biodiversity: Science and Society

Paleogene

Cretaceous

Neogene Miocene

Oligo

1 2 3 4 5 6 7

Eocene

Late

Paleo

Early

0 Ma

50

100

Period Epoch

Functional feeding groups External foliage feeding Piercing-and-sucking Boring Leaf mining Galling Seed predation Oviposition

8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 Interval

Nymphaeales Arecales Juncales Poales Zingiberales Chloranthiales Magnoliales Laurales Ranunculales Proteales Vitales Caryophyllales Saxifragales Malpighiales Fabales Rosales Malvales Myrtales Fagales Sapindales Cornales Ericales Garryales Gentianales Lamiales Aquifoliales Dipsicales Asterales Unknown

Plant hosts

Odonatoptera Orthopteroidea Hemiptera Hymenoptera Coleoptera Diptera Lepidoptera Acari

Inferred arthropod herbivores

Foliage feeding Piercing-sucking Boring Leaf mining Galling Seed predation Oviposition

Functional feeding groups

78–88 68–77 63–67 54–62 49–53 46–48 43–45 38–42 35–37 27–34 22–26 18–21 17 15–16 13–14

11–12 10 7–9 5–6 3–4 2 1

Biota

24  The Fossil History of Insect Diversity

and increasingly is used to understand the evolution of morphology in insects – not only in model organisms such as Schistocerca, Tri­ bolium, Drosophila, and Manduca, but in rel­ evant, non‐model organisms. Recently, in the course of exploring trace‐fossil assemblages in the fossil record, disparity has been used to explore the extended phenotype of behavior, particularly in ascertaining how insects mod­ ify their sedimentary environment through burrows, tracks, and trails, termed ich­­no­logic disparity, or “ichnodisparity” (Bua­ tois and Mángano 2013). 24.2.5.1  Size Disparity

Wings are perhaps the best example of insect size and shape disparity. Wings can explain much about the biology of insect locomotion, and there are several fundamental modes that describe how insects fly (Dudley 2000). One noteworthy study is an assessment of disparity trends in insect body size, which is correlated to wing length, for the 323‐million‐year history of pterygote insects (Clapham and Karr 2012). Maximum wing lengths were calculated for each of 32 ten‐million‐year‐long time intervals that provided a trend line indicating the rapid and early increase in wing lengths (and thus body size) under varying atmospheric O2 levels for the insect fossil record. From the Late Pennsylvanian to Middle Permian, wingspans ranged up to 70 cm for one Early Permian inter­ val, and the robust correlation between wing length and atmospheric O2 during the Late Carboniferous continued into the Late Jurassic (Fig. 24.11). This trend was followed by a reset­ ting of maximum wing length during the

Cretaceous and into the Cenozoic to considera­ bly less than half of the Paleozoic value. When wing length was compared to atmospheric O2 concentrations of the past (Berner 2009), it was concluded that during the first 150 million years of insect evolution, body size was linked to elevated levels of atmospheric O2. Maximum insect body size was correlated with a maxi­ mum atmospheric concentration of 33% O2 at the same Permian interval mentioned above. (The present atmospheric concentration of O2 is 21%.) After this phase, during the Cretaceous and Cenozoic, there was decoupling of body size with atmospheric O2 levels. During this lat­ ter interval, body size likely was controlled by aerial predators such as birds during the latter half of the Mesozoic and Cenozoic, and possibly bats during the Cenozoic (Fig. 24.11). 24.2.5.2  Structural Disparity

One of the best examples of structural disparity in insects is their mouthparts. As feeding is a primary task of insects, considerable informa­ tion regarding the life habits of insects is bound up in their mouthpart structure. Insect mouth­ parts consist of about 36 broad categories of structural divergence (Fig. 24.12) and serve as morphological proxies for a wide variety of feed­ ing strategies (Labandeira 1997). In contrast to the record of insect family‐level diversification, which is characterized by a semilogarithmic, constantly increasing rate expressed as a “hol­ low curve” (Fig. 24.1a), mouthpart disparity is typified by an inverse, S‐shaped logistic trend, whereby most new mouthpart types are pre­ sent relatively early in insect history, and then plateau at mid‐history (Labandeira and Sepkoski

Figure 24.10  Ecological diversity 7. The expansion of insect herbivores with angiosperms, showing the diversity of feeding associations. The distribution of functional feeding groups is illustrated within specified biotas (bottom), on accompanying plant hosts within successive, specified, five‐million‐year intervals (at top) from the mid‐Early Cretaceous to the Recent. Inferred insect herbivores are provided in a middle panel that links functional feeding groups with plant hosts within each interval. The arrow refers to the early associational biota, the Early Cretaceous Dakota Formation of the mid‐continental United States, which documents the early record of this expansion phase. Labandeira (2006a) gives additional details. Adapted from Labandeira (2006a), under the Attribution‐Share Alike Creative Commons License of the Facultat de Geologia, Universitat Autònoma de Barcelona.

753

200 150 100 0

50

Wing length (mm)

250

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Insect Biodiversity: Science and Society

Permian

Triassic

Jurassic

Cretaceous

Paleogene

Neog

10

20

30

Carbonif

Atmos. pO2 (%)

754

300

250

200

150 Age

(106

100

50

0

yr)

Figure 24.11  Morphological diversity 1. Phanerozoic trends in insect wing length at top (Clapham and Karr 2012), with atmospheric environmental oxygen concentration (pO2), using the GEOCARBSULF model (Berner 2009), at bottom. The maximum size for each 10‐million‐year bin that contains more than 50 measurements is demarcated by a black line. From Clapham and Karr (2012), used by permission of the National Academy of Sciences, USA.

1993). The contrast between these two patterns of taxic diversity versus structural disparity indi­ cates that most new morphologies, such as the complex feeding structures of mouthparts, occur early in insect history. One of the generators of biodiversity in insects is evolutionary convergence, whereby the same structure recurs multiple times in unrelated lineages. The origin and develop­ ment of a distinctive mouthpart type associated with the long‐proboscid pollination syndrome is one of the best examples of the origination of evolutionary convergence in the fossil record. The multiple origins of long‐proboscid mouth­ parts also are an object lesson about how mor­ phologically based biodiversity is generated in

deep time. Modern examples of the long‐pro­ boscid syndrome famously include the 28‐cm‐ long tubular proboscis of the hawk moth that pollinates a Madagascaran orchid (Nilsson et al. 1987), and species of nemestrinid, tabanid, and apiocerid flies that pollinate deep‐throated members of the Geraniaceae (geraniums) and Iridaceae (irises) in the Cape Floristic Province of South Africa (Manning and Goldblatt 1996, Morita 2008). Other examples abound from other orders of modern insects (Labandeira 2010). Nevertheless, the long‐proboscid mouthpart structure has a history that extends deep into the mid‐Mesozoic, back to the small, earliest representative from the mid‐Permian of Russia (Labandeira 2010). Long‐proboscid

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Figure 24.12  Morphological diversity 2. The deep‐time distribution of 34 extant mouthpart classes (left panel), their mouthpart class (central column) and assigned mouthpart group (right column), including two extinct classes, from a phenetic analysis of insect mouthparts (Labandeira 1990). Solid black segments represent the occurrence of mouthparts as body fossils in well‐preserved deposits; gray shade indicates their presence based on sister‐group relationships; white segments indicate indirect evidence for the presence of a mouthpart type, including trace‐fossil evidence and the inferred presence of the mouthpart type in one life stage (e.g., larva) when fossils of another life stage (e.g., adult) are present for the taxon in question. Details are provided by Labandeira (1997). Adapted from Labandeira (1997), copyright © 1997 by Annual Reviews, used with permission.

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mouthparts have originated minimally 13 times during the mid‐Mesozoic in association with pollination of a variety of gymnosperms (Fig.  24.13). These originations in­­ clude three independent origins in Neuroptera, four to more

likely five originations in Mecop­tera, at least four originations in Diptera, and evidently one in Lepidoptera (Ren et  al. 2009, Labandeira 2010, Yang et al. 2014, Peñalver et al. 2015, Lin et al. 2016).

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Figure 24.13  Morphological diversity 3. The convergently acquired long‐proboscid pollination mode in six separate lineages of preangiospermous, mid‐Mesozoic insects from the Middle Jurassic Jiulongshan Formation and mid‐Early Cretaceous Yixian Formation in northeastern China. (a) Lichnomesopsyche gloriae of the Mesopsychidae (Order Mecoptera). (b) Pseudopolycentropus janeannae (Mecoptera: Pseudopolycentropodidae). (c) Jeholopsyche liaoningensis (Mecoptera: Aneuretopsychidae). (d) Florinemestrius pulcherrimus (Diptera: Nemestrinidae). (e) Protonemestrius jurassicus (Diptera: Nemestrinidae). (f ) Kalligramma aciedentatus (Neuroptera: Kalligrammatidae). From Labandeira and Currano (2013, fig. 6), copyright © 2013 by Annual Reviews, used with permission.

24  The Fossil History of Insect Diversity

24.2.5.3  Developmental Disparity

Although developmental diversity often has been considered a separate category of biodiver­ sity (Erwin 2008), it also can be viewed within the broader ambit of morphological disparity. Developmental disparity has only recently been recognized (e.g., Davidson 2006), and has been formalized in a number of ways. Developmental disparity is deployed in two common modes. First is the conservation of genes typically across multiple lineages in a major clade (Raff 1996). An example is the genes that provide for an ancestral, efficient tracheal respiratory system that allowed for the incredible success of hexa­ pods (Alexander 1971). A second mode is recombination of genes to provide a new devel­ opmental morphology  –  and its phenotypic expression  –  within a single, derived lineage. One example is the abdominal spring mecha­ nism in collembolans (Konopova and Akam 2014), which likely occurred during the Late Silurian, not long after acquisition of respiratory tracheae. A closely related, second example simi­ lar to the collembolan abdominal spring com­ plex, but with wider‐ranging implications, is the origin of abdominal appendage patterning in holometabolous insect larvae and adults. This process extended the diversity of abdominal appendage occurrence, number, size, structure, and specialization in holometabolous insects that initially began as a full complement of paired, leg‐like abdominal appendages with minimal Hox‐gene regulation (Fig. 24.14). A third example of this second mode of providing dramatic developmental changes involves the diversification of long‐proboscid mouthparts in four orders of insects (Fig. 24.13) that occurred during the mid‐Mesozoic (Labandeira and Currano 2013). This repeated developmental process of long‐proboscid mouthpart formation likely is attributable to a common genetic mechanism in the ancestral adult holometabolan, probably similar to that described by Rogers and Kaufman (1997) for the evolution of elongate piercing‐and‐sucking mouthparts. This process probably represents a latent gene system responsible for the anatomic

merging of paired labial appendages (glossae and paraglossae) or paired maxillary append­ ages (galeae, possibly palps) into conjoined tubular structures able to imbibe fluid plant products (Lin et al. 2016). Also demonstrating the importance of the co‐optation of evolution­ ary‐developmental structures is a fourth exam­ ple involving wing spots and eyespots in extinct, mid‐Mesozoic kalligrammatid lacewings (Fig. 24.15). The observed pattern represents a gene system responsible for the developmental ori­ gin of mid‐Mesozoic kalligrammatid lacewing spots and eyespots (Labandeira et  al. 2016c), and the convergence of near‐identical spot and wing‐eyespot patterns with nymphalid butter­ flies that first appeared approximately 60 mil­ lion years later during the Paleogene Period (Oliver et  al. 2012). These four examples of genetic mechanisms reveal the profusion of ways that developmental disparity is a funda­ mental source for the generation of insect diversity. 24.2.5.4  Key Innovations

Most structural novelties do not rise to the level of key innovations (Gao et al. 2016). Occasionally, a particular, newly arising morphological feature can be a key innovation, and consequently can be a source of morphological diversity (Wagner 2011). Indeed, the four evolutionary develop­ mental features mentioned above each can be considered a key innovation that expanded the possibilities for new life‐habits and evolutionary diversification, although appropriate tests would be required for confirmation. At a fundamental level, metamorphosis probably is the most suc­ cessful key innovation, responsible for the diver­ sification of the Insecta (Rainford et  al. 2014, Haug et al. 2015), defined as the clade that bears the egg to larva to pupa to adult developmental sequence. The success of this key innovation already was present during the Early Penn­ sylvanian to Permian interval, 323 to 252 mya (Labandeira 2011, Haug et  al. 2015), in which the  earliest lineages of Holometabola (Laban­ deira 2011) (Fig. 24.14) began to displace con­temporaneous non‐holometabolous insects.

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(a)

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Figure 24.14  Morphological diversity 4. (a) Fossil evidence for the earliest Holometabola, placed within a broad phylogenetic context. The earliest reliable occurrences of taxa (solid dots, followed by a thick, black vertical line) are found in various sources (Labandeira 2011, Haug et al. 2015). The horizontal stippled bar at bottom represents the initial diversification and earliest fossil occurrences of holometabolan insects in the fossil record. (b) A recent reconstruction of the earliest known holometabolous larva, Srokalarva berthei, based on Haug et al. (2015). The black indicates sclerites; the gray indicates softer, more weakly sclerotized regions between sclerites. White lines designate possible intersegmental junctures that have been preserved. (c) Variation in expression domains of abdominal Hox genes in the larvae of the earliest Holometabola (S. berthei (i) and Metabolarva bella (ii)) and various modern taxa. The most closely related model species affiliated with the earliest Holometabola is probably the beetle Tribolium castaneum; its wild‐type pattern of Hox gene development shown in (iii), and the expression pattern of an Ultrabithorax (Ubx)/Abdominal‐A (Abd‐A) mutant is shown in (iv). A sample of the Hox gene effects on abdominal leglet development is provided in (v–vii), showing the variety of expression patterns on abdominal appendages in taxa of Hymenoptera, Diptera, and Lepidoptera, respectively. The expression domains of Distalless (Dll) and the Hox genes Ubx, Abd‐A, and Abd‐B in holometabolous larvae is shown in (viii). The gray‐scale hues represent, from anterior to posterior: black, antennae and mouthparts; medium gray, thorax and abdomen; dark gray, thoracic legs; darkest gray, abdominal leglets; and light gray, cerci. Sources: (a) from Labandeira (2011), reproduced by permission of the New York Entomological Society; (b,c) from Haug et al. (2015), used under terms of the Creative Commons Attribution 4.0 international license (http://creativecommons.org/publicdomain/ zero/1.0/).

24  The Fossil History of Insect Diversity

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Figure 24.15  Morphological diversity 5. The diversity of wing spots and eyespots in Mesozoic Kalligrammatidae (Neuroptera), with a phylogenetic context and comparisons to modern Lepidoptera. (a) The most parsimonious tree of the Kalligrammatidae phylogeny, with the right forewing eyespot/spot condition mapped onto terminal clades, indicating likely wing spot and eyespot origins. Wing eyespot‐ and spot‐type symbols are designated at the upper left; crosses are wing eyespot or spot absences. Examples (b–f ) are right forewings with wing eyespots or spots from mid Mesozoic Kalligrammatidae and modern Psychopsidae (g). (b) Type 1 eyespot. (c) Type 2 eyespot. (d) Type 3 eyespot. (e) Type 4 eyespot. (f ) Two Type 5 spots (Kalligrammatidae). (g) Two Type 5 spots (modern Psychopsidae). Examples (h–k) represent eyespots and spots from modern Lepidoptera. (h) Type 6 eyespots. (i) Multiple Type 5 spots. (j) Absence of wing spots or eyespots. (k) Enlargement of modern pigmentation of the owl butterfly showing a pigmentation pattern similar to Type 2 and 3 eyespots in mid Mesozoic Kalligrammatidae. Additional details are given by Labandeira et al. (2016c). Unrestricted use of Labandeira et al. (2016) is provided by terms of the Creative Commons Attribution, under a license of the Royal Society.

Another key innovation is the parasitoid life habit that commenced during the mid‐Early Jurassic (Labandeira 2002b), particularly among  apocri­ tan Hymenoptera and brachyceran Diptera. This novel feeding guild later was supplemented by

the oldest fossil evidence of animal parasitoidism by fungi (Hughes et al. 2010), which supports a Late Jurassic–Early Cretaceous origin for this macabre fungus–arthropod relationship (Sung et al. 2008) (Fig. 24.4).

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24.2.6  Functional Diversity

Functional diversity is the range of particular operations that insects accomplish or the mechanical roles that they serve in communi­ ties or ecosystems. The measurement of func­ tional diversity has always been challenging (Erwin 2008), and historically has depended on the particular role that a group of species has assumed. The fossil record documents sev­ eral types of functional diversity, and three approaches are presented below that involve assessments of functional diversity through time. The first example is the record of herbi­ vore‐feeding guilds as they compete for and partition food resources through time. The sec­ ond instance is the extension in space and time of various, sediment‐based life histories as insects have colonized novel substrate ecospace in lacustrine ecosystems. The third is the evolu­ tion of insect trophic roles in food webs, and in particular the consequences for food‐web func­ tion from the emergence of the parasitoid guild during the mid‐Mesozoic. These three aspects of insect functional diversity share similar fea­ tures, such as competition and partitioning of resources through time. 24.2.6.1  Functional Feeding Groups

No aspect of insect functional diversity is better understood than their feeding biology. One way of understanding feeding diversity in deep time is through insect functional feeding groups and their DTs. A functional feeding group is the broader category of insect‐feeding style that is defined by how an insect feeds, such as external foliage feeding, piercing and sucking, galling, leaf mining, seed predation, and wood boring (Labandeira et  al. 2007). Functional feeding groups are subdivided into fundamental, diag­ nosable, explicitly defined occurrences of insect‐mediated damage, or DTs, which are the common currency by which insect damage is qualitatively and quantitatively assessed in the fossil and modern records. In ecosystems from the Middle Devonian to the present, studies of the diversity of functional

feeding groups and damage types assess various patterns in deep time, such as plant‐host tissue use through time (Labandeira 2013b), the iden­ tities of the major herbivorized plant‐host taxa through time (Labandeira 2006a), the role of global climate change across regional land­ scapes (Wilf and Labandeira 1999, Wilf et  al. 2001), herbivory in site‐specific habitats (Prevec et  al. 2009, Wing et  al. 2009, Schachat et  al. 2014), and the evolution of a component com­ munity on a group of architecturally identical plant hosts (Ding et al. 2015). Also studied are shifts in herbivory at major extinction events (Labandeira et  al. 2002b, Wilf et  al. 2006, Donovan et  al. 2014) and during intervals of prolonged global climate change (Currano et al. 2008, 2010; Wappler et  al. 2009). In modern ecosystems, DT diversity has been used to assess latitudinal patterns of herbivory intensity and richness (Adams and Zhang 2009), the rela­ tionship between DT and predator richness in wet tropical forest (Bachelot and Kobe 2013), the establishment of a correlation between DT richness and respective insect‐herbivore diver­ sity (Carvalho et al. 2014), and the consequences of global climate change on insect herbivory (Blois et al. 2013). 24.2.6.2  Lacustrine Ecospace Occupation

One way of understanding the evolution of insect functional diversity involved the degree to which ecospace is occupied through time. One such process was the evolution of diversity in freshwater lotic (moving water) and lentic (still water) ecosystems and in and on their sedi­ mentary substrates. An evaluation of this pro­ cess is done through the measure of ichnodisparity during the crucial interval from the Late Triassic to the Early Cretaceous, a time interval that encompassed the prelude and cul­ mination of the Mesozoic Lacustrine Revolution (MLR). During the Permian and into the Jurassic, a shallow tier of taxonomically depau­ perate, burrowing infauna, as well as track‐ and trail‐making epifauna, occupied underused ecospace in freshwater biotas along lake mar­ gins (Miller and Labandeira 2002).

24  The Fossil History of Insect Diversity

Profundal depths in lake settings, however, bear more penetrative burrows, indicating greater substrate tiering – that is, occupation by organisms of discrete sediment layers  –  of deeper infaunal communities. Lake‐margin and profundal settings were colonized exclusively by detritivores and consisted of simple food‐ web structures (Sinitshenkova and Zherikhin 1996). During the Late Jurassic to Early Cretaceous, depending on locality, there was a trophic transformation of ichnodisparity in lacustrine communities such that sediment bio­ turbation, tiering, and an increase in track and trail activity dramatically increased in all benthic lacustrine environments. This major increase in functional diversity, or ichnodispar­ ity, indicates that greater metabolic activity and diversification in lacustrine habitats were asso­ ciated with a functional shift from detritivore‐ to herbivore‐dominated food webs that created the MLR (Buatois et  al. 2016). The increase in functional roles associated with the MLR resulted in a sizable extension in the carrying capacity of lacustrine ecosystems and signifi­ cant increases in the diversity of their inhabiting organisms, including infauna, epifauna, plank­ ton, nekton, neuston, and consumers of live plant tissues (Zherikhin and Sinitshenkova 2002). 24.2.6.3  Parasitoids and Trophic Roles in Food Webs

The relatively new field of fossil food‐web recon­ struction has combined exceptional, well‐pre­ served data from particular fossil deposits (Dunne et  al. 2008) with modern, quantitative techniques of food‐web analyses (Camacho et al. 2002, Stouffer 2007). A recent study established the food web from the noted 47‐million‐year‐old Middle Eocene site at Messel, Germany (Dunne et  al. 2014), based on exceptionally well‐pre­ served trophic data (Labandeira and Dunne 2014) (Fig. 24.6). One of the conclusions from this study was the importance of parasitoid insect taxa that linked consumers to trophically lower levels in the terrestrial portion of the food web, but not for the aquatic food‐web, which remained predator driven.

In modern terrestrial ecosystems, consumer taxa consist of pathogens, predators, parasites, and parasitoids, but as the Messel (Dunne et al. 2014) and modern webs (Memmott et al. 1994, 2000) demonstrate, insect parasitoids were the key consumers in an Eocene terrestrial ecosys­ tem, just as they are in extant ecosystems. Assess­ments of food‐web structure from a vari­ ety of modern ecological contexts has consider­ ably underrepresented the roles of parasitoids in contributing to food‐web structure (Dunne et  al. 2014). For the Eocene Messel food web, this issue has been brought to the fore: the top consumer of the terrestrial portion of the food web was a parasitoid fly, which contrasted with the lacustrine part of the same overall food web, in which the top consumer was a large croco­ dilian (Dunne et  al. 2014). The emergence of insect parasitoids overwhelmingly dominates the third, great tier of consumers in terrestrial food webs and is where the dominant repository of insect‐consuming diversity is lodged (Mem­ mott and Godfray 1993, Lafferty et  al. 2006). The origin of this guild is attributable to a major diversification event during the Jurassic and Early Cretaceous (Labandeira 2002b), which had a major effect on the efficiency of food webs in the deep past. The ecological significance of parasitoid functional diversity has been as an efficient, consumer‐driven, top‐to‐down regula­ tor of herbivory, present since the mid‐Mesozoic, which differs from earlier, pre‐Jurassic, bottom‐ to‐up regulation of herbivores that were limited by plant‐resource allocation (Labandeira 2002b). 24.2.7  Behavioral Diversity

Behavioral diversity consists of how insect indi­ viduals or social groups possess unit cohesion, respond to external stimuli such as predators, or behave in a particular environmental context, such as camouflage or pollen‐ or foliage‐foraging strategies. The types of behavioral diversity are quite varied, and several aspects of behavioral diversity are explored below. Features such as the degree of sociality, types of close resem­ blance of one organism to another organism or

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inanimate object, and pollination strategy are all aspects of behavioral diversity. Although Erwin (2008) indicated that one common perception of behavioral diversity is that it is “irretrievably lost” in the fossil record, data from the sedimen­ tary ichnological and plant–insect association records retain significant behavioral diversity from the deep past. 24.2.7.1 Sociality

There is a significant body‐fossil history of eusocial insect clades, such as termites, ants, and bees, as well as subsocial clades, notably some lineages of beetles (Grimaldi and Engel 2005). An important part of this history is the trace‐fossil record of these same insect groups where crucial evidence for behavior is available from nest and tunnel networks in subterranean sedimentary substrates (Duringer et  al. 2007); from borings in wood and adjacent tissues con­ taining entry and exit burrows, nuptial cham­ bers, congregation galleries, larval tunnels, and pupal cells (Boucot and Poinar 2010); and from particular types of insect damage on plant tis­ sues (Labandeira and Prevec 2014). These cate­ gories of evidence typically point to a type of behavior assignable to a particular group of insect fabricators. Insect lineages with varying degrees of social­ ity produce complex ichnologic structures and plant‐associational damage records that can yield significant evidence for behavioral diversity (Genise et al. 2000, Labandeira 2007b). Subsocial clades, such as some ambrosia and bark beetles associated with complex borings in wood, pro­ vide a variety of evidence that indicates parental care and caste structure, implying various degrees of sociality (Labandeira et  al. 2001). Eusocial termites and ants also create stereo­ typed and complex nests with galleries, tunnels, and special chambers in sedimentary substrates and in woods that occasionally are preserved (Francis and Harland 2006, Roberts and Tapanila 2006, Duringer et al. 2007). In addition, leafcut­ ter bees leave precise, circular excision holes on leaves; these leaf discs are used as food resources and for the construction of larval chambers for

pupation (Sarzetti et al. 2008). Zombie ants leave distinctive death‐grip marks on the undersides of leaves, caused by the gripping of major veins with their mandibles, thus revealing behaviors resulting from the attack of this social species by a fungal parasitoid (Hughes et  al. 2010). Eusociality generally is not related to an increase in diversity when compared to a non‐eusocial sister clade, a pattern that is particularly notice­ able for bees (Engel 2001). 24.2.7.2  Mimicry and Warning Coloration

In modern ecosystems, a significant amount of biodiversity is connected with mimicry (Meyer 2006). Mimicry results when an organism (the mimic) closely resembles another, similarly con­ structed organism or a non‐living feature (the model), resulting in a high degree of similarity in appearance, behavior, sound, smell, or position in the environment of the two or more entities (King et al. 2006). As a general designation, mim­ icry can refer to a variety of phe­nomena, includ­ ing true mimicry, camouflage, mimesis, and crypsis, as well as eyespots that are not necessar­ ily attributable to any particular model. Several recent studies have identified multiple examples of fossil mimicry, including crypsis in leaf insects from the German Middle Eocene (Wedmann et al. 2007), foliar mimesis between the wings of a saucrosmyline lacewing (Neuroptera) and either cycadalean or bennettitalean pinnules from the end of the Middle Jurassic of northeast­ ern China (Fig. 24.16a–e) (Wang et al. 2010), and whole‐body mimicry between a hangingfly (Mecoptera) and an entire  Ginkgoites leaf from the same locality (Fig. 24.16f) (Wang et al. 2012). In the case of wing eyespots among 17 species of kalligrammatid lacewings (Neuroptera) from the late Middle Jurassic to mid‐Early Cretaceous of Eurasia, there is a heightened diversity of species with wing‐spot and especially wing‐eyespot pat­ terns that are nearly identical in structure to those of extant nymphalid butterflies (Lepidoptera) (Labandeira et al. 2016c). The pos­ session of predator‐deterring wing eyespots (Labandeira et al. 2016c) apparently was associ­ ated with elevated diversity levels of the

24  The Fossil History of Insect Diversity (a)

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Figure 24.16  Behavioral diversity 1. Examples of the diversity of insect mimesis involving cycadophyte and ginkgophyte models from the late Middle Jurassic of northeastern China. (a) The insect mimic Bellinympha filicivora (Neuroptera: Saucrosmylidae), with left wing re‐ illustrated in (b) for comparison to cycad foliage models. Potential co‐occurring plant models for this mimic from the Middle Jurassic Jiulongshan Formation of northeastern China are shown in (c–e), with (e) perhaps indicating the closest fit. (c) Holozamites sp. (Cycadales). (d) An unnamed cycadophyte. (e) Nilssonia sp. (Cycadales). Wang et al. (2010) provide additional details. (f ) A reconstruction (© Wang Chen of Capital Normal University) of the mimesis between Ginkgoites leaves of Yimaia capituliformis and the hangingfly Juracimbrophlebia ginkgofolia, from the Jiulongshan Formation. Wang et al. (2012) give additional details. Permission for reproduction of parts (a–f ) granted by the National Academy of Sciences, USA and Wang Chen of Capital Normal University.

Kalligrammatidae and their likely higher levels of speciation (Fig. 24.15). 24.2.7.3  Pollen‐Collection Strategies

One fundamental evolutionary feature of polli­ nating insects such as bees is the behavioral pat­ terns that are revealed during the course of their  deep‐time history. One such feature is stereotyped patterns of pollination‐collection strategies, such as foraging specialization (Cane

and Sipes 2006). Two major extinct lineages that represent six species of pollen‐basket‐bearing electrapine bees from the early to Middle Eocene of Germany were examined for pollen on their exquisitely preserved bodies, including their metatibial pollen baskets (Wappler et  al. 2015). The results indicated that a twofold pat­ tern of pollen‐collection strategies simultane­ ously was present as they foraged on the local flora. Pollen on the bee’s bodies, except for their

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pollen baskets, was ­collected in a generalized, incidental, and random manner, plastered on their heads, thoraces, and abdomens. Such pol­ len was destined to provide a pollination service for the ambient, local flora. By contrast, pollen occurring on the legs, particularly pollen bas­ kets, reflected a much narrower, restricted suite of pollen taxa from evergreen hosts with a spe­ cific, uniform flower structure (e.g., Parker et al.

2015). Eventually, this subset of collected spe­ cialized pollen was delivered as larval food to the hive. This dual foraging pattern (Fig. 24.17) probably was responsible for the increased diversity of pollen‐basket‐bearing bees, as this clade possesses a dual pollen‐collection strat­ egy, whereas bees lacking pollen baskets possess a single pollen‐collecting behavior and are less diverse (Michener 2007).

Thorax Abdomen

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Figure 24.17  Behavioral diversity 2. A plot of the first two orthogonal component axes with positions of pollen‐load variables of plant families versus bee‐body regions. The bees (Apidae) represent two genera from an extinct tribe of bees and the pollens are from recent taxa of mostly dicot plant families. Note the taxonomically broad spectrum of collected pollen on the heads (lightest gray), thoraces (dark gray), and abdomens (medium gray) of the bee bodies, showing a polylectic collection syndrome. This stands in contrast to the taxonomically narrow range of pollen on the legs (light gray), particularly in the pollen baskets, of the same two bee genera, representing a narrow (monolectic) pollen collection syndrome. Source: Wappler et al. (2015). Reproduced under an open‐access agreement with Cell Press, © by Elsevier, Inc.

24  The Fossil History of Insect Diversity

24.3 ­Biodiversity Changes Through Time Two basic modes of change affect biodiversity in the fossil record: (i) prolonged intervals of envi­ ronmental perturbation and (ii) sudden and cat­ astrophic events. Each of these modes of change has two components that affect biodiversity. First is the event that caused dramatic change in biodiversity. Second is the resulting aftermath, which also results in biodiversity change. For diverse continental organisms with a significant fossil record, such as land plants, arthropods, and fungi, both types of sudden and prolonged change can have a redounding effect on the future course of diversity contraction or expan­ sion. For insects, the taxonomic, taxic, ecologi­ cal, morphological, functional, and behavioral components of diversity are dramatically affected by these two basic forms of change. 24.3.1  Long‐Term Environmental Change

Global biodiversity is affected by long‐term change through its protracted modification of the physical environment, or by important bio­ logical events that profoundly restructure the basic relationships of life (Schopf 1983). The dis­ tinction between physical and biological causal­ ity is murky, however, as long‐term environmental change often has both physical and biological aspects. One example is the late Paleozoic expan­ sion of herbivore diversity, which likely was accentuated by glacial–­ interglacial cyclicity (Schachat and Labandeira 2015, Schachat et  al. 2015). However, other examples, such as rapid biodiversity increases associated with the Parasitoid Revolution and the early expansion of angiosperms, seem to be overwhelmingly or sin­ gularly attributable to a particular biological development, such as, respectively, the invention of the parasitoid ovipositor and the origin of the flower. Both of these developments were innova­ tions that spurred major biodiversity increases. Several such prolonged changes have resulted in major diversity increases in insects through time.

24.3.1.1  Mid‐Paleozoic Beginnings of Terrestrial Ecosystems

The earliest macroscopic terrestrial ecosystems consisted of a Lilliputian world in which subaerial life emerged as land plants, microarthropods, and fungi, through various pathways, formed simple food webs along a gradually vegetated coastline (Little 1983). Major exceptions to this dwarfed world were the giant columnar fungus Prototaxites (Hueber 2001) and perhaps the arthropleurid myriapod Eoarthropleura (Wilson and Shear 1999). The earliest, well‐documented ecosystem is the Rhynie Biota (Fayers and Trewin 2004) of Early Devonian age from southeastern Scotland, interpreted as a hot‐spring deposit that was less diverse than modern counterparts. These and other Late Silurian and Devonian ecosystems have modest taxonomic diversities that consist of several major plant and arthropod–herbivore lin­ eages (Laban­ deira 2006b), with only a few arthropod–­herbivore groups that were consum­ ing a fraction of the available plant tissues (Fig. 24.9). The Paleozoic pattern of arthropod her­ bivory suggests major lags between the appear­ ance of plant tissues and their eventual consumption by arthropod herbivores (Labandeira 2007a). Although site‐specific taxonomic diversi­ ties were low, ecological diversity and morpho­ logical disparity of the arthropod component might have been moderate. (Truly terrestrialized vertebrates, by contrast, did not appear in conti­ nental ecosystems until the end of the Devonian.) Little is known of the functional diversity of Devonian organisms, although a liverwort species from the Middle Devonian of New York state shows evidence of three herbivore feeding mecha­ nisms: external foliage feeding, piercing and suck­ ing, and galling (Labandeira et al. 2013). Virtually nothing is known of behavioral diversity in these earliest of terrestrial ecosystems. 24.3.1.2  Initial Taxic Radiation of Insects

The single continental insects in the  course

greatest biodiversity event for the realm was the initial radiation of the Pennsylvanian Period over of approximately 24 million years

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between 323 and 299 mya. For insects, the Pennsylvanian Period represented the first sig­ nificant uptick in dominance of any major ani­ mal group in terrestrial ecosystems, as measured by total family‐level diversity (Fig. 24.1a), origi­ nation (Fig. 24.1b) (Labandeira 2005b), includ­ ing the first appearances of 12 orders of insects (Labandeira 1999, Grimaldi and Engel 2005). Major insect lineages that have their first appear­ ances during this interval include zygentomans, odonatopterans, paleodictyopteroids, ephemer­ opterans, plecopteroids, orthopteroids, hemip­ teroids, and, as studies during the past decade have attested, holometabolans (Fig. 24.14) (Nel et al. 2013, Haug et al. 2015). This last clade, the Holometabola, had cryptic beginnings during the Pennsylvanian and Permian (Fig. 24.14a), and their larvae were important exophytic and endophytic herbivores (Fig. 24.14b). Much of early holometabolan evolution was molded by developmental evolutionary genetics that allowed a bewildering diversity of mouthpart types (Labandeira 1997), abdominal appendage expression (Haug et al. 2015) (Fig. 24.14c), flight mechanisms (Dudley 2000), and other struc­ tures (Kristensen 1999). The Holometabola con­ tinued their diversification during the Permian, particularly herbivorous and predaceous forms, invading a wide variety of habitats. By the end of the Triassic, almost all of the modern ordinallevel lineages were present (Fig. 24.2), rendering the lineage the most diverse multicellular clade of all time. Today, the Holometabola are the most taxically diverse element and the dominant trophic player in terrestrial and freshwater eco­ systems (Kristensen 1999), although the group has never successfully invaded open marine habitats. The Holometabola also display ele­ vated levels of morphological disparity and functional and behavioral diversity. 24.3.1.3  Late Paleozoic Expansion of Herbivore Functional Feeding Groups

During the Middle Pennsylvanian, most equato­ rial habitats of the paleocontinents straddling the paleoequator, particularly Euramerica, had coal‐swamp biotas. The iconic elements of these

biotas were pole‐like lycopod and horsetail trees, a variety of architecturally diverse seed plants, large dragonfly, paleodictyopteroid, and orthop­ teroid insects, and an occasional amphibian‐ grade vertebrate, barely emergent on land. In this ecosystem, insect herbivory initially expanded, most of it documented from the exceptionally well‐preserved coal‐ball and shale deposits of the Illinois Basin (Labandeira and Phillips 1996a, 1996b, 2002; Labandeira 1998a, 2001). A significant turnover in the floras at the  Middle Pennsylvanian–Late Pennsylvanian boundary resulted in a successor tree‐fern‐dom­ inated biota that supported an increase in the intensity and diversity of herbivory. One particu­ lar Late Pennsylvanian, Illinois Basin source plant, the marattialean tree fern Psaronius ­chasei, bore a variety of arthropod detritivores and insect herbivores that constituted the earliest diverse‐component community documented from the fossil record (Labandeira 1998b). The insect functional feeding groups of P. chasei included mite detritivores, unknown root feed­ ers, cockroach pith borers, a holometabolous insect galler, paleodictyopteroid piercers and suckers, and orthopteroid feeders on foliage and sporangia (Labandeira 1998a, 1998b, 2001). It seems that all major plant organs and accessible tissue types were consumed by these herbivores, while hard tissues, such as sclerenchymatous root mantle, were bored and consumed by detri­ tivorous oribatid mites (Labandeira et al. 1997). Later, during the early Permian, herbivory further expanded onto significantly different ­ assemblages of plants from north‐central Texas. These four assemblages largely contain ferns, horsetails, and mostly extinct seed‐plant taxa such as medullosans, peltasperms, cycado­ phytes, gigantopterids, and early conifers (Beck and Labandeira 1998; Labandeira and Allen 2007; Schachat et  al. 2014, 2015; Schachat and Labandeira 2015). The environments from which insect herbivory has been recorded at these sites typically are associated with a variety of fluvial habitats, and the vegetation was sub­ strated on mineral soils, rather than the peats of the earlier clastic swamps, such as the Illinois

24  The Fossil History of Insect Diversity

Basin to the northeast. Accordingly, there was an expansion in the diversity of herbivory (as DTs) inflicted on plant hosts from the Texan early Permian (Fig. 24.9), perhaps reflecting an increase in mouthpart types (Fig. 24.12), but no  change in the functional feeding groups that  were established during the earlier Late Pennsylvanian. Diversity and levels of insect‐ mediated damage have been quantified for these Texan early Permian floras, and strongly indi­ cate that the insect damage disproportionately targeted seed‐plant hosts, was variable across habitats, and overall was about one‐third that of modern levels (Schachat et al. 2015). These data differ from those of a study by Turcotte et  al. (2014), who indicated that early Permian her­ bivory levels might be closer to modern levels than formerly thought.

of sedimentary substrates) and especially the deployment of more complex and abundant behavioral diversity in a variety of lacustrine hab­ itats (Buatois and Mángano 2013). Toward the end of this increase in behavioral diversity, there is substantial behavioral convergence across lake‐margin and fully lacustrine habitats in which phylogenetically unrelated lineages of insects and other arthropods ecologically, functionally, and behaviorally arrived at the same solutions for occupation and exploitation of the benthos, epib­ enthos, plankton, nekton, neuston, and a variety of herbivore niches (Buatois and Mángano 2013). The MLR was a significant driver of freshwater insect diversity, as detritivore, herbivore, preda­ tor, and other niches were established along lake margins and deeper lacustrine zones.

24.3.1.4  Ecological and Behavioral Changes from the Mesozoic Lacustrine Revolution

The Parasitoid Revolution had a considerable effect on terrestrial insect diversity, particularly during the Jurassic and Cretaceous, when an explosion of families of apocritan Hymenoptera, and to a lesser extent brachyceran Diptera, entered the newly created parasitoid feeding guild. Insect parasitoids did not exist before the Jurassic, and insect consumption of insects was accomplished either by predation if immediate death of the prey ensued or possibly by rare parasitism if the host survived. The novel feeding and dietary mode of parasitoidism uniquely combined elements of pre­ dation and parasitism and is associated in modern ecosystems with significantly elevated diversities of not only parasitoid species, but also their troph­ ically lower herbivore hosts and trophically higher consumers such as hyperparasitoids (Memmott et  al. 1994, 2000). Although there has been an increase in mid‐Mesozoic to modern diversity of parasitoid insects (Labandeira 2002b), there also has been a heightened diversification of parasitoid fungi that similarly have resulted in insect victims (Fig. 24.4a). For example, during the Late Cretaceous, some insect lineages likely became zombified by a process initiated by the e­ volutionary radiation of the fungal parasitoid lineage Ophiocordycipitaceae (Sung et  al. 2008) (Fig.  24.4b,c). By Middle Eocene times, fungal

Within the continental realm, freshwater ecosys­ tems have always been taxonomically and eco­ logically less diverse than terrestrial ecosystems (Cohen 2003). Nowhere is this more evident than in the exceedingly delayed time lags of certain aquatic life habits occurring in freshwater habi­ tats when compared to terrestrial habitats (Miller and Labandeira 2002). (This delay in taxic, taxo­ nomic, ecological, and behavioral diversity also is true when the freshwater realm is contrasted with that of the marine realm (Vermeij and Lindberg 2000, Buatois and Mángano 2011).) The most notable of these delays was herbivory, which was well established by Pennsylvanian time in terrestrial habitats (Labandeira 1998a), but took approximately 200 million years longer to be comparably established in shallow, lake‐ margin habitats (Fig. 24.18a), and a longer delay for an equivalent presence in deeper, fully lacus­ trine habitats (Fig. 24.18b) (Buatois et al. 2016). This delay represents a protracted process rather than a singular event, and occurred on different paleocontinents at various times during the mid‐ Mesozoic (Buatois et al. 2016). The same delayed timespans are recorded for infaunalization (the degree of burrowing and trace‐fossil penetration

24.3.1.5  The Parasitoid Revolution

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(a)

Lake margin

(b)

Fully lacustrine

Figure 24.18  Functional diversity 1. Phanerozoic changes in bioturbation in lake‐margin and fully subaqueous lacustrine deposits resulting in the diversification of herbivorous, mostly insect clades during the mid‐Triassic to mid‐Cretaceous, as a result of the Mesozoic Lacustrine Revolution in lake‐margin habitats (a). The effects of the Mesozoic Lacustrine Revolution appeared later, during the Jurassic–Cretaceous boundary interval in fully lacustrine habitats (b). Additional details are provided by Buatois et al. (2016). This figure is used with permission and is a modification of Buatois et al. (2016).

­ arasitoid lineages also targeted ant hosts (Hughes p et  al. 2011). In addition to the evolution of this novel repertoire of behavioral diversity, the Parasitoid Revolution contributed to a significant increase in taxic diversity, as heightened Hymenoptera family‐level diversity during the mid‐Mesozoic (Fig. 24.2) is accounted for over­ whelmingly by parasitoid family origi­ nations (Labandeira 2002b). Parasitoids also enhanced mid‐Mesozoic morphological diversity through an increase in mouthpart classes, partly attributa­ ble to newly originating parasitoid ­ lineages

(Labandeira 1990) (Fig. 24.12). The introduction of the new parasitoid functional feeding group came to dominate many ecosystems, as seen in modern food webs (Valladares et  al. 2001, Lafferty 2006). 24.3.1.6  Biodiversity Ramifications of the Early Expansion of Angiosperms

The frequently used estimate of extant angio­ sperm species richness is 250,000 (Gaston and Spicer 2004), although values as high as 400,000 appear more reasonable (Govaerts 2001). This

24  The Fossil History of Insect Diversity

likely high estimate is approximately a fourth that of extant, described insect diversity, indi­ cating a coupling of the diversities of these two most hyperdiverse and prominent continental macro‐organismic groups. This linkage is but­ tressed by a massive literature of plant–insect interactions (Price et al. 2011), which is particu­ larly rich in herbivory studies (Futuyma and Agrawal 2009, Bagchi et al. 2014, Turcotte et al. 2014). As the preeminent continental organis­ mic groups, angiosperms and insects form a bewildering array of agonistic, neutral, and mutualistic relationships; but angiosperms are a relatively recent group in land‐plant evolution, initially diversifying sometime during the Early Cretaceous, based on fossil (Friis et  al. 2011) and time‐calibrated molecular phylogenetic analyses (Magallón 2010). Put in an appropriate deep‐time perspective, angiosperms are about 30% as old as land plants, the latter of which have their earliest fossil occurrences in the mid‐ Silurian, about 430 mya. From the evidence of several studies men­ tioned earlier, the general pattern of initial angi­ osperm diversification did not induce a parallel diversification with insects, as judged by the flat trend in family‐level taxic diversity throughout the Cretaceous (Fig. 24.1a). This trend also is supported by stasis in family‐level diversity for  such plant‐associated insect taxa as the Coleoptera and Lepidoptera during the initial angiosperm radiation interval (Fig. 24.2). The absence of a parallel increase in angiosperm and insect diversification is even more marked when plant‐associated insect families and their plant‐ host affiliations and shifts (i.e., fern/cryptogam only, fern–cryptogam to angiosperm transition, gymnosperm only, gymnosperm to angiosperm transition, angiosperm only) are taken into con­ sideration through a 174‐million‐year‐long his­ tory that encompasses the Jurassic, Cretaceous, and Paleogene periods (Fig. 24.5). This study (Labandeira 2014b) indicated that earlier insect lineages with gymnosperm hosts were replaced by later insect lineages with newly acquired angiosperm hosts, resulting in significant time‐ lag effects. This changeover produced a flat

trend in phytophagous insect diversity asso­ ciated with the switch to an angiosperm‐­ dominated global flora. This effect also is seen in the relationships among angiosperm hosts and their inferred plant‐associated arthropod taxa beginning during the mid‐Late Cretaceous (Fig. 24.10). Post‐Cretaceous floras are consid­ erably richer in the ordinal diversity of insect‐ consumed plant hosts as well as the ordinal diversity of their inferred arthropod herbivores and their membership in a greater number of functional feeding groups (Labandeira 2006a). The cautionary conclusion of these studies is that the biodiversity relationships between two presumably coevolved groups might not repre­ sent tandem codiversification; rather, in the case of angiosperms and insects, dampening factors caused stasis in the initial diversity level of plant‐associated insect lineages for tens of mil­ lions of years before a new global equilibrium was established. 24.3.1.7  Expansion of the Grassland Biome

Although grasses (Order Poales) were present at the end of the Cretaceous in dinosaur copro­ lites (Prasad et al. 2011), a distinctive grassland biome did not originate until a few tens of mil­ lions of years later. The expansion of the grass­ land biome occurred during the late Oligocene to Middle Miocene in a time interval from about 26 to 14 mya, depending on the particu­ lar continent (Strömberg 2011). During this interval, insect communities that were ecologi­ cally linked to grasses began to evolve. The numerous grass–insect interactions that were established at this time were varied and likely included aphid transmission of the predeces­ sor of barley dwarf virus, which causes bar­ ley  yellow dwarf disease in cereal grains (Labandeira and Prevec 2014); the association between dung beetles and the feces of large grazing ungulates for the creation of brood balls in soil‐tunnel and gallery networks (Genise et  al. 2000, Sánchez et  al. 2010); and  leaf‐mining interactions between agro­ myzid flies and their poaceous hosts (Winkler et al. 2010). Although the e­ stablishment of the

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e­cologically well ­ integrated grassland biome took approximately 12 million years as it expanded across vast continental interiors, there apparently were time lags in the evolu­ tion of grass–insect associations, just as there were in the development of high‐crowned den­ tal batteries in grazing artiodactyls and peris­ sodactyls to accommodate the tooth‐abrading effects of silica‐rich grasses (Strömberg 2011). Once the grassland biome was present, insect herbivory on the Poales declined, as measured in the percentage of removed foliar surface area (Turcotte et  al. 2014). This trend also is reflected in a comparison of herbivory on Poales versus other angiosperm orders (Fig. 24.10), attribu­table as well to the antiherbivore effects of grass  foliar architecture and basal meristematic growth. The data tentatively sug­ gest that the initial expansion of the grassland biome is associated with an increased richness of plant–insect interactions, but not necessar­ ily an increase in herbivory levels. 24.3.2  Short‐Term Environmental Change

In addition to long‐term environmental change, global biodiversity is affected by sudden, cata­ strophic events that can dramatically alter the future course of life. Most notable of these are the five great Phanerozoic extinction events and associated ecological crises that have affected life during the past 541 million years (Raup 1986). For the biodiversity of continental organ­ isms, such as plants and insects, the two most relevant and important of these events are the P–Tr and K–Pg events (Erwin 2006, Nichols and Johnson 2008). These two global events had repercussions up and down the taxonomic hier­ archy, resulting in major alterations of food‐web structure and considerable reductions of mor­ phological disparity, and they placed constraints on or eliminated much of the functional and behavioral diversity of continental organisms. Such major and sudden events have been attrib­ uted to physical changes rather than biological causes such as pathogenic wipeouts. These

events involve either terrestrial triggers, for example, the dramatic changes in global tem­ perature associated with the Pleistocene extinc­ tion, or extraterrestrial causes, including a 10‐km‐diameter bolide impact that eliminated much of terrestrial life at the end of the Cretaceous Period. Estimated durations for the deployment of these geochronologically instan­ taneous events range from hours to days for the K–Pg event (Labandeira et al. 2016b), to 103 years for the major effect of the PETM (Currano et al. 2008) and the end‐Pleistocene extinctions (Martin 1984), and perhaps 103–105 years for the destruction involved in the P–Tr event (Erwin 2006). In all instances, there were signifi­ cant changes in biodiversity. 24.3.2.1  Permian–Triassic Global Crisis and Reductions in Biodiversity

The single most devastating ecological crisis in the history of life was the P–Tr event, per­ haps represented by a major event at the end of the Permian Period 252 mya (Erwin 2006). Marine ecosystems were devastated (Alroy et al. 2008), and the biological consequences of the P–Tr event were also far reaching in conti­ nental ecosystems, involving extinction of the  overwhelming majority of insect taxa (Labandeira 2005b) and their associations with plants (Labandeira et al. 2016a), the deci­ mation of ecosystem structure  (Miller and Labandeira 2002, Labandeira 2006b), severe reductions in morphological disparity and functional diversity, and the imposition of a major bottleneck on behavioral diversity. This elimination of a significant portion of terres­ trial biodiversity was abetted by a deteriora­ tion of the continental physical environment, including depressed atmospheric O2 levels (Berner 2009), deteriorating continental con­ ditions (Looy et al. 2001), and the imposition of acidic freshwater systems (Twichett 2006). These processes caused the loss of much bio­ logical history embedded in the genetic port­ folios of the deceased species (Erwin 2008).

24  The Fossil History of Insect Diversity

The Permian ecological crisis essentially reset the clock for a previously expanding biotal diversification process, such that each of the five types of biodiversity gradually re‐evolved in major ways in the wake of the event’s after­ math and as the recuperation process started noticeably during the Middle Triassic, contin­ uing into the mid‐Mesozoic (Looy et al. 2001, Hochuli et  al. 2010, Botha and Smith 2006). The most significant evidence for recovery comes from Late Triassic biotas, where the contrast is the starkest with the earliest Triassic interval immediately after the Permian. Several effects that pertain to the devastation of insect diversity at the P–Tr event are note­ worthy. In addition to global taxic and site‐­ specific taxonomic losses during the Early Triassic, there was a major restructuring of ter­ restrial ecosystems to much simpler food webs (Twichett 2006, Visscher et  al. 2011), although there are alternative views (Wardle et al. 2011). This ecological change restricted opportunities for insect invasion of new niches. In freshwater ecosystems, there was initial contraction but subsequent expansion of functional diversity; later on in the Triassic, there was an expansion of substrate‐use strategies and accompany­ ing  insect behaviors in lacustrine ecosystems (Fig. 24.18) (Buatois et al. 2016). Morphological disparity became severely limited after the eco­ logical bottleneck at the end of the Permian, evi­ denced by a decrease in average insect size, as measured by wingspan lengths (Fig. 24.11; Clapham and Karr 2012), and the absence of new Triassic mouthpart types (Fig. 24.12; Labandeira 1997), suggesting stasis in mouthpart disparity and the absence of broad‐scale feeding innova­ tions. The related pattern of increased packing of insect functional feeding groups consuming particular tissue types (Fig. 24.9) followed the P–Tr event (Labandeira 2013b), suggesting the repackaging of insect feeding strategies. A full return to the Late‐Permian levels of plant–insect associational diversity apparently took about 25 to 30 million years, and diversity levels did not re‐equilibrate until the Late Triassic.

24.3.2.2  Cretaceous–Paleogene Global Crisis and Reductions in Biodiversity

The end‐Cretaceous (K–Pg) ecological crisis occurred 66 mya and had a far greater effect on the ecological state of affairs than on the major insect lineages, which were far less affected than in the catastrophic P–Tr event (Labandeira et al. 2016b). But the recentness of the K–Pg event means that its impact on the fossil record is simi­ lar to that of the far older P–Tr event. Although the taxonomic and taxic reductions of the P–Tr event were far greater than that of the K–Pg event, the latter still left a significant imbalance in ecological communities in its wake, revealed in evaluations of insect‐mediated damage at two North American communities within about 1 million years of the event (Wilf et al. 2006). This reduction in ecological scope also is demonstrated by the significant decline in angiosperm–insect associations when evaluated globally at the ordinal level (Fig. 24.10) (Labandeira 2006a), and by the greater packing of insect functional feeding groups into plant component communities, following the K–Pg event (Fig. 24.9; Labandeira 2013b). For inten­ sively studied floras across the K–Pg boundary, there is evidence for a major reduction of spe­ cialized insect associations (Labandeira et  al. 2002b), although the pattern is less clear for Western Europe (Wappler et al. 2009). This con­ clusion is buttressed by the extinction of leaf‐ mining associations and their angiosperm hosts after the K–Pg event and corresponding subse­ quent time lags for the establishment of other leaf‐mining associations (Lopez‐Vaamonde et  al. 2006; Donovan et  al. 2014). Evidently, by the middle Eocene, this trophic imbalance was alleviated, as data from the Lake Messel food web indicate (Fig. 24.6), with the development of modern terrestrial and lacustrine food webs (Dunne et al. 2014, Labandeira and Dunne 2014). These ecological data document a significant reduction in associational diversity (Labandeira et al. 2002b), which is at odds with the absence of family‐level taxic reductions in insect diversity during the K–Pg mass extinction

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(Labandeira and Sepkoski 1993). This distinc­ tion is true whether the family‐level taxic data are evaluated by total insect diversity (Fig. 24.1a), by insect order (Fig. 24.2), or by the diversification rates of certain insect lineages (Fig. 24.3) (Song and Liang 2013). Unlike the P– Tr event, there are no detectable post‐event reductions in insect size (Fig. 24.11) (Clapham and Karr 2012), in numbers of mouthpart types (Fig. 24.12) (Labandeira 1997), or behaviors of substrate use in lacustrine ecosystems (Fig. 24.18) (Buatois et  al. 2016). Consequently, the  K–Pg event, unlike the P–Tr event, is a case  where significant changes in insect eco­ logical diversity were largely decoupled from unchanged insect taxic diversity. 24.3.2.3  Biodiversity Realignments During the Paleocene–Eocene Thermal Maximum

Ecological diversity of insects can track short‐ term climate change in deep time (Wilf and Labandeira 1999, Wilf et al. 2001, Currano et al. 2010). These studies demonstrate that insect herbivores express a heightened feeding response during times of transient increases in temperature and atmospheric CO2 concentra­ tions (Fig.  24.7, Fig. 24.8) that typically occur within longer greenhouse intervals (Currano et  al. 2010). During such climatological spikes, insect‐herbivore populations and species expand as latitudinal belts characterized by temperature increases shift poleward. One such event, the PETM, occurred at the Paleocene–Eocene boundary 56 mya, during a period of elevated temperatures and CO2 levels that was present for 105 yr, and associated with a near‐complete turnover of floras at mid‐latitudinal localities in western North America. At these localities, resulting invasions of insect herbivores tracked migrating floras poleward during major global climate‐change conditions. In addition, social insect groups became ecologically linked to newly developed her­ bivore communities and included termites (Engel et  al. 2009), ants (Moreau et  al. 2006), and bees (Wappler et  al. 2015). Once long‐term global climate‐change conditions were established at Paleogene mid‐­

latitudes, there were major insect diversity rea­ lignments of insect herbivores. 24.3.2.4  End‐Pleistocene Extinctions and Their Meaning for the Modern World

The Pleistocene Epoch toward the end of the Neogene Period was the prelude for evolutionary and ecological processes that characterize our own Holocene Epoch. The Pleistocene was typi­ fied by several major glacial and interglacial intervals, each representing a major climato­ logical shift that resulted in a significant biogeo­ graphic realignment of vertebrate species ranges (Martin 1984). Insects were no exception to these regional biogeographic movements, and such movements are instructive for understanding insect evolutionary biology during the Pleistocene (Elias 1996). Working mostly with beetles from Pleistocene deposits that represent multiple gla­ cial–interglacial cycles, Coope (1978) demon­ strated that insect species essentially did not evolve throughout the 2.6 million years of the Pleistocene, whereas the environments inhabited by those beetle species underwent profound physical changes. Beetle species responded to these changes by migrating to new regions that matched their former habitats (Coope 1978), of which perhaps the best example is an extant Tibetan beetle species adapted to cool, alpine conditions that formerly occurred in a Pleistocene glacial deposit from Britain (Coope 1973). Another example is the more recent inva­ sion of northern Europe by Mediterranean insect species after the last glaciation (Coope 1990). These observations indicate that one response to the creation of new habitats – even if they pre­ viously existed elsewhere in time and space – is to relocate rather than evolve. As a consequence, the diversity of insect taxa, as illustrated by Coope’s work on beetles, remained relatively sta­ ble, even though many of the local assemblages inhabited by those insects were different from and not ecologically linked to any community of today (Williams and Jackson 2007). The climatological shifts at the end of the Pleistocene eliminated much of the existing vertebrate megafauna and their community ­

24  The Fossil History of Insect Diversity

s­tructure, and megafaunal extinctions (espe­ cially of birds and particularly on islands) car­ ried on, albeit at a lower rate, throughout the Holocene due to the highly altered ecological conditions. However, insects were relatively unaffected, using shifts in biogeographic range distributions as a hedge against extinction and diversity losses.

24.4 ­Current Societal Aspects of Fossil Insect Biodiversity The fossil record of insect biodiversity has features that simultaneously engage human understanding of the world and are instructive to the current situation that faces human society. Several previously mentioned deep‐time pro­ cesses and events that eventually affected the course of human development were the K–Pg ecological crisis, the expansion of the grassland biome, and the end‐Pleistocene extinctions. Five additional aspects of fossil‐insect diversity, men­ tioned below, involve the waxing and waning of biodiversity that impinge on societal concerns of today. These features express current research programs that link the insect‐fossil record and past insect diversity with the joys of understand­ ing insect natural history. But more importantly, these topics address the current environmental dilemma that confronts humankind. 24.4.1  Human Interests and Biases

A fascinating aspect of the insect‐fossil record was the Pennsylvanian–Permian interval of the late Paleozoic during which many myriapod, arachnid, and insect species were sizably larger than their counterparts of the more recent Mesozoic and Cenozoic fossil record. The reason for this gigantism probably involves elevated atmospheric O2 concentrations, which at some points reached 33%, compared with the present oxygen level of 21% (Dudley 2000). One morpho­ logical manifestation of this gigantism was among the paleopteran dragonfly‐like aerial insects, including one paleodictyopteran with a wingspan

of 66  cm from the Late Pennsylvanian Commentry deposit of northern France, and odonatopterans with wingspans extending to 70 cm in the case of one meganeurid species from the early Permian Elmo deposit of central  Kansas, United States (Beckemeyer 2000, Dudley 2000). Other aspects of the insect‐fossil record and biodiversity involve exceptional fossil deposits that contain splendidly preserved insects. Notable mid‐Mesozoic deposits are the Jiulongshan Formation (late Middle Jurassic, 165 mya) and the Yixian Formation (mid‐Early Cretaceous, 125 mya). These deposits provide excellently preserved specimens of long‐pro­ boscid, butterfly‐like kalligrammatid lacewings with differentially pigmented wing eyespots that display evolutionary convergence with Cenozoic butterflies after a 60‐million‐year‐long gap (Yang et  al. 2014, Buchmann 2015, Labandeira et  al. 2016c), and scorpionflies that contain excep­ tional details of mouthpart structures used in the pollination of extinct gymnosperms (Ren et  al. 2009, Labandeira 2010, Lin et  al. 2016). More popular in the public’s mind is amber (Labandeira 2014a; Fig. 24.19), made famous by the movies Jurassic Park and Jurassic World. Although there are several inaccurate statements regarding the paleobiological understanding of amber, such as its geological provenance, presence of DNA, and techniques for extraction of insect inclusions, the films of the Jurassic Park series did instill in the lay audience’s mind the high level of insect pres­ ervation and charm of amber (Labandeira 2014a). 24.4.2  Tools for Understanding Evolutionary and Ecological Diversification

Although examples of fossil‐insect diversity studies are too numerous to recount here, sev­ eral areas of research provide new perspectives toward understanding the relationship between past insect diversity and their relationships with other organisms. One recent example is the often conspicuous insect damage on fossil leaves. Studies of fossil plant–insect interactions currently are experiencing a renaissance in research,  particularly examinations of herbivory

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(Table 24.1), pollination (e.g., Compton et  al. 2010; Labandeira 2010; Peñalver et al. 2012, 2015; Yang et  al. 2014), and mimicry (e.g., Wedmann et  al. 2007; Wang et  al. 2010, 2012; Labandeira et al. 2016c). Other aspects of recent interest in plant–insect interactions include studies of the timing of insect herbivory, such as the delayed response of lepidopteran Gracil­lariidae leaf min­ ers to colonizing their plant hosts during the Late Cretaceous and Paleogene (Lopez‐Vaamonde et al. 2006), and the trophic status of the basal‐ most lineage of the overwhelmingly herbivorous lepidopteran lineage Ditrysia, which apparently consumed animal products such as keratin, hair, and skin (Regier et  al. 2014) rather than being consummate herbivores. Other recent discoveries provide different themes for the importance of fossil‐insect stud­ ies for understanding the exceptional history of insects. The discovery of a fungus–termite association with galleries preserving fungal comb material from Late Miocene–Early Plio­ cene Chad is noteworthy (Duringer et al. 2007). This find documents mesic and humid condi­ tions in the Sahara Region just 5 mya, repre­ senting a different climate regime from today (Sutherland 2003). Tritrophic interactions involving a dicot plant, its herbivore (a zombi­ fied ant), and the herbivore’s parasitoid fungus have been documented in the fossil record (Hughes et al. 2010); such strange parasites have tapped into the public’s fascination with zom­ bies, exemplified by the cult classic Night of the Living Dead. Part of the interest in this study (Hughes et  al. 2010) is that it features dead, zombified ants existing as an extended pheno­ type of the ant‐consuming parasitoid fungus (Anderson et al. 2009). 24.4.3  Detection of Insect‐Borne Diseases in the Fossil Record

Pathogens are a major source of diversity (Bagchi et al. 2014), and insect‐borne pathogens are a major cause of plant disease in modern ecosystems. However, little is known about their

fossil record, their past and present diversity, and how they can be potentially diagnosed in the fossil record. The basic elements of these insect‐borne diseases are the host plant, the pathogen, the environment, and the insect vec­ tor that facilitates the introduction of a particu­ lar pathogen into a plant (Labandeira and Prevec 2014). Invasive insect‐borne pathogens can be viruses, bacteria, fungi, or nematodes, which can elicit a repertoire of responses on the host plant, including necroses, callus‐tissue forma­ tion, and production of an array of defensive compounds that seal off lesions exposed to the environment (Agrios 2005). Some of these modifications of plant‐host tissues can be cate­ gorized into DTs that can be diagnosed in the fossil record. Such tracking of pathogens, their plant hosts, and their potential insect vectors may provide insights into the historical origins of such insect‐borne diseases as barley yellow dwarf disease (a virus), cucurbit bacteria wilt (a  bacterium), Dutch elm disease (a fungus), and  red ring disease of palms (a nematode) (Labandeira and Prevec 2014). Knowledge of extinct host‐plant lineages and the types of host‐plant shifts in these and other insect‐borne diseases can contribute useful approaches toward their current control. On a somewhat human‐focused theme, insect vectors of vertebrate diseases, including those that severely afflict human health, have been found in the fossil record (Poinar 2014). It has been long known that species of Glossina, the current primary insect vector for sleeping sick­ ness and nagana in Eastern Africa, was present in the intermontane west of the United States during the latest Eocene approximately 34 mya (Cockerell 1917). Although these well‐preserved glossinid specimens occur as compression– impression fossils, virtually all other evidence for insect‐borne and pathogen‐borne diseases in the fossil record originates from amber deposits (Labandeira 2014a). One such incidence comes from mid‐Cretaceous Burmese amber, which provides evidence for malaria  –  a debilitating disease consisting of a red‐blood‐cell‐invading

24  The Fossil History of Insect Diversity

apicomplexan parasite that is transmitted by a mosquito. In this case, a 99‐million‐year‐old api­ complexan parasite Paleo­haematopus burmacis was found in the body cavity of a Protoculicoides ceratopogonid midge (Poinar and Telford 2005), suggesting that a type of malaria was being transmitted, probably to a reptilian ultimate host. From more recent Dominican amber, about 21 million years in age, another example was the blood‐feeding phlebotomine sand fly Lutzomyia (Psychodidae), which was associated with the hair of an unknown solenodon, a relict group of Insecti­vora from the Greater Antilles (Peñalver and Grimaldi 2006). Modern Lutzomyia is associated with Insectivora such as solenodons, which host the protistan pathogen Leishmania, the cause of the disfiguring human disease leishmaniasis. Also occurring in Dominican amber was a species of the anophe­ line mosquito Anopheles (Culicidae), females of which bore the same distinctive egg morphol­ ogy, including float structures, as modern malaria‐carrying mosquitoes (Zavortink and Poinar 2000). These and other occurrences (reviewed by Labandeira 2014a) provide circum­ stantial evidence that insect‐ and mite‐borne diseases have ancient histories in non‐human vertebrate hosts, occasionally extending as far back as the mid‐Cretaceous. However, many of the claims need to be substantiated with addi­ tional, more compelling evidence. Nevertheless, emerging human species during the later Neogene, such as Austra­lopithecus afarensis and Homo erectus, likely acquired, through lat­ eral transfer from other vertebrates, some of these long‐standing diseases. 24.4.4  Insect Herbivory and Global Warming

Deep‐time studies indicate that insect‐medi­ ated herbivory resulted as a consequence of global warming and associated elevated levels of atmospheric CO2 (Currano et  al. 2008, 2010). These findings have significant implications for the current climate change crisis (Blois et  al. 2013). The conclusion that insect herbivory sig­

nificantly increases during transient or longer‐ term spikes in temperature and atmospheric CO2 concentrations in the fossil record can pro­ vide estimates regarding the potential duration, affected areas, as well as level of damage expected with the current upward trend in tem­ perature and atmospheric CO2 rise. The mecha­ nism that drives the increase in insect herbivory under conditions like those of the PETM 56 mya and of modern global climate change is largely known. In particular, as CO2 levels rise, nitrogen becomes limiting in plants, inducing increased consumption levels and diversity of damage to plants by insect herbivores to maintain their metabolic homeostasis (Lincoln et  al. 1993, Watt et  al. 1995). However, the relationship might be more complex than formerly appreci­ ated (Stiling and Cornelissen 2007). The pat­ terns of insect herbivory during the PETM and other analogous time intervals have direct application to similar global climate trajectories occurring today. 24.4.5  The Current Biodiversity Crisis

The alarming rate of biodiversity losses across the planet is becoming well known. In continen­ tal ecosystems, the rate of species loss might be approaching that of the K–Pg extinction, when the event that occurred 66 mya is normalized for its duration. Another way of expressing this crisis is that about 50% of all plant and animal species would be extinct by about 2100 if extinction rates during the late 1990s continued unabated (Wilson 2002). Of the five major extinctions affecting life during the past 541 million years, two are discussed in this review that are relevant for fossil insect biodiversity – the P–Tr and K–Pg events. Each of these events affected insect diver­ sity in different but nevertheless devastating ways. When both extinction events are collec­ tively considered, global taxic, taxonomic, eco­ logical, morphological, functional, and behavioral diversities were major casualties. Although data on rates of extant insect extinction are sketchy, the pervasiveness of the global event is n ­ onetheless

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evident, and its consequences for insect diversity should assiduously be avoided.

24.5 ­Conclusions From this review of the nature of and fossil record of insect biodiversity, five major conclusions are provided. These conclusions summarize the major points of this review but also indicate departure points for further exploration of past insect biodiversity. 24.5.1  The Importance of the Insect Fossil Record for Understanding Insect Diversity

The fossil record of insects is the only direct evi­ dence for understanding the past history of insect biodiversity and offers an unparalleled, albeit under‐studied, way of teasing apart the fundamental types of insect diversity. 24.5.2  The Five Fundamental Types of Diversity in the Insect Fossil Record

Taxonomic and taxic diversity are methods by which the variety of insects are enumerated in the taxonomic hierarchy, including species, gen­ era, families, and orders that are measurable at a site, in a habitat, or across time. Taxic and taxo­ nomic diversity are considered methodologically separate types of species or lineage diversity. Ecological diversity is comprised of the differ­ ences among insects that describe their trophic, community‐level, and environmental inter‐rela­ tionships at scales ranging from populations through habitats to biomes. Morphological diversity principally includes structural disparity, which is the range of anatomical, architectural, developmental, molecular, or other phenetically based form of a particular group or evolutionary lineage of insects. Functional diversity is the range of particular operations that insects accomplish or roles that they serve in communi­ ties or ecosystems. Behavioral diversity consists of those elements of biodiversity that condition how insect individuals or social groups have unit cohesion, respond to external stimuli such as predators, or comport within various interac­

tions. These five aspects of insect biodiversity have a significant fossil record that extends into deep geological time. 24.5.3  The Effect of Long‐Term Environmental Change on Insect Diversity

Long‐term environmental change has a global effect on insect diversity that involves major, protracted modification of the physical envi­ ronment such as climate change, significant alteration of the landscape, or physiochemical changes of water bodies. Major, prolonged bio­ logical events, such as the MLR and initial angi­ osperm diversification, also have an effect, and probably are more important. However, distinc­ tions between long‐term physical and biological effects often become confounded because cau­ sality cannot be pinned down in many cases. 24.5.4  The Effect of Short‐Term Environmental Changes on Insect Diversity

Global biodiversity is affected in deep time by brief, catastrophic events that dramatically change the future course of life. Extinction events are the most notable of these short‐term environ­ mental changes. As an example, major, short‐ lived, transient spikes of exceptional global warming and shifts in atmospheric CO2 concen­ trations have a signal effect on organisms through their responses, such as dramatic increases in insect herbivory. 24.5.5  How Fossil Insect Biodiversity Affects Us All

The fossil record of insect biodiversity contains aspects that engage human comprehension of the world and are instructive toward mitigating the current situation that faces human society.

Acknowledgments Finnegan Marsh expertly produced the figures. This is contribution 302 of the Evolution of Terrestrial Ecosystems consortium at the National Museum of Natural History.

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Wilf, P., C. C. Labandeira, K. R. Johnson, P. D. Coley and A. D. Cutter. 2001. Insect herbivory, plant defense, and early Cenozoic climate change. Proceedings of the National Academy of Sciences USA 98: 6221–6226. Wilf, P., C. C. Labandeira, K. R. Johnson and N. R. Cúneo. 2005. Richness of plant–insect associations in Eocene Patagonia: a legacy for South American biodiversity. Proceedings of the National Academy of Sciences USA 102: 8944–8948. Williams, J. W. and S. T. Jackson 2007. Novel climates, no‐analog communities, and ecological surprises. Frontiers of Ecology and Environment 5: 475–482. Wilson, E. O. 2000. The Diversity of Life. Second edition. Harvard University Press, Cambridge, Massachusetts. Wilson, E.O. 2002. The Future of Life. Vintage, New York, New York. 256 pp. Wilson, H. M. and W. A. Shear. 1999. Microdecemplicida, a new order of minute arthropleurideans (Arthropoda: Myriapoda from the Devonian of New York State, USA. Transactions of the Royal Society of Edinburgh 90: 351–375. Wing, S. L., F. Herrera, C. A. Jaramillo, C. Gómez‐Navarro, P. Wilf and C. C. Labandeira. 2009. Late Paleocene fossils from the Cerrejón Formation, Colombia, are the earliest record of Neotropical rainforest. Proceedings of the National Academy of Sciences USA 106: 18627–18632. Winkler, I. S., C. C. Labandeira, T. Wappler and P. Wilf. 2010. Distinguishing Agromyzidae (Diptera) leaf mines in the fossil record: new taxa from the Paleogene of North America and Germany and their evolutionary implications. Journal of Paleontology 84: 935–954. Wittry, J. 2006. Mazon Creek Fossil Flora. Earth Science Club of Northern Illinois, Chicago, Illinois. 164 pp. Wong, W. O., D. L. Dilcher, C. C. Labandeira, G. Sun and A. Fleischmann. 2015. Early Cretaceous Archaeamphora is not a carnivorous angiosperm. Frontiers in Plant Science 6: 326.

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the Entomological Society of America 93: 1230–1235. Zherikhin, V. V. and N. D. Sinitshenkova. 2002. Ecological history of the aquatic insects: Cainozoic. Pp. 417–426. In A. P. Rasnitsyn and D. L. J. Quicke (eds). History of Insects. Kluwer, Dordrecht, Netherlands.

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25 Phenotypes in Insect Biodiversity Research István Mikó and Andrew R. Deans Department of Entomology, Pennsylvania State University, State College, Pennsylvania, USA

Phenotypes – here defined as the micro‐ or macroscopically observable characteristics of organisms, including morphology, behavior, and physiological products (e.g., nests and galls) – i.e., the results of gene expression, in an environment, are the primary data for research on the evolution and taxonomy of insects (Deans et al. 2012a). The shapes, textures, colors, or simply the presence or absence of anatomical features or behaviors help us determine taxa and guide our hypotheses regarding their evolutionary history. Phenotype data are relevant to a broad range of most other research domains on insects, such as biomechanics (Labonte and Federle 2015), bioinspired engineering (Lau et al. 2014, Werfel et al. 2014, Winegard et  al. 2014), genotype–phenotype associations (Houle and Fierst 2013), genetic modification (Wang and Jacobs‐Lorena 2013), human‐disease research using insect models (Washington et al. 2009), and insect–plant interactions (Whitney and Federle 2013). Despite their broad relevance, phenotype data generally lack the standards necessary for computation or even broad understanding by humans. That is, one cannot simply query across phenotype data (e.g., “what are the most frequently described color phenotypes for Hymenoptera?”) with the aid of computers, as one can do with molecular data (e.g., “what are the most ­frequently sequenced protein‐encoding

genes for Hymenoptera?”). Researchers increasingly recognize that lifting this barrier will result in more meaningful, integrative research across the life sciences (Deans et al. 2015) as computational mechanisms are increasingly implemented in biodiversity research. This chapter provides a state‐of‐the‐art report on ontology based efforts to generate more accessible phenotype data and some insights into challenges and barriers to effective ­implementation of semantic approaches in phenotype representation.

25.1 ­Phenotype Data: Past and Present Phenotypes have long been the primary data for classifying biodiversity. Even a casual reading of Aristotle’s History of Animals, arguably one of the first hierarchical, scientific classifications of life, reveals rich descriptions of arthropod phenotypes. The following description of a decapod, for example, reads almost like a modern morphological treatment (Barnes 1984, Book IV: 526a1–526a12): In the crayfish the male differs from the female: in the female the first foot is bifurcate, in the male it is undivided; the belly‐fins in the female are large and overlapping on the neck, while in the male they are smaller and

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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do not overlap; and, further, on the last feet of the male there are spur‐like projections, large and sharp, which in the female are small and smooth. Both male and female have two antennae in front of the eyes, large and rough, and other antennae underneath, small and smooth. The eyes of all these creatures are hard, and can move either to the inner or to the outer side. The eyes of most crabs can do the same, to an even greater degree. Biodiversity researchers continue to publish prosaic descriptions like this, including hypotheses of function and evolutionary relevance, but most taxonomic descriptions are written in a quasi‐standardized, telegraphic style (Mayr and Ashlock 1991). Consider this descriptive statement of a bee species, also from Aristotle (Barnes 1984, Book IX: 627b23‐971): “another kind is what is called the robber‐bee, black and flat‐bellied”. In a contemporary taxonomic ­context, these characters would be written as follows: Body black. Metasoma flattened.

Although this simplified prose is arguably more concise and accessible than full sentences, phenotype annotations (data) continue to be generated using personalized lexicons, often impossible to reference or reconstruct, and without a truly standard syntax. Synonymy and homonymy are rampant (Yoder et al. 2010, Seltmann et al. 2013), and the interpretation of descriptions requires significant human brain power. One could express the above phenotype – that the body is black and the metasoma is flattened – in myriad ways (not even considering multiple languages): Body black, abdomen flattened. Blackish, with dorsoventrally compressed metasoma. Fully melanized; posterior tagma flat. At this point in time, only a human would consistently recognize these as roughly equivalent statements. And only a human reader could look past the lack of precision or outright inaccurateness of these descriptions. The entire body, for example, is rarely uniform in color. The parasitoid wasp in Figure 25.1 was described

Figure 25.1  Bright‐field micrograph of Gryonoides glabriceps, originally described by Dodd (1920) as being, in part: “Black; thorax and base of abdomen slightly suffused reddish; leg wholly yellow; antenna1 scape yellow.” Image by I. Mikó.

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by Dodd (1920) as being “black” (a few sclerites excepted, including the legs and scape). Yet the cuticle of the wing blade and the compound eye, each evidently (logically) part of the body, are transparent. Other parts of the body – the ocelli, processes, articulations, or setae – likewise are not black, though a reader could assume that they are, based on this description. Despite the imprecision of these annotations, most scientists could read them and understand that the body, and most likely the external cuticle alone, are “mostly black.” The broader scientific enterprise, however, depends on proper contextualization, especially when analysis relies on computers. For phenotype data generated by taxonomists to be repurposed outside of taxonomy (i.e., beyond a small community of highly trained domain experts), those data must be generated and made available in a more logically accessible and contextualized (meaningful) form.

25.2 ­Phenotype Data: Present and Future Taxonomists are not the only scientists who generate phenotype description in natural language. Despite some calls for a shift toward representing phenotypes as images, rather than text (MacLeod et al. 2010, Riedel et al. 2013), the practice is pervasive throughout the life sciences. Communities that work on model species recognized early on that computable phenotypes would increase the potential for novel discoveries (e.g., Washington et  al. 2009), and they have developed several approaches to make phenotype data more accessible. Most of these systems are built around ontologies, which increasingly facilitate biodiversity research (Deans et al. 2015). 25.2.1  Biological Ontologies

An ontology, at least in this context, is a formal representation of concepts, usually referred to as classes, in a particular domain, and of the relationships between those concepts. One can think

of an ontology as “a set of well‐defined terms with well‐defined relationships” (Ashburner et  al. 2000). Consider this entry from a well‐known informal ontology, The Torre‐Bueno Glossary of Entomology (Nichols 1989, p. 654; edited here for simplicity): scape, scapus; shaft (R. W. Brown); the first or basal segment of the antenna The definition (the first or basal segment of the antenna) is a class in the domain of entomology, which most researchers refer to by the terms “scape” or “scapus” (except R. W. Brown, who used the term “shaft”). The eight‐word definition also contains information regarding the relationships of this class to other entomological classes. One can understand that this class is a type of, or subclass of, “segment”, and it has the property of being part of the “antenna”. After reading the definition of segment (“subdivision of…an appendage…associated with muscle attachments”; Nichols 1989, p. 667) the reader can infer that because the scape is a type of segment, it must be associated with muscle attachments. Because of its natural language syntax (i.e., way it is written), the substantial knowledge represented in The Torre‐Bueno Glossary of Entomology (Nichols 1989) is only applicable, at least in the intended way, by human actors. A computer could not pick up a paper copy of the book, flip to the definitions of scape and segment, and logically conclude that scapes must have muscle attachments. Classes and their relationships must be formalized – written in a form that computers can process – to facilitate machine‐based computation across this kind of data. Only a handful of formal ontologies exist for insect biodiversity research. The Hymenoptera Anatomy Ontology (HAO) (Yoder et al. 2010) is the most well‐developed multispecies arthropod anatomy. Several others exist in other domains relevant to phenotypes, specimens, and so forth (Table 25.1). Formal ontologies can be developed using a number of approaches, but the most widely used standard in biology is Web

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Table 25.1  Some ontologies relevant to insect biodiversity research; note that several other ontologies are available or required for proper formalization of insect phenotype data. Ontology

Description

Reference

Hymenoptera Anatomy Ontology (HAO)

Used for descriptive taxonomy and comparative morphology of Hymenoptera

Yoder et al. (2010)

PATO

Ontology of phenotype descriptors, used in multiple research domains

Mungall et al. (2010)

Biospatial Ontology (BSPO)

Biospatial terms, such as dorsal and ventral

Mungall (2015)

Environment Ontology (ENVO)

Includes classes relevant to environment and habitat

Buttigieg et al. (2013)

Biological Collections Ontology

Includes classes relevant to natural history collections

Walls et al. (2014)

Gene Ontology (GO)

Covers genes, their products, and processes

Ashburner et al. (2000)

Ontology Language (OWL; World Wide Web Consortium 2015). An OWL representation of an ontology allows one to compute across phenotype data sets to address scientific questions and infer relationships that are not explicitly stated (Fig. 25.2; Balhoff et al. 2013). 25.2.2  Ontologies in Biodiversity Research

Insect biodiversity scientists have yet to integrate large‐scale, machine‐based computation into their research on phenotypes, but the potential for discovery is substantial. Taxonomic and other morphological descriptions are unparalleled sources of information about evolutionary novelties and adaptations (Deans et al. 2012a, 2012b), which remain hidden from most researchers. The following use‐cases, some of which are simple to implement, serve as examples where data generation and the process of discovery could benefit from an ontological approach. 25.2.2.1  Referencing a Glossary

Although formal ontologies are primarily designed for machine‐based computation, the process of developing these ontologies and the development of user‐friendly tools can facilitate human‐based computation as well. In a recent analysis of 428 papers on arthropod systematics, Deans et al. (2012a) found that fewer than 3% of authors included a glossary of morphology terms used in character descriptions. Almost one‐third

of the papers did not cite any references for term meanings. This situation is problematic, especially when terms frequently refer to more than one concept (e.g., “paramere” can mean at least five different body parts (Yoder et al. 2010), and the “forearm” of Lepidoptera is quite different from the forearms of Mammalia). With community‐selected “preferred terms” for classes, an ontology could function as a controlled vocabulary, which unifies within‐ and cross‐domain knowledge (Noy and McGuiness 2001, Smith et  al. 2007) and improves small‐ and large‐scale (interdomain) communication. Even without preferred terms, an ontology can play a central role in word‐sense disambiguation of anatomical terminology, for example as an online glossary. An online description, marked up with Web links (URLs) that reference explicit concepts can provide more information that cannot be stored in the ontology itself (e.g., images). There are tools (Seltmann et al. 2012) that match natural language descriptions to concepts in the HAO, so that a table of terms, definitions, and URIs (in this case, links to Web pages with more information) can be generated. In our experience, this resource is already useful for insect lineages outside of the Hymenoptera, because many homologous structures are shared widely across hexapods (e.g., Mikó et al. 2014). In this sense, the HAO forms a semantic core from which anatomy ontologies for related groups can be developed.

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Figure 25.2  Hymenoptera Anatomy Ontology (HAO). Partial graph of the HAO, illustrating the explicitly stated properties of the scape. The curators of this ontology explicitly relate (solid arrows) scape to antennal segment, using the “is a” relationship. There is no explicitly stated “part of” relationship for the scape, but one can infer, through properties of anatomical entities higher in the hierarchy, that the scape is part of the antenna (dashed arrow). The term/class “Scape” itself is expanded to reveal its URI (resolvable identifier, at top), instances (annotated images), labels used in the literature (bold name is preferred term), and genus differentia definition. Image by A. R. Deans.

25.2.2.2  Generating Logically Consistent Phenotypes

Appending a glossary of logical, structured morphology definitions to manuscripts in preparation increases the accessibility of those data to future users. Adding structured phenotype descriptions is the next step in making data available for computation. This practice also improves the character development by forcing the researcher to think carefully about the phenotypes being described. Is the character circumscription overly complex? Are the states consistently formulated and diagnostically relevant?

Consider the color of the scape of the wasp in Figure 25.1. One could create a character called “scape color”, with the following states: (0) scape brown and (1) scape black. Brown and black are both children of “color” in the Phenotypic Quality Ontology (PATO; Mungall et al. 2010), and, therefore, these states are perhaps more likely to be diagnostically coherent. Attempts to add character states not referring to color (pigmentation), for example (2) scape hairy and (3) scape rugose, would create an inconsistency in state identity, with downstream challenges for computational processing (i.e., stand out as inappropriate). Hairy

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is a child of pilosity, and rugose is a child of texture; the states are unlikely to be semantically coherent. The example is a bit silly, perhaps, but problematic character states are pervasive in insect systematics. This character state partitioning from Whiting et al. (1997), edited for clarity, is a bit more subtle: Ovipositor (0) absent, (1) present, (2) vestigial, (3) modified (reduction in the second valvulae, the third valvulae serving as the functional components of the ovipositor), (4) fused. The character combines phenotypes that are related to count (presence, absence), structure (fused, vestigial), and detailed morphology (modified). Are these states in precise semantic correspondence, or should the phenotypes be scored as separate characters? Another character from this data set suffers from similar problems: trochantin (0) absent, (1) present, (2) trochantin‐episternal sulcus present. To a naive reader, this character appears to refer to counts of two different anatomical entities (i.e., the states are not in ontological correspondence), the trochantin itself and a sulcus that is associated with the trochantin. We are not suggesting that these phenotypes are not phylogenetically relevant; however, the way in which they were individuated might hinder both human and computational processing.

In our own research, we found many examples where character states were too loosely described to be meaningful, and a semantic approach improved their description. Mullins et  al. (2012) and Balhoff et  al. (2013) provide examples of biodiversity research that incorporates explicit, logical representations of phenotypes, composed in OWL, using many of the ontologies in Table 25.1. Mikó et al. (2015) provide details on how to generate data in this format (Table 25.2). 25.2.2.3  Reasoning Across Phenotype Data

When phenotype data are generated logically, using multiple ontologies and standards of the  Semantic Web, one can leverage the logic inherent in those ontologies for computation. ­ Numerous real and hopeful examples of computation have been published, illustrating a range of possible use‐cases, including image management (Ramírez et al. 2007), connecting human‐disease phenotypes to mutant model organisms (Washington et al. 2009), species determination through successive queries (Deans et al. 2012b), enhancing taxonomic practice (Balhoff et  al., 2013), tracing structural complexity along branches of a tree (Ramírez and Michalik 2014),

Table 25.2  Typical character statements and their semantic representations. Character

Semantic annotation

Antennal shelf count: present

has part some antennal shelf

Anterolateral mesopectal projection 2‐d shape: square shape

has part some (anterolateral mesopectal projection and (has part some (lateral side and (bearer of some square))))

Body length universal: 4.6–8.0 mm

has part some (body and (has part some (median anatomical line and (bearer of some (length and (is quality measured as some ((has measurement unit label value millimeter) and (has measurement value some (float[≥4.6f ] and float[≤8.0f ])))))))))

Male petiole length versus width: 2.5× as long as wide

has part some (petiole length and (bearer of some (length and (is quality measured as some ((has measurement unit label some (width and (inheres in some abdominal segment 2))) and (has measurement value 2.5))))))

Mandibular teeth count: 4

has part some (mandible and (has component exactly 4 tooth))

Ventrolateral region of has part some (mesosoma and (has part some (ventrolateral region and (bearer of mesosoma texture: areolate some areolate))))

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ontology‐based partitioning for phylogenetic analysis (Tasarov and Génier 2015), linking morphometric data to descriptions (Csösz et al. 2015), correlating phenotypes with environment (Thessen et al. 2015), a similarity‐based approach for recognizing comparative homologs (Vogt 2015), and correlating phenotypes with genetics and environment (Deans et al. 2015). Prospecting for non‐model species is another process that could be facilitated by a semantic phenotype approach. Genomic and developmental approaches are becoming increasingly important, with an extraordinary amount of research done on insect models, such as Drosophila and Tribolium species. Each of these systems, coincidentally, has its own phenotype and anatomy ontologies (Dönitz et  al. 2013, Osumi‐Sutherland et al. 2013), but are they the most well‐suited model species for research on certain phenotypes? Could vision impairment in humans be better understood by studying lineages with natural variation in eye development? If one could query across known insect phenotypes – for example, “show me all insects that exhibit eye reduction”  –  other models might reveal themselves and perhaps facilitate compelling comparisons. Research on other developmental processes (e.g., branching morphogenesis or epithelial folding) likewise could benefit from increased accessibility to phenotype data from the natural world (Deans et al. 2015). Workflows are now available for generating computable phenotype annotations, using either manual or automated approaches. Human‐mediated annotations, using applications such as Phenex (Balhoff et al. 2010) or Protégé (http://­ protege.stanford.edu/), yield relatively precise and accurate data, but the approach is not scalable to  the vast corpus of legacy phenotype treatments. Formulating semantic statements currently requires some knowledge about OWL ontologies and Manchester syntax (Table 25.2; http://www. w3.org/TR/owl2‐manchester‐syntax/), and probably requires a more streamlined and user‐friendly workflow to be adopted more broadly. Machine‐ mediated annotation tools, such as CharaParser

(Cui 2012, Cui et al. 2016), are less time c­ onsuming, but, due to the high number of homonyms in biodiversity descriptions, these methods have a higher error rate.

25.3 ­Challenges and Future Directions Insect biodiversity research already uses ontologies (e.g., our hierarchical classification of life), yet there remains some nescience or even skepticism regarding their application. Below we outline some misconceptions and real barriers  to effective implementation of semantic approaches to phenotype representation. We also offer ideas for how to surmount these barriers, where possible, and describe a vision for the future of phenotype data in biodiversity research. 25.3.1  Social Challenges to “Standardization”

A common misconception is that the ontology implementation is an attempt to force a research community to use a unified lexicon. Biodiversity research is extraordinarily heterogeneous, and each domain has its own history of language and “camps” of experts. Sociologically, it is impossible to get widespread agreement on terminology. An ontology can act as a controlled vocabulary, and best practices call for a single “preferred” term for each concept. The sensu model and glossary development described above, however, allow for the retention of personalized lexicons, while also facilitating communication through a standard set of concepts. 25.3.2  Ontology Development Barriers

Several taxon‐focused phenotype ontology development projects have been funded by the United States National Science Foundation, each employing a large staff of informaticians, postdoctoral researchers, and other high‐level personnel. Given this history, there is a perception that developing a new ontology, for hemipteran anatomy, for example, is a substantial challenge. Much of the heavy

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lifting  –  development of concepts, standards, tools, and documentation – has been done, however, which lowers barriers to future implementation. The Hymenoptera Anatomy Ontology, for example, could be cloned and modified with relative ease and applied to almost any hexapod taxon. This approach is being done for Coleoptera and  Neuroptera, with relatively few resources. Documentation and examples of implementation are forthcoming. 25.3.3  Ontology Implementation Barriers

Researchers who generate phenotype data, especially taxonomists, already document their science extensively, perhaps more so than any other domain in the life sciences (e.g., voucher‐specimen management). Adding yet another task, the generation of structured phenotype data, might further prolong the descriptive process at a time when biodiversity is rapidly disappearing. It is true that current workflows are somewhat complex, requiring multiple applications, and they use unfamiliar standards and syntax (Mikó et al. 2015). There have been several calls for investment in phenotype‐data collection and analysis (Deans et  al. 2015), however, and tool development is ongoing. The Gene Ontology project (www.geneontology.org), for example, has implemented a templates‐based approach (Dietze et al. 2014) that facilitates ontology implementation without technical knowledge of syntax and the ontologies themselves. We anticipate a relatively seamless, user‐friendly integration of these approaches into biodiversity research in the near future. 25.3.4  Phenotype Complexity

Phenotypes can, indeed, be elaborate, and many natural language descriptions cannot be understood without figure references or the specimens in hand. Characters with graded quantifiers  –  e.g., “Scutellum strongly curved in lateral view (fig. 6A)” versus “Scutellum weakly curved in lateral view (fig. 6B)” (Baur et al. 2014) – and terms referring to the putative evolutionary

­ istory, rather than pointing to an observable h anatomical structure  –  e.g., “parossiculi fused” (Mikó et  al. 2013)  –  rely on non‐textual references (i.e., images). The real meaning of these descriptions remains hidden behind the text (scutellum is more curved in species A than in species B, and an area of the integument is divided into two sclerites by a median conjunctiva) and cannot be translated adequately into semantic statements. Taxonomic descriptions are flooded with similar culprits that must receive only course‐grained annotations (“scutellum curved”) or be revised (“median conjunctiva present”). 25.3.5  Communicating Primarily with Semantic Phenotypes

The default approach to describing phenotypes is currently natural language, and most researchers assume that the subtleties and extent of many characteristics can adequately be described only in prose. The learning curve for telegraphic natural language, however, might be at least as steep as the learning curve for formal description. We understand the aversion of domain experts against structured representations, especially with its unfamiliar cadence and heavy usage of parentheses, but these representations provide a much more objective and standardized way to describe phenotypes. Our current approach is to complement, rather than replace, natural language descriptions with semantic phenotype data. 25.3.6  No Clearinghouse for Phenotype Data

At this time, there is no phenotypic analog to GenBank®, a well‐developed, high‐demand database with a suite of tools that facilitate genetic research (Benson et al. 2005). Phenotype data for most insects currently reside in individual publications, distributed across more than 1000 scientific journals (Deans et al. 2012b) or as files in different data repositories (e.g., the Dryad Digital Repository [www.datadryad.org]; Balhoff et  al. 2013, Mikó et al. 2014). Researchers with an interest in a particular taxon must seek out these data to perform

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any analyses. For broader computation, one must rely on other methods (text mining), and this remains an opportunity for innovation. 25.3.7  Reasoning Challenges

Balhoff et al. (2013) demonstrate that for small data sets one can make inferences computationally, using existing semantic reasoners and in a short period of time, and this approach has utility for error checking and small‐scale discovery. Larger data sets, however, require substantial computing resources. There is a substantial community effort to improve reasoning algorithms, through grand challenges and other mechanisms, and we anticipate that useful, large‐scale computation is on the horizon. None of these barriers are insurmountable. Given the increasing interest in semantic approaches to phenotype representation, we predict that this aspect of the taxonomic enterprise  –  describing the diversity of life  –  will remain as vibrant and relevant as ever.

Acknowledgments This material is based on work supported by the US National Science Foundation, under grant numbers DBI‐0850223, DEB‐0956049, DBI‐1356381, and DEB‐1353252. Any opinions, findings, and conclusions or recommendations expressed in this material are those of the authors and do not necessarily reflect the views of the National Science Foundation. James P. Balhoff (RTI International) provided invaluable comments on an earlier version of this manuscript.

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the Hymenoptera Anatomy Ontology: utility, clarification, and future directions. Journal of Hymenoptera Research 27: 67–88. Seltmann, K. C., Z. Pénzes, M. J. Yoder, M. A. Bertone and A. R. Deans. 2013. Utilizing descriptive statements from the Biodiversity Heritage Library to expand the Hymenoptera Anatomy Ontology. PLoS ONE 8: e55674. Smith, B., M. Ashburner, C. Rosse, J. Bard, W. Bug, W. Ceusters, L. J. Goldberg, K. Eilbeck, A. Ireland, C. J. Mungall, N. Leontis, P. Rocca‐ Serra, A. Ruttenberg, S.‐A. Sansone, R. H. Scheuermann, N. Shah, P. L. Whetzel and S. Lewis. 2007. The OBO Foundry: coordinated evolution of ontologies to support biomedical data integration. Nature Biotechnology 25: 1251–1255. Tarasov, S. and F. Génier. 2015. Innovative Bayesian and parsimony phylogeny of dung beetles (Coleoptera, Scarabaeidae, Scarabaeinae) enhanced by ontology‐based partitioning of morphological characters. PLoS ONE 10: e0116671. Thessen, A. E., D. E. Bunker, P. L. Buttigieg, L. D. Cooper, W. M. Dahdul, S. Domisch, N. M. Franz, P. Jaiswal, C. J. Lawrence‐Dill, P. E. Midford, C. J. Mungall, M. J. Ramírez, C. D. Specht, L. Vogt, R. A. Vos, R. L. Walls, J. W. White, G. Zhang, A. R. Deans, E. Huala, S. E. Lewis and P. M. Mabee. 2015. Semantic linking of phenotypes and environments: a review. PeerJ 3: e1470. Vogt, L. 2015. Assessing similarity: a semantic approach to non‐evolutionary comparative homology. PeerJ PrePrints 3: e1523. Walls, R. L., J. Deck, R. Guralnick, S. Baskauf, R. Beaman, S. Blum, S. Bowers, P. L. Buttigieg, N. Davies, D. Endresen, M. A. Gandolfo, R. Hanner, A. Janning, L. Krishtalka, A. Matsunaga, P. Midford, N. Morrison, E. O. Tuama, M. Schildhauer, B. Smith, B. J. Stucky, A. Thomer, J. Wieczorek, J. Whitacre and J. Wooley. 2014. Semantics in support of biodiversity knowledge discovery: an introduction to the biological collections ontology and related ontologies. PLoS ONE 9: e89606.

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26 Global Change and Insect Biodiversity in Agroecosystems David R. Gillespie1, Matthew J. W. Cock 2, Thibaud Decaëns 3, Philippa J. Gerard 4, Sandra D. Gillespie 5, Juan J. Jiménez 6 and Owen O. Olfert 7 1

Agassiz Research Centre, Agriculture and Agri‐Food Canada, Agassiz, British Columbia, Canada Centre for Agriculture and Biosciences International, Egham, UK 3 Centre d’Ecologie Fonctionnelle et Evolutive, Montpellier Cedex 5, France 4 Biocontrol & Biosecurity, AgResearch Ltd., Ruakura Research Centre, Hamilton, New Zealand 5 Biology Department, University of the Fraser Valley, Abbotsford, British Columbia, Canada 6 Instituto Pirenaico de Ecología (IPE), Consejo Superior de Investigaciones Científicas (CSIC), Jaca, Spain 7 Agriculture and Agri‐Food Canada, Saskatoon, Saskatchewan, Canada 2

In this chapter we consider the possible effects of global change on the biodiversity of insects in agroecosystems and the implications of these effects for human society. We focus mostly on plant‐based agricultural systems. In the context of human societies, negative effects on the bio­ diversity of insects in and around agricultural systems have direct consequences for food and fiber supplies, and food security for humans. Insects are integral to the functioning of agroecosystems (Cock et al. 2012). Herbivorous insects cause crop losses (Oerke 2006), and insects that are predators and parasitoids are integral to ecosystem‐based regulation of pest populations (Costanza et al. 1997, Losey and Vaughan 2006). Insects pollinate agricul­ tural crops. Although honeybees are the main pol­linators in many crops, other insect taxa can be  important pollinators in agroecosys­ tems  (Nicholls and Altieri 2013). Insects are intimately involved in leaf‐litter degradation, manure recycling, and soil function (Cock et al. 2012), and they are important food sources for  many vertebrates and invertebrates in and

around agroecosystems (Hallmann et al. 2014). Changes in the biodiversity of insects in agroe­ cosystems have effects on both the human food supply and agroecosystem health.

26.1 ­Global Change Global climate change is one of the domi­ nant issues affecting human societies (IPCC 2013, 2014). It is a complex and multifaceted phe­nomenon. The accumulation of anthropo­ genic greenhouse gases, such as carbon dioxide, nitrous oxide, and methane, is responsible for an increasing fraction of the Sun’s heat energy being retained by the planet, thus driving a steady increase in global average temperature. The mean global surface temperature is pro­ jected to increase throughout the 21st century and to be accompanied by more frequent warm days, less frequent cold days, and increases in the frequency and duration of extreme weather events (IPCC 2013). Important regional dif­ ferences in how climate change is projected to

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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progress are especially relevant to a discussion of the effects of climate change on the biodiver­ sity of insects. A recent report on the physical basis  of climate change from the IPCC (2013) highlights the following regional trends. First, greater increases in seasonal mean and annual mean temperatures are expected in the trop­ ics  and subtropics than in the mid‐latitudes. Second, the Arctic will warm to a greater degree than the rest of the globe. Third, the amount of warming will be greater over land areas than over oceans. Fourth, mid‐latitude and subtropi­ cal dry regions probably will see a decrease in precipitation, whereas mid‐latitude wet regions will see an increase in precipitation. And fifth, extreme precipitation events over mid‐latitude and wet subtropical regions will become more intense and more frequent. Changes in sea‐ice cover and permafrost extent, and changes in the duration and extent of important regional weather events, such as the monsoon seasons, are other important and relevant trends (IPCC 2013). We suggest that the combined effects of changes in seasonal and annual heat and precipitation patterns proba­ bly will have the greatest effects on insect bio­ diversity. Although critically important, other climate change trends, such as decreasing ocean pH and increasing sea levels, are unlikely to affect insect biodiversity directly. Global climate change is only one part of the ongoing disruption of the abiotic and biotic sys­ tems of planetary homeostasis by human activi­ ties, which collectively constitute global change (Tylianakis et al. 2008, Barnosky et al. 2012). The human population is increasing; as of 2015, the world population exceeded 7 billion and is projected to exceed 9 billion by 2050 (United Nations 2014). Through sheer numbers alone, humanity is consuming an increasing fraction of  the planet’s land resources, and the largest part of this is for food and fiber production (Pimm 2001). Industrialization is the second major driver of global change. For example, the global per capita gross domestic product in US$ (Fig. 26.1a) and the global per capita energy consumption in kg of oil equivalents (Fig. 26.1b)

demonstrate the increase in industrialization on a per capita basis. Humans are producing more goods, and are consuming more energy to do so. Land‐use extent and intensity are also changing. An increasing amount of the world’s land is devoted to human habitation and food produc­ tion. In Australia, China, Europe, New Zealand, and North America, increases in agricultural production depend on the use of increasing amounts of pesticides and fertilizers and increases in the ratio of irrigated to non‐­ irrigated land. In South America, Sub‐Saharan Africa, and parts of Asia, increases in produc­ tion are being achieved by clearing new land (FAO 2009). Forestry and clearing for additional food production are depleting forests at an alarming rate. From 1990 to 2011, the world’s forested lands decreased by 1.16 million ha2 (World Bank Group 2016b), which is an area greater than the total land area of many coun­ tries, and equal to 0.8% of the world’s land area. Increases in trade and population are breaking down biogeoclimatic barriers to movement and dispersal and, together with land‐use changes, are resulting in an alarming increase in the accidental or deliberate introduction of invasive alien species to new habitats (Hobbs 2000, McNeely 2000). The responses of species affected by global climate change are limited to moving to new ranges and adapting to (or coping with) chang­ ing climatic conditions in situ (Davis et al. 2005). These responses, however, play out against the mosaic of global change impacts  –  urbaniza­ tion, increasingly intensive agriculture, destruc­ tion of key habitats, and interactions with invasive species, to name a few. The equatorial range limit of a herbivore impacted by climate change might, for example, shift poleward in response to increased frequency of heatwaves, but if necessary host plants are absent from that  potential new range because of habitat destruction or differential dispersal or tempera­ ture tolerances, there might be impacts on the abundance and distribution of the herbivore (Davis et al. 1998, Warren et al. 2001, Pelini et al. 2009). In response to shifts in flower phenology,

26  Global Change and Insect Biodiversity in Agroecosystems

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Figure 26.1  Two metrics that illustrate the global increase in industrial activity. Both per capita economic activity (a) and per capita energy consumption (b) increase over time. These are per capita measures; thus, the increases are not due to increases in numbers of humans, but to increases in industrial activity of all sorts. (Original by authors; data sources: World Bank Group 2016a, 2016c.)

individuals of a specialist pollinator might, for example, feed from a broader range of flowers, but that could bring them into competition with invasive pollinators and expose them to invasive diseases. At regional scales, the effects of cli­ mate shifts on the biodiversity of insects are occurring on a landscape that is itself rapidly changing, and many of these landscape changes have severe implications for biodiversity.

26.2 ­Insect Biodiversity in Agriculture Agricultural activities have greatly changed our  planet, with 12% of the ice‐free land sur­ face  under cropland and 22% under pasture (Ramankutty et al. 2008). Pastoral lands that are managed for grasses and forbs consumed by grazing animals make up an important and

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substantial fraction of agricultural land use. The sheer area of habitat that is now agriculturally influenced means that conserving biodiversity on a global scale requires conserving biodiver­ sity in agroecosystems (Matson et al. 1997, Wray et al. 2014). Overall, there are roughly 7000 plant species grown for food, with rice, corn (maize), wheat, and potatoes being the dominant crops (FAO 2015). This extraordinary plant biodiversity supports a rich community of herbivorous insects. The herbivore species are predominantly in the insect orders Coleoptera (beetles), Diptera (flies), Hemiptera (e.g., aphids, mealybugs, planthoppers, psyllids, scales, and true bugs), Hymenoptera (sawflies), Lepidop­ tera (moths), Orthoptera (grasshoppers and locusts),  and Thysanoptera (thrips). In grass­ lands, Coleop­tera, Lepidoptera, and Orthoptera are common, especially in dry subtropical and temperate regions. Important insect natural enemies of these insect pests are in the Coleoptera (predatory families), Diptera (pred­ atory and parasitoid families), Hemiptera (predatory true bugs), and Hymenoptera (para­ sitoid wasps). Soil communities are dominated by predators and herbivores in the orders Coleoptera and Diptera and by members of  these two orders that are important in decom­ position and nutrient recycling (e.g., Scarabaeidae and Stratiomyidae). Although the domesticated honeybee Apis mellifera L. domi­ nates in the commercial pollination of crops, a multitude of other species play a role in pol­ lination, predominantly species in the Dip­tera (particularly flower flies, Syrphidae) and Hymenoptera (mostly bees, Apoidea). 26.2.1  What Do We Mean By “Biodiversity”?

At its simplest, the diversity of species in a region is the total number of species at all sites or habitats within the region (richness). Other measures of diversity take into account both the  abundance and evenness of species in the  habitat (Magurran 2004). Both of these sorts of measures are relevant in agroecosys­ tems. Species richness and relative abundance

determine the identity and strength of pair­ wise species interactions that influence vari­ ous ecosystem functions and services. Core ecosystem functions, such as carbon fixation, nutrient cycling, water filtration, and productivity, are supported by various biogeo­ chemical processes (Hooper et al. 2005). These processes can also be regulated by organisms within the ecosystem itself, and where this occurs, the consensus among scientists is that the rates or directions are a direct function of biodiversity (Hooper et al. 2005). That portion of the species richness in a habitat that contrib­ utes to ecosystem processes is functional biodi­ versity. When human interests are promoted by an ecosystem process, the process is an ecosys­ tem service. Insects in agroecosystems are closely tied to ecosystem services through the promotion of plant reproduction by transfer of pollen from anther to stigma in angiosperms (pollination), through the conversion of leaf material to detri­ tus (herbivory), and through contribution to the regulation of populations in food webs (preda­ tion and parasitism). Moreover, species‐spe­ cific traits moderate these services. Functional trait biodiversity takes into account these spe­ cies‐specific traits that mediate the contribution of a species to a particular ecosystem function or service. For example, different species of nat­ ural enemies might attack different life stages of a herbivore, which distributes mortality and potentially increases the biocontrol service. Adding another species that attacks the same stage can increase biodiversity without increas­ ing functional trait biodiversity and, thus, not affecting the biocontrol service. Overall, func­ tional trait biodiversity is a better predictor of service than species counts (Frainer et al. 2014, Deraison et al. 2015). In agroecosystems, the diversity of insects is influenced by both the cropping system and proximity to unmanaged plant communities. The deliberate reduction of biodiversity is an operational assumption in high‐input monocul­ tures (Swift et al. 2004). These are consequently dominated by a few key pest species (Bengtsson

26  Global Change and Insect Biodiversity in Agroecosystems

et al. 2005, Batáry et al. 2012), and ecosystem services such as biological control and nutrient cycling tend to be replaced by petrochemical energy (Swift et al. 2004). At the other end of the spectrum, insect diversity tends to be compara­ tively high in low‐input polycultures, such as organic crops, rangelands, or pastoral grazing lands (Bengtsson et al. 2005, Batáry et al. 2012). Proximity to low‐input or unmanaged habitat tends to increase species richness of insects in intensively managed cropping systems (Blitzer et al. 2012, Batáry et al. 2013, Le Féon et al. 2013). In agroecosystems, diversity of insect species is positively correlated with ecosystem services such as pollination (Martins et al. 2015) and pest‐population regulation (Harrison et al. 2014). This functional biodiversity is important in maintaining the agroecosystem function and ecosystem services on which humans rely.

26.3 ­Effects of Global Change on Biodiversity – What Do We Know? Changes in the distribution and abundance of insect species driven by global change can lead to changes in community composition and diversity at local and regional scales, with impli­ cations for ecosystem processes. In this section, we review these patterns and their underlying causes in the context of several important eco­ logical processes in agroecosystems: predator– prey interactions, soil maintenance, and crop pollination. 26.3.1  Crop Pests and Natural Enemies 26.3.1.1 Distribution

One of the best‐documented consequences of global climate change is the ongoing poleward and altitudinal shift in distributions of many insect species (Mason et al. 2015). Estimated rates of poleward movements have ranged from 0.6 km per year (Parmesan and Yohe 2003) to 1.7 km per year (Chen et al. 2011), and a rapid 2.7 km per year for crop pests (Bebber et al. 2014). Changes in species distributions

so  far do not seem to be even across groups. Although most groups of insect pests, including Coleoptera, Diptera, Hemiptera, Lepi­doptera, and Thysanoptera, have consistently moved poleward in the Northern Hemisphere over the past 50 years, this pattern has not so far been evident in Blattodea (as Isoptera) or Hymenop­ tera (Bebber et al. 2014). Upslope movement has been estimated at 11 m per decade (Chen et al. 2011) to 50 m per decade (Whittaker and Tribe 1996, Menéndez et al. 2007, Colwell et al. 2008). Root et al. (2003) reported that 694 animal and plant species have already responded to regional climate changes that have occurred during the 20th century. Walther et al. (2002) suggested that invertebrate responses to a changing climate will be related more to regional temperature and precipitation shifts than to approximated global averages. As a consequence of this poleward and upslope movement of insects, the biodiversity of insects present at any given place on the globe is stead­ ily changing. Concurrent with poleward range expansion is a general trend toward earlier seasonal occur­ rence of insects. Fleming and Tatchell (1995) showed that five aphid species in Britain were dispersing from overwintering hosts earlier in the year, probably in response to climate warm­ ing, and models by Harrington et al. (2007) sug­ gest that this is probably a general trend in the Aphididae. Analysis of 50 years of aphid flight‐ monitoring data from the Rothamsted insect survey (Bell et al. 2014) showed that the date of first aphid flight has shifted earlier, at a rate of approximately −0.6 days per year. Seasonal phe­ nology determines, to a large degree, the effects of invasive species on competing resident spe­ cies, where early phenology allows the invasive or migrant to pre‐empt a resource (Gidoin et al. 2015). This is particularly true for invasive plants (Wolkovich et al. 2013), which has knock‐ on effects on the surrounding insect communi­ ties. The effects of warmer temperatures on plant communities and insect food webs, com­ bined with increased land‐use intensity, could have dramatic and negative effects on insect

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communities in agricultural settings (De Sassi et al. 2012). Jeffs and Lewis (2013) reviewed the effects of climate warming on host–parasitoid interac­ tions. Parasitoids are expected to respond to cli­ mate warming in ways similar to those of other insects, but the authors suggested that responses to climate change will be linked to the dispersal ability of host and parasitoid and to the host specificity of the parasitoid. The ability of insects to respond to changes in  climate will vary by species. In general, the potential distribution of insects is, and will continue to be, constrained by their response to extremes in temperature and precipitation, and  to an increase in the frequency of such extremes  (Hance et al. 2007). However, most insect species probably will have the ability to shift their geographical distributions in response to changes in climate so as to remain in geogra­ phical areas to which they are well adapted (Gillespie et al. 2013). The magnitude of predicted temperature change associated with changing climate is not within the historical experience of modern agri­ culture (Olfert et al. 2012). As a result, the use of  historical data as analogues to classify bio­ geographic zones that might be susceptible to establishment of invasive pest species or to identify regions that are best suited for intro­ duction of classical biological control agents is somewhat restricted (Gillespie et  al. 2013). Bioclimate simulation modeling (also known as ecological niche modeling) can assist in predict­ ing the potential impacts of changes in tempera­ ture and precipitation on invasive alien species and the related system vulnerability. Bioclimatic simulation models have been used successfully to predict the population distribution and establishment of insect pests and their natural enemies in new environments (Haye et al. 2013) and in response to a changing climate (Olfert et al. 2015). Simulation soft­ ware, such as CLIMEX, enables the develop­ ment of models that describe the potential distribution and seasonal population density of a species of interest based on its geographic

range, phenology, seasonal abundance, and empir­ical data (Hearne 2016). This modeling allows researchers to develop an overview of climatic factors (including climate change) that affect species distributions and abundance. In addition, the analysis allows the documentation of non‐climatic factors that might be limiting species distribution ranges. To date, the majority of bioclimate simulation modeling has focused on pest species, in part due to the increased complexities of tritrophic systems (host–pest species–natural enemies). Owing to species‐specific differences in con­ straints imposed by changing climates and responses to those constraints, bioclimate sim­ ulations should be interpreted with caution (Jeschke and Strayer 2008). These differences in responses might cause currently interdepend­ ent species to become uncoupled if their responses to the changing climate diverge (Parmesan 2007). The use of different general circulation mod­ els (GCMs) covering a range of projections to assess the potential range expansions of pest groups in response to changing climate is well documented (Olfert et al. 2011). As a result of the variability of climate projections between models, climate‐change impact studies benefit from using multiple GCMs (Olfert et al. 2012). A complementary approach to studying the effects of future climate on species distribution and abundance is an incremental sensitivity analysis (Olfert et al. 2011). This approach typi­ cally creates scenarios across a range of possible combinations of temperature and precipitation. Olfert and Weiss (2006) showed that an incre­ mental analysis provided additional insight by systematically assessing insect responses across ecoregions of interest. The ability of insect species to successfully establish in a new region will depend on a number of factors, including the number of consecutive years that suitable climate is availa­ ble (Venette and Hutchison 1999). As a result, populations on the periphery of their geo­ graphic ranges may experience fewer seasons of suitable climate than those near the center of

26  Global Change and Insect Biodiversity in Agroecosystems

their ranges. The analysis of incremental sce­ narios offers insights into insect responses within these peripheral geographic ranges. For example, Olfert et  al. (2015) showed that instances of predicted range expansion are most prevalent in northern regions of North America. Conversely, model output predicted that the range and relative abundance of the organisms under study could also contract as a result of heat stress in regions where climate condi­ tions  become limiting owing to warmer, drier climates. Native insects characteristically have stable tritrophic relationships that developed under specific, stable climate norms, and these include a suite of natural enemies. Higher trophic levels are more likely to be affected by climate change because they require the lower trophic levels to adapt to climate change (Gutierrez et al. 2008). Bioclimatic simulation of a North American pest grasshopper, Melanoplus sanguinipes (Fab­ ricius), predicted that it would increase in abundance, and its distribution would shift sig­­ ni­ficantly poleward but contract in the south (Olfert et  al. 2011). However, the impact of changing climate on biotic factors, such as dis­ eases and parasites of a native species (e.g., M. sanguinipes), must be considered. A number of major grasshopper outbreaks in western Canada were terminated by epizootics of the fungus Entomophaga grylli (Riegert 1968). Even though future climates might be conducive to grass­ hopper populations, they might also provide favorable conditions for natural enemies. The alternative can also occur, and the natural‐ enemy relationships can become unlinked. To accurately estimate pest‐population range and abundance, bioclimate modeling of insect pests requires a multitrophic approach (host plants– pest species–natural enemies). Migratory species are typically unable to per­ manently establish in ecoregions into which they have migrated, but flourish until unfavora­ ble conditions result in population decline. In northern latitudes, these migratory insect spe­ cies cannot survive the cold winters, and inva­ sions during spring and summer are the result

of northerly migrations on favorable wind pat­ terns (Dosdall et al. 2013). The diamondback moth Plutella xylostella (L.) (Plutellidae) is a migratory pest of vegetable and oilseed cruciferous crops. Its populations are likely to be affected by climate change (Liu et al. 2002). The species can complete three to four generations per year in temperate regions or as many as 20 per year in tropical regions (Harcourt 1957), depending on temperature. In North America, it overwinters in the southern United States and Mexico, where it reproduces con­tinuously south of approximately 36°N lati­ tude. A bioclimate simulation model predicted a northerly extension of the range of continu­ ous diamondback moth reproduction in North America by approximately 125 km for each 1.0 °C of ambient temperature increase, along with increases in the numbers of generations throughout vast regions of agricultural pro­ duction in North America with a warmer cli­ mate, particularly at more northern latitudes (O. O. Olfert, unpublished data). Invasive alien species are a threat to ecosys­ tem function and to world economies (e.g., agri­ culture and forestry), and many studies have reported on the impacts of a changing climate on invasion (Guo et al. 2012). Global trade and climate change greatly increase the potential for new invasive species to be introduced and become established. Differences in risk of inva­ sions among countries can arise due to the diversity of international trade routes, human destinations, and sources of visitors. Kriticos (2012) concluded that with warming climates, the southern latitudes of New Zealand will become suitable for the colonization of invasive alien species that are adapted to the warmer climates. By extrapolation, northern and south­ ern latitudes, in general, might become similarly suitable for colonization by invasive species that currently are restricted to warm‐temperate or subtropical regions (Kriticos 2012). Owing to  their potential to disrupt ecosystems and threaten biodiversity, invasive species are agents of global change, and climate change will increase that threat.

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Range shifts, contractions, and expansions, and changes in relative population densities resulting from a changing climate have been predicted for several North American insect species associated with agricultural ecosystems (Olfert and Weiss 2006; Mika et al. 2008; Olfert et  al. 2011, 2012, 2016). The wheat midge Sitodiplosis mosellana (Géhin) (Cecidomyiidae) is currently expanding its range in North America, and predictions of its eventual range vary from 19.6% to 27.3% of North America, depending on the underlying climate model (Olfert et  al. 2015). Similarly, 17% of North America would have a favorable habitat for the cereal leaf beetle, Oulema melanopus (L.) (Chrysomelidae), under current climates, but up to 35% of North America would be favorable under various future climate scenarios (Olfert et al. 2012). Across higher latitudes, ecological suitability is constrained by growing‐season fac­ tors and cold and heat. For example, Olfert and Weiss (2006) reported that cold stress limits the ability of O. melanopus to survive winters north of 60oN latitude. In general, at higher latitudes, distributions of invasive alien species are pre­ dominately limited by cold stress, and as this lessens, invasive insect species have opportuni­ ties to move poleward (Hance et al. 2007, Olfert et al. 2012). Model predictions of species expanding their range poleward in response to climate change are often associated with poleward range con­ tractions at the equatorial limits of their range (Olfert et al. 2012). The lack of suitable overwin­ tering sites, lower soil moistures and high soil temperatures at the equatorial limits resulting from a changing climate may be important fac­ tors for many species. In southern Saskatchewan, for example, current, mid‐summer soil temper­ atures can exceed the upper optimum tempera­ ture threshold for a number of invasive alien species (Olfert et al. 2012). Going forward, there are a number of issues of  vital importance to insect biodiversity due to  range expansion under a changing climate. In  relation to agricultural pests, considera­ tion must be given to the viability of agriculture,

based on climate and soil constraints, in the regions that are predicted to become newly infested. The range expansion or shift of insect pest populations is moot if the new region is not  suitable for host‐plant production. Hence, it  is  important to begin implementation of tritrophic modeling approaches (host–pest species–­ natural enemies) to range‐expansion studies. There is some evidence from northern continents, where agricultural production is currently not possible due to cold stress, that a suitable arable land base would be available for successful establishment of crop pests under a changing climate. Mills (1994) conducted a study of arable soils in northwestern North America (north of 55oN and west of 110oW) and predicted that if CO2 levels double (i.e., +3.8 °C; +17% rain), the availability of arable land could substantially increase in area to approximately that of the current amount of arable land on the  Canadian prairies. This prediction implies impacts on regional biodiversity and ecosys­ tem  function as a result of land‐use changes, increases in intensity of cropping practices, and invasions. The frequency of single or multiple species outbreaks requiring mitigation strategies (for economic or environmental reasons) under a changing climate has not been quantified to a large extent. Some insights can be gained from bioclimate modeling by overlaying predicted regions of favorable habitat for multiple species to determine whether there is greater overlap of the areas under a changing climate, compared with current climate. Changes in abundance and generation time are likely to have implica­ tions for pesticide applications, with direct and indirect effects on biodiversity. 26.3.1.2  Community Composition

The crops produced in most regions are expected to change over time as growers select and develop plant species and varieties that optimize yield and economic returns under the prevailing conditions (Cock et  al. 2013). Intensification of agriculture normally has adverse effects on insect biodiversity, often

26  Global Change and Insect Biodiversity in Agroecosystems

mediated through plants – reduced diversity of plants inevitably leads to a reduced diversity of  insect herbivores, which in turn leads to a  reduction in specialist natural enemies. Fertilization and liming of grassland encourage those grasses that grow most rapidly and push out the diverse community of plants that would support a corresponding diversity of insects. Removal of hedges and minimization of field edges reduce habitat diversity and eliminate nectar and pollen sources for natural enemies and pollinators (Stoate et al. 2001). More effec­ tive weed management does the same. The application of insecticides has strong negative effects on most insects (Chagnon et al. 2015), and the targets of such treatments are more likely to rapidly evolve resistance than are other insect species in the crop. Natural biological control and conservation biological control are highly dependent on landscape‐level processes and can be negatively affected by disturbance, landscape fragmentation, and loss of biodiver­ sity (Letourneau and Bothwell 2008; Letourneau et al. 2009, 2011). The biodiversity of insects is highly sensitive to the underlying and adjacent plant commu­ nities (Stamps and Linit 1997, Thomas and Marshall 1999, Söderström et al. 2001). For example, the removal of shade trees in coffee and tea plantations greatly reduces insect bio­ diversity. Coffee is traditionally produced under a canopy of shade trees, the diversity and architecture of which support a high biodiver­ sity of associated organisms, including insects. In Latin America, the recent trend of reducing this shade cover to increase production raises concerns about the potential loss of biodiversity (Perfecto et  al. 2004, 2005). Similarly, the con­ servation of even small forest fragments (e.g., in gullies and beside water courses) improves cof­ fee pollination by indigenous pollinators from these fragments (Klein et al. 2003, Ricketts 2004, Ricketts et al. 2004). Global change‐induced shifts in vegetation will modify habitat availability for insects (Woo­key et al. 2009). The responses are likely to be site‐specific and context‐dependent. The

response of invertebrates to regional climate shifts will probably be guided by human activi­ ties such as land‐use changes (habitat loss or fragmentation) and reduction of overall genetic diversity (Thomas 2010). For example, Lauss­ mann et al. (2010) reported that a decrease in butterfly species diversity occurred between the 1800s and 1900s after the introduction of large‐scale farming in Europe. In Australia, Hoffmann et al. (2008) reported the occurrence of shifts in the status of invertebrate pests over a 30‐year period, which was attributed to a changing climate and changing agronomic practices. The diversity and composition of insects and assemblages are expected to change with chang­ ing plant communities due to climate change variations (Kardol et al. 2011) and farmer responses. For example, the availability of leaf shelters in the canopy of white oak (Quercus alba) saplings can be an important determinant of species richness for the associated insect‐ herbivore community (Lill and Marquis 2003). If elevated CO2 (eCO2) levels increase plant productivity (i.e., they result in more leaf pro­ duction), then the number and diversity of shel­ ter‐building caterpillars in white oaks might increase, although no study has been conducted to test this hypothesis. If, due to climate change or the impacts of invasive species or other global change effects, a crop is no longer grown in a region, then the associated insect pest and ben­ eficial species will suffer similar declines. There are concerns for the persistence of some plant species in some European pastoral and low‐ input agroecosystems (Rühl et al. 2015). Because many of these weed species also support insect biodiversity (Marshall et al. 2003), this scenario has implications for the persistence and perfor­ mance of insect species associated with ecosys­ tem functions. An important constraint to range expansion for specialist herbivores and natural enemies might be the rate of movement by their host plant or prey (Harrington et al. 1999). In natural ecosystems, plants move at a slower rate than do their herbivores; specifically, the seed dispersal

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range is generally lower for plants than is the dispersal range of phytophagous insects (Kinlan and Gaines 2003). Specialist herbivorous insects might be able to follow a shift in temperature isoclines, but still be constrained because their plant resource will lag behind (Berg et al. 2010). In agroecosystems, farmers might take advan­ tage of global warming to grow new crops in a region, and therefore benefit from the absence of specialist herbivores. Although more com­ plex to analyze, multispecies‐interaction net­ works might help to forecast global warming effects on communities, with responses from positive (facilitation) to negative (inhibition) or  neutral. Furthering the mechanistic under­ standing of how interactions between spatially separated biota regulate responses to climate change is a tantalizing prospect for future research (A’Bear et al. 2014). Tracking the effects of global change on com­ munities is complicated. Communities of at least some insect groups in temperate regions, particularly Lepidoptera, are highly variable in space and time (Taylor and Taylor 1977, Woiwod and Harrington 1994, Conrad et al. 2004). It is not clear whether this has always been the case, or to what extent this reflects an ongoing response to change. It is certainly an added level of complexity for the measurement of changes in insect diversity and populations. 26.3.1.3  Other Responses to Climate Change

Faced with disruptive changes in their habitat, some species may be constrained from dispers­ ing and will either adapt to the new conditions or decrease and eventually become locally extir­ pated (Davis et  al. 2005, Cock et al. 2013). Implicitly, some species might both move and adapt; although we know of no examples from agroecosystems, a Californian butterfly moved uphill and adapted to a new food plant (Par­ mesan et al. 2015). Some in situ adaptation, either through phenotypic plasticity or evo­ lutionary selection, is expected under global change, especially where movement is not an option (e.g., on low, isolated islands) and when species have a short generation time and a high

rate of reproduction. In their review, Donnelly et al. (2012) found many reports of direct observations of phenotypic plasticity to climate change in species, but less conclusive evidence of genetic adaptation. Phenotypic plasticity constitutes a critical survival mechanism for adaptation (Thomas et al. 2001, Valladares et al. 2014). Moreover, evolutionary and plastic responses to climate change are not mutually exclusive. The concept of evolutionary rescue to arrest population decline and allow population recovery (Gonzalez et al. 2013) may be particu­ larly relevant, especially to island populations challenged by climate change. Species may adapt to the new conditions in situ. There are abundant examples of adapta­ tion, in short time frames, to rapid shifts in environment. Perhaps the most famous was Kettlewell’s (1955) study on the melanic adapta­ tion of the peppered moth Biston betularia (L.) (Geometridae) to soot‐blackened tree trunks, mediated by bird predation. For many years this example was considered a classic demonstra­ tion of the mechanisms of evolution in action, until doubts were raised about the experimental methods and design. Nevertheless, new experi­ ments by Michael Majerus reported after his death by Cook et  al. (2012) confirmed that natural selection by insectivorous birds was the driving force of this dramatic change. When the vegetable leaf miner Liriomyza tri­ folii (Burgess) first colonized Senegal in the early 1980s, it achieved outbreak levels as a result of poor performance of natural enemies, exacer­ bated by the use of insecticides. Following cessa­ tion of insecticide sprays, indigenous parasitoids of indigenous Liriomyza spp., now adapted to the new introduction, brought the pest under control (Neuenschwander et al. 1987). Intro­ duced biological control agents go through a genetic bottleneck in the process of introduc­ tion and establishment, and may go through another in adapting to their new environment, and sometimes to a new host or prey. Releases of  Pareuchaetes pseudoinsulata Rego Barros, an  erebid arctiine moth from Trinidad, for the biological control of the weed Chromolaena

26  Global Change and Insect Biodiversity in Agroecosystems

odorata (Asteraceae) in India and elsewhere were successful only in Sri Lanka. A population collected from Sri Lanka was released in India and then readily established. This material was subsequently mixed with fresh material from Trinidad, cultured, and released in Guam. New cultures from this stock were subsequently used successfully in other countries where introduc­ tions had previously failed (Zachariades et al. 2009, Cock et al. 2010). In the context of extinction as a consequence of climate change, species that live close to ther­ mal maxima are highly sensitive to increases in temperature, because these tend to have little variation in the upper critical temperature at which death occurs (Araujo et al. 2013, Vasseur et al. 2014). Species in the tropics might, there­ fore, be highly vulnerable to extinction as a result of increases in the severity and frequency of extreme heat events. For example, the cold‐ adapted bumblebee Bombus bellicosus Smith has become extinct in the former northern portion of its range in Brazil, whereas two co‐ occurring species with wider tolerances have become more abundant (Martins and Melo 2010). Climate‐change impacts on major vege­ tation types within biomes were modeled by Malcolm et al. (2006) who showed that a two‐ fold CO2 increase may increase extinction rates from  70%), which requires herbivores to have finely tuned host‐finding abilities. Chemical defenses are highly important; for example, the Neotropical tree genus Inga (Fabaceae) invests up to 50% dry weight of young leaves in secondary metabolites (Lokvam and Kursar 2005). Increases in tem­ perature, atmospheric CO2, and the length of the dry season are all likely to alter plant chem­ istry. For example, increased CO2 will shift the balance from nitrogen‐based defenses (e.g., alkaloids) to carbon‐based defenses (e.g., tan­ nins) (Coley 1998). 26.5.1  Climate Tolerances in Tropical and Temperate Species

We know less about how tropical ecosystems may respond to climate change than we do for mid‐ to high‐latitude systems (Stange and Ayres 2010). Tropical species are likely to be particu­ larly sensitive to global warming because they are adapted to limited geographic and seasonal

variation in temperature, they already live at close to the highest temperatures on Earth before global warming began, and they are often isolated from cool refuges. The tropics are now entering a set of human‐mediated changes (pri­ marily deforestation) and climatic changes, which in combination are without precedence; our future climate could have temperatures as high as in the Late Eocene, but with the possibil­ ity of widespread aridity and without the miti­ gating presence of megathermal moist forests in the high latitudes (Maslin et al. 2005). By 2100, under a moderate greenhouse gas emissions scenario, 75% of the area covered with tropical forests will experience mean annual tempera­ tures that are greater than the highest mean annual temperature that currently supports a closed‐canopy forest (Wright et al. 2009). Vulnerable components of forest communities might move upward and poleward to remain in their optimum temperature range, as new areas become suitable for tropical forests. However, these movements will be constrained by changes in rainfall patterns, patterns of human activity such as farming, and availability of refuges. A number of studies comparing species per­ formance from different climatic zones have concluded that warming in the tropics is likely to have more deleterious effects on inver­ tebrate communities that remain in situ than in temperate zones (Deutsch et al. 2008, Amarasekare and Sifuentes 2012, Hoffmann et al. 2013, Kingsolver et al. 2013, García‐Robledo et al. 2016). At least some tropical species that seem to have a broader temperature tolerance (and hence altitude range) are complexes of temperature‐specialized cryptic species, dis­ tinguishable using barcodes, but not morpho­ logically (García‐Robledo et al. 2016). Tropical insects are relatively sensitive to temperature change, are currently living close to their opti­ mal temperature, and are not expected to per­ sist in warmer tropical zones (Araujo et  al. 2013). By contrast, species in temperate zones have broader thermal tolerance and live in climates that are currently cooler than their

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physiological optima, and might benefit from a warmer climate (Deutsch et al. 2008). For example, figs (Ficus spp.) have an obligate pollination mutualism with tiny, short‐lived (1–2 days) fig wasps (Agaonidae). Adult females of four species of fig wasps from equatorial Singapore were tested for thermal tolerances. The results suggest that an increase of 3 °C or more above the current temperatures experi­ enced across much of the equatorial tropics would markedly decrease the active adult lifes­ pan of all four species. Fig plants are the center of an intricate web of specialist and generalist animals. Unless fig wasps acclimate or adapt to warmer temperatures, these responses could disrupt the mutualism, potentially affecting multiple trophic levels (Jevanandam et al. 2013). Using a large data set of field and museum records of rolled‐leaf beetles (Cephaloleia and Chelobasis in the family Chrysomelidae) in Costa Rica, García‐Robledo et al. (2016) found that species previously thought to have phe­ notypically plastic thermal tolerances were actually complexes of cryptic species with dis­ crete elevational ranges and thermal tolerances adapted to local temperatures. With a tempera­ ture increase of 3–6 °C predicted for Costa Rica in the next century, the authors concluded that populations of endemic cryptic species that have narrow elevational distributions and that live at the highest elevations face the highest risks of extinction as their thermal habitat dis­ appears and they are pushed off the tops of these mountains. Contrary to theory, when responses to ther­ mal extremes have been compared within related species, both temperate and tropical species are affected. For example, a study of 10 closely related Drosophila species with known tropical or widespread distributions found that adult tolerance to thermal extremes was corre­ lated with current distributions, with most spe­ cies experiencing heat exposure close to, but rarely above, their upper heat limit. Among widespread species, adult cold resistance proved a good predictor of species distributions in cooler climates (Overgaard et  al. 2014). Thus,

both tropical and temperate species could face a similar proportional reduction in range under future warming. Above‐optimal temperatures can have numerous effects on invertebrates, and repro­ duction might be at the crux. Although insect oviposition ceases at temperatures near the upper temperature threshold, experiments with other invertebrates indicate reproductive disruption at lower temperatures. Zeh et  al. (2012) subjected the Neotropical pseudoscor­ pion Cordylochernes scorpioides (L.) to the 3.5 °C increase predicted for the tropics. They found survival was significantly reduced and development accelerated at the cost of reduced adult size. There was a dramatic decrease in level of sexual dimorphism. The most striking effects, however, involved reproductive traits. Males produced 45% as many sperm as con­ trol  males, and females failed to reproduce. Sequencing of the mitochondrial NADH dehy­ drogenase 2 (ND2) gene revealed two highly divergent haplogroups that differed substan­ tially in developmental rate and survivorship but not in reproductive response to high temperature. Drought is expected to become more com­ mon in many tropical regions, and a lack of genetic variation in desiccation resistance is a further trait predicted to limit climate adapta­ tion of tropically restricted species, especially rainforest inhabitants. When exposed to humid­ ity levels at ecologically realistic levels of cli­ matic change and desiccation stress, significant evolutionary responses were found in two spe­ cies of rainforest‐restricted Drosophila (van Heerwaarden and Sgro 2014), suggesting that evolution might be an important adaptation by sensitive rainforest‐restricted species to climate change. Leaf‐litter quality, a major determinant of the size and diversity of decomposer communities (Boyero et al. 2011), is expected to alter under eCO2 levels and climate‐induced changes in plant‐community composition. Climate change is expected to result in higher rates of popu­ lation growth and have positive effects on the

26  Global Change and Insect Biodiversity in Agroecosystems

abundance of some species of temperate millipedes (Diplopoda) and woodlice (Isopoda) (David and Handa 2010). At low latitudes, inter­ actions with more severe droughts are likely and could affect community composition. Land cover changes, mainly due to deforestation in the tropics and land abandonment in Europe, are critical to habitat specialists and could over­ ride any other effect of global change (David and Handa 2010). Both tropical and temperate species are responding to global warming through altitudi­ nal shifts. In one of the few documented exam­ ples from the tropics, the average altitudes of 102 montane moth species in the family Geometridae collected on Mount Kinabalu in Borneo increased by a mean of 67 m between 1965 and 2007, during which period the average temperature for the 5° cell that includes Mount Kinabalu increased by 0.7 °C (Chen et al. 2009). However, the consequences of these shifts for whole communities are largely unknown. Using current altitudinal data for six taxonomic groups spanning 90 degrees in latitude, Sheldon et al. (2011) examined the potential impacts of climate‐driven range shifts on community change, or “disassembly,” across latitude. They concluded that tropical communities are more sensitive to temperature increases, compared with temperate communities. Mountain height can affect the amount of community disassem­ bly, with greater change occurring on smaller mountains. Endemic species currently occur­ ring on top of isolated mountains will be most at risk of extinction, as they cannot move upward (García‐Robledo et  al. 2016). However, these altitudinal shifts might lead to lowland biotic attrition. Assessment of plant and insect data on an altitudinal transect in Costa Rica led Colwell et  al. (2008) to conclude that tropical lowland biotas could face significant net attrition, while at higher latitudes the range shifts might be compensated for by species from lower latitudes. Although tropical species may have a narrow thermal tolerance compared with similar taxa at temperate latitudes, high spatial heterogene­ ity in temperature is correlated with greater

warming tolerance in insects globally (Bone­ brake and Deutsch 2012). Therefore, spatial heterogeneity and invertebrate behavioral responses could play a critical role in thermal adaptation and climate‐change impacts, par­ ticularly in the tropics. This heterogeneity can be over relatively small scales, as seen in other invertebrate species. During field observations of the tropical snail Littoraria scabra (L.), sub­ strate‐surface temperatures were heterogene­ ous (22.5 to 53.1 °C) at the centimeter scale, and L. scabra selected thermally favorable sites (i.e., between 22.5 and 33.4 °C) rather than micro­ habitat type (Chapperon and Seuront 2011). Tropical insects are well known to vary their activity patterns seasonally in response to high temperatures and low humidity, and differences between the wet and dry seasons can be pro­ nounced (Wolda 1988). Just as there is scope for individual species to show behavior that enables them to persist at higher temperatures in the tropics, so too is there scope to change agricultural methods, such as soil preparation, water management, and use of shade trees, to enable continued production at raised ambient temperatures. Recent observations on coffee growing illustrate this. Coffee production generates income for 100  million people, mainly in the developing world. However, the combination of climate change and the impacts of urbanization, with its resultant landscape modifications, creates a feedback loop whereby coffee production sys­ tems are adversely affected. This situation has accen­tuated damage by the coffee berry borer Hypothenemus hampei Ferrari (Curculionidae), the most serious biotic threat to global coffee production. However, in a traditional coffee production area in greater Nairobi, Kenya, Jaramillo et al. (2013) showed that the relatively simple strategy of planting shade trees decreased mean plantation temperature by 2  °C and increased berries per branch by more than 10%, compared with coffee grown in monoculture. Most importantly, whereas infestation levels of H. hampei in the open plantation exceeded the economic threshold on most sampling dates, it

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was never reached in the shaded plantation, possibly because shade trees act as a refuge for beneficial arthropods (native and introduced), leading to higher levels of biological control. Thus, several studies indicate that tropi­ cal  insects might have problems tolerating increased temperatures, leading to the risk of local extinction in situ. Alternatively, poleward and uphill movement can be expected and has been documented for many species, but where this is not possible due to geographical barriers, will tropical species become extinct? Not neces­ sarily; the world, at least in Antarctica, has been warmer in some previous interglacial peri­ ods, perhaps by as much as 3 °C (Petit et  al. 1999) – although estimates for the actual inter­ glacial temperatures in the tropics are not as robust as those for high latitudes (Metcalfe and Nash 2012). By implication, the species still pre­ sent today have been able to survive these past conditions. Behavioral adaptation and exploita­ tion of habitat heterogeneity can be expected to play an important role, enabling many species, including many in agroecosystems, to persist in situ in the tropics in a world that does not become warmer than it has done during the past 2.6 million years of the Quaternary Period.

26.6 ­Some Concluding Viewpoints For insects, changes in distribution and timing, declines in abundance, and outright extinctions are, and will continue to be, outcomes of ongo­ ing global change processes. Society depends on the networks of interactions that drive the ecosystem processes in agriculture, many of which are regulated by insect biodiversity. As these functions are diminished through declines in insect biodiversity, they may be supported with increased reliance on petro­ chemical energy (Swift et al. 2004), potentially exacerbating the outputs of greenhouse gases. There are, however, encouraging signs for con­ servation of biodiversity of insects and aware­ ness of the importance of ecosystem function in agroecosystems.

Among farmers, perhaps only the biodiversity of pollinator insects is universally viewed as positive and essential. Implementing mitigation strategies in this area might be relatively easy because there is a direct relationship between pollination and crop yields and because pollina­ tion services cannot easily be replaced with pet­ rochemical energy. Mitigating the effects of both climate change and agricultural intensifi­ cation on pollination requires the development of strategies to maintain pollinator diversity in highly managed landscapes. The best recom­ mendations for conserving pollinator diversity in agriculture involve managing multiple risk factors – that is, minimizing the risk to natural populations from pesticides, a return to IPM practices, and using land conservation or resto­ ration to maintain food and nesting resources throughout the year (Goulson et al. 2015). The results of habitat restoration in the form of hedgerows and pollinator enhancements are promising; such efforts increase the local abundance of pollinators, and pollinators on nearby flowering crops (Hannon and Sisk 2009, Morandin and Kremen 2013). All these strate­ gies are compatible with the overall goal of maintaining ecosystem services provided by insect biodiversity in agroecosystems (Wratten et al. 2012). However, intensification of agricul­ tural production in some parts of the world is likely to work counter to this. In contrast to awareness of pollinators, natu­ ral enemies and soil inhabitants are almost uni­ versally ignored. In general, IPM programs have almost exclusively addressed insects that are pests of crops, in contrast to considering the state of the community of organisms rele­ vant to the wider agroecosystem. This general lack of awareness means that pesticides used to control insect pests also have effects on non‐ target organisms through the network of inter­ actions in terrestrial food webs and can eliminate the natural enemies of some pests (Hunter 1996). In some cases, pesticide depend­ ence increases pest populations (Settle et  al. 1996, Han et  al. 1998). This present state of affairs is contrasted against an increasing

26  Global Change and Insect Biodiversity in Agroecosystems

awareness of the importance of insect biodiver­ sity in agriculture and the development of IPM strategies and programs that take ecosystem state into consideration (Gurr and Kvedaras 2010, Ragsdale et  al. 2011, Pons et  al. 2013, Brévault and Bouyer 2014, Roubos et al. 2014, Schellhorn et al. 2015). There is no doubt that, on the whole, global change will have serious negative effects on small island nations, especially on socioeco­ nomic conditions and biophysical resources, although some of these impacts might be reduced through effective adaptation measures (Nurse et  al. 2014). Although there are many studies of climate‐change impacts on mid‐ and high‐latitude islands (Webb et al. 1998, Le Roux et al. 2005, Bokhorst et al. 2007, Bokhorst et al. 2008), there are few equivalent studies on tropi­ cal small islands, especially with insects. As in continental regions, increasing global tempera­ tures could lead to altitudinal species range shifts and contractions on high islands, with an upward creep of the tree line and associated fauna (Benning et al. 2002, Krushelnycky et al. 2013). Reduction in the numbers and sizes of endemic populations caused by such habitat constriction and changes in species composi­ tion in mountain systems might result in the extinction of endemic species (Pauli et al. 2007, Sekercioglu et  al. 2008, Chen et  al. 2009, Krushelnycky et  al. 2013). Many small islands depend on a limited number of economic sec­ tors such as tourism, fisheries, and agricultural crops, all of which are climate‐sensitive. Thus, on the one hand, climate‐change adaptation is integral to social stability and economic vitality on islands, but on the other hand, government‐ and community‐adaptation efforts are con­ strained because of the high costs. This may constrain the mitigation of the effects of global change on biodiversity to a much greater extent than in continental regions, with consequent threats to the diversity of endemic island species. The greatest impact of climate change on world food security probably will be in tropical agroecosystems. The length of the growing

period is expected to decline by 5% or more across a broad area of the global tropics, includ­ ing heavily cropped areas of Brazil, Mexico, Southern and West Africa, the Indo‐Gangetic Plains, and Southeast Asia (Ericksen et al. 2011). Country poverty, rather than region, will dictate the potential to mitigate, with African and South Asian countries less able to respond than China or Latin America (Ericksen et  al. 2011). Conservation tillage and a ban on burning sug­ arcane crops are two strategies implemented in Brazil to preserve soil systems (Cerri et al. 2007). By contrast, areas of suitability for corn and beans, two key staple crops in Africa, could decline by 20–40% relative to the period 1970– 2000, while population growth, dietary change, and increased globalization of trade will further drive cropping‐system change and compound the risk of pest and disease outbreaks (Dinesh et al. 2015). The biodiversity of insects in agroecosystems is clearly important to both ecosystem sustain­ ability and food production. This biodiversity – as expressed by numbers of species, their abundance, and their functional linkages  –  is threatened by ongoing global change processes. If science is to deliver strategies that preserve biodiversity and enhance agroecosystem sus­ tainability for the benefit of future generations, then knowledge gaps need to be addressed and the resulting information and best practices developed, freely exchanged, and shared across borders.

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27 Digital Photography and the Democratization of Biodiversity Information Stephen A. Marshall School of Environmental Sciences, University of Guelph, Guelph, Ontario, Canada

The science of taxonomy has traditionally pro­ gressed by organizing species diversity into a Linnaean hierarchy that reflects the origin of biodiversity through sequential speciation events. The resultant biodiversity information is made broadly accessible through a taxo­ ­ nomic  framework that involves properly named, meaningful, and recognizable taxa, with at least  the first of these three criteria (“properly named”) rigorously regulated by the  rules of nomen­clature (for entomologists and other zoologists, the International Code of Zoological Nomenclature (ICZN)) so that taxa have proper (unique, universal, and stable) names. The recent history of taxonomy also has seen a great deal of emphasis on methodology for estab­ lishing meaningful (generally inter­ preted as “monophyletic”) higher taxa. But tax­ onomy has had a spotty record with regard to the third criterion, and there has traditionally been little emphasis on making biodiversity information broadly accessible, especially at the species level. As a result, many, if not most, named insect species are recognizable only by a few specialists at best. The anachronistic nam­ ing of species in sparsely illustrated monographs or, worse yet, in isolated and out of context spe­ cies descriptions such as those that still some­ times appear in regional faunal works, has been

one of the major impediments to making biodi­ versity information broadly accessible. But regional faunal works developed with the objective of making names accessible, rather than just making names for their own sake, represent an important means by which biodiversity information is rendered useful. Authoritative faunal works and reviews that interpret an existing taxonomic framework, translating it into a form that renders taxa recognizable in a regional context, can effi­ ciently render names and associated informa­ tion broadly accessible, as long as there is an underlying taxonomic framework to interpret. If the taxa of interest have not been revised, and higher taxa remain inadequately defined or spe­ cies remain either unnamed or unrecognizable, then regional faunal works are impractical. Good taxonomy provides the necessary founda­ tion for the study of biodiversity, and taxonomic revisions are generally a prerequisite to good taxonomy. Improving and broadening access to biodiversity information, then, must begin with increasing the coverage and accessibility of tax­ onomic revisions. Once revisions are in place, then reviews, faunal works, and digital libraries will follow, opening access to biodiversity infor­ mation to increasingly broad user groups and ultimately leading to a desirable democratization

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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of the discipline. A dearth of current, complete, and accessible taxonomic revisions and useful faunal reviews currently creates a bottleneck, or taxonomic imped­ iment, slowing or blocking identifications and, thus, blocking access to bio­ diversity information. Several relatively recent technologies have been touted as revolutionary solutions to the taxonomic bottleneck or taxonomic impedi­ ment, but the greatest acceleration of taxonomic progress and the biggest boost to democrati­ zation of biodiversity information has been driven by the combination of digital photogra­ phy and  effectively universal Internet access. I have argued elsewhere (Marshall 2008) that the widespread adoption of digital photography is equivalent to the widespread adoption of the microscope in its impact on insect taxonomy, and that impact is soon to be magnified as all aspects of the discipline become open to a much broader spectrum of participants. This process will happen through a corresponding explosion of digital insect collections, changing approaches to taxonomic description, and the development of new digital tools that will render more and more groups of insects as identifiable and acces­ sible as birds are today. This chapter is based on information and ideas available up to the sub­ mission date (13 September 2014) of the origi­ nal chapter manuscript.

27.1 ­The Digital Insect Collection Revisions provide the necessary foundation for meaningful and accessible taxon names, and collections of specimens have traditionally provided the foundation for revisions. Most published revisions are essentially interpreta­ tions and summaries of the characters and data associated with physical specimens, either bor­ rowed from museums or collected by taxono­ mists specifically for their work. The online posting of digital images of museum speci­ mens, such as online‐type libraries or libraries of specimen images for particular taxa, has recently begun to expedite the study of museum

specimens, making it more selective, faster, cheaper, and more accessible. Examples include the Museum of Comparative Zoology entomol­ ogy type database (http://insects.oeb.harvard. edu/mcz/), the Tachinidae image gallery (http:// www.nadsdiptera .­o rg/Tach/WorldTachs/ Tachgallery/Tachgalleryhom.html), and other image galleries such as the Carabidae of the World (http://carabidae.org/). But, with or without online images of museum specimens, the depth and breadth of a revision has tradi­ tionally been a function of the data associated with available specimens, either newly col­ lected or older museum specimens. The study of specimens from collections is fundamental to a number of biodiversity‐related disciplines and especially critical to revisionary taxonomy. Good revisionary taxonomy is impossible with­ out specimens on which to carry out detailed morphological study, and taxonomists rou­ tinely use collections to ascertain intraspe­ cific  variation and to work out phenology and geographic ranges. There is no question but that museum collections will remain central to  taxonomy and related bio­diversity studies. Museums, however, are no longer the only source of insect species records. Libraries of digital photographs of living insects are already taking on many of the functions of specimen collections, and these virtual col­lections will soon rival museums full of dead specimens as sources of data for revisionary taxonomy. The museum specimens traditionally used for revisionary taxonomy are typically accumu­ lated by a combination of general and special­ ized collecting, often supplemented by the results of intensive regional surveys. By way of  illustration, I am currently revising the New World Taeniapterinae, a mostly Neotrop­ ical subfamily of the acalyptrate fly family Micropezidae. Micropezids are relatively large and attractive flies, and thus routinely taken by general insect collectors. About a quarter of the specimens I have accumulated for this project are bycatch specimens collected casually by other taxo­nomists, reflecting the long‐standing tradition of putting precious field time to good

27  Digital Photography

use by collecting all sorts of things rather than just the one group of interest to the taxonomist doing fieldwork. This pleasant and cost‐effective tradition has been effectively extinguished over the past couple of decades as most bio­diverse countries have implemented strict require­ ments for collecting permits, often allowing a collector to take only limited numbers of one focal taxon and frequently imposing onerous limitations on export of specimens. Another quarter of the micropezid specimens used in my revisions were borrowed from museums that accumulated them through surveys, pri­ vate collector donations, student collections, or other means. Most of that material (other than that taken during recent survey projects in Colombia and Costa Rica) is more than 50 years old. The rest of the available specimens, includ­ ing many species not represented in museum collections, are from my own recent collecting trips undertaken with the Micropezi­dae as a main focus. This sort of dependence on a com­ bination of museum specimens with a broad but incomplete geographic coverage and more focused special purpose collections with a nar­ rower geographic coverage is typical of recent revisions, most of which suffer from significant gaps in distributional coverage. These gaps, which often lead to critical deficiencies in phy­ logenetic coverage of the taxon under study, often correspond to countries where collecting permits are difficult or impossible to obtain, or  areas where other factors render inter­ national research work impractical. Current trends toward increasingly restrictive require­ ments for collecting permits in many countries suggest that it will be harder and harder to fill these sorts of gaps with newly collected physi­ cal specimens. This serious problem is exacer­ bated by prohibitively obstructive regulations regarding export of any biological specimens from some biodiverse countries. Digital collec­ tions (of live insect images) offer a partial solu­ tion to these problems while at the same time providing new opportunities for taxonomists to make their work more extensive and acces­ sible, and providing opportunities for vastly

broader participation in original taxonomic research. The impact of digital insect collections is only starting to be appreciated by taxonomists because the technology has only recently reached the point where detailed photographs of living specimens can be taken by almost any­ one. One obvious application of this technology is for taxonomists working on appropriately field‐identifiable taxa (e.g., Micropezidae) to undertake fieldwork in permit‐unfriendly coun­ tries, using only photography. As a case in point, appropriately angled, adequately sharp photos of living micropezid flies belonging to recently revised groups can usually be identified to spe­ cies, and for a few selected genera it would be practical to confidently describe new species on the basis of photographs alone (yes, the ICZN does allow for the description of species based on “lost” or “released” living type specimens). But here, too, there are obstacles. It is unlikely that any granting agency is going to support entomological fieldwork just to take pictures, and photography for research purposes is also illegal without a (difficult or impossible to obtain) permit in some countries. But no such obstacles stand in the way of the armies of ama­ teurs, hobbyists, students, ecotourists, and oth­ ers currently accumulating millions of images, some of which will be compiled into accessible and useful collections. And it is these collec­ tions of digital photographs of living insects that eventually will become almost as impor­ tant to taxonomy as museums full of dead spec­ imens as sources of data. This is not a popular prediction among some entomologists, who offer the following usual objections: Insect taxonomy is based on properly and permanently labeled specimens, which meet at least minimum standards for locality information. Photographs are not as good. In fact, collections of photographs are equiv­ alent to collections of specimens in this regard. Both can be inadequately or inaccurately labeled. But many cameras today have GPS

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units that embed latitude and longitude in the image metadata. My images are much more carefully georeferenced than most of my speci­ mens. Many of the biologists, naturalists and photographers currently assembling image collections are doing so with the same sort of care they would put into specimen collections. There are tens of millions of specimens in museums, covering all taxa and all regions. For most taxa, relatively few identifiable digi­ tal photos are available. True, but this is changing with incredible speed. A few years ago a Google search on “Micropezidae images” yielded only a few dozen images, now it yields over a thousand, including hundreds of high‐quality images that warrant the same attention as the miscellaneous pinned micropezids in museum collections. I expect that a decade from now I will find more useful distributional records on Flickr and other photo sharing sites than can be extracted from speci­ mens in the British Museum or the Smithsonian. Currently private offline image collections are likely to have an even greater impact as they become publicly available through museum websites. Live images are nice, but they do not have taxonomic value like a museum specimen. Again, this is changing fast, and varies from group to group. In the Micropezidae, a good photo of a live individual can offer more useful information than can be extracted from a shriveled pinned specimen, but of course for some taxa it is impossible to separate closely related species without dissections of genitalia or other structures usually not visible in pho­ tos, and species in some groups apparently dif­ fer only at the genetic scale. Digital collections can only complement museum collections, not replace them. Live images are fine for big insects such as dragonflies, but inadequate for small insects.

Again, this is only partially true and is chang­ ing fast. The standard 100‐ or 105‐mm macro lenses now carried by many naturalists can cap­ ture a good image of a mosquito‐sized insect. With added extension, crisp shots of aphids and smaller things are already within reach of any serious photographer, and the technology is moving forward with breakneck speed. I fully expect that any amateur naturalist with a smart­ phone will be able to take useful photos of Drosophila‐sized insects in the near future. Most image collections are in private hands, and cannot be borrowed for revisionary taxonomy. Possibly true, for the moment, but this was also once true for many important insect speci­ men collections that started out as private col­ lections and are now publicly accessible in major museums. It is cheaper for a museum to acces­ sion an image collection, especially a digital one, than a specimen collection. Estates or individu­ als routinely contact me about donating private insect specimen collections to the University of Guelph Insect Collection. But even though our collection, like most regional collections, had its start as an aggregation of amateur collections, it is not practical to accept most such donations. Even if we had the space and time to incorporate and curate the material, amateur specimen col­ lections are often in bad shape and lack ade­ quate data. The same is likely to be true for some image collections, but my guess is that the new generation of digital insect collectors will ultimately leave an enormous legacy of high‐ quality images and associated data that will lend themselves to organization and archiving just as physical insect collections are curated. It is difficult to estimate how much effort is currently going into the development of digital insect collections. But let us make a conserva­ tive guess that at least 1% of the serious natural­ ists now combing every bit of natural habitat on the planet to photograph birds and other verte­ brates will make the natural transition to pho­ tographing more diverse groups of animals.

27  Digital Photography

According to US Fish and Wildlife Service fig­ ures, in 2006 some 48 million birdwatchers spent 36 billion dollars on trips and equipment (Carver 2009), and birding and related nature watching is a fast‐growing hobby worldwide. Birding is the number one hobby in the United Kingdom, and similar wildlife watching activi­ ties are on the upswing around the world, so it is not unreasonable to estimate that there are at least 100 million active wildlife watchers world­ wide. Many, if not most, of these birders and other naturalists now carry digital cameras with macro functions, and part of this unprec­ edented pool of observers inevitably find that insect watching is more satisfying than bird watching; many insect watchers will go on to develop serious and useful image collections and some will accumulate collections that run into the tens of thousands. My conservative guess of 1% means a million insect photogra­ phers, of whom tens, if not hundreds of thou­ sands, are quietly accumulating significant collections, often identifying their images with paper and digital guides. Most of these collec­ tions will, at least in the short run, remain offline and in private hands, but some idea of the explosion in insect imaging can nonetheless be gleaned by looking at those insect image col­ lections that are being made available online. The explosive growth of online image collec­ tions is perhaps best illustrated by looking at the statistics for the popular (and insect‐rich) photo‐sharing site Flickr, which reported host­ ing more than 6 billion images as of August 2011. According to a more recent report in The Verge (Jeffries 2013), more than 3.5 million new images are uploaded to Flickr daily, so you can add another billion or two to that total. Flickr houses the largest aggregation of insect images from throughout the world, but many other photo sharing sites, such as PBase, also house some important collections of high‐quality insect images. Some of these are already having a significant impact. The Tom Murray collec­ tion in PBase, for example, includes tens of thousands of images covering more than 8000 identified species, and parallels the best private

specimen collections for species coverage and data quality. Many other important online digi­ tal insect collections can be found on personal websites, such as Stephen Cresswell’s “American Insects” site covering New World insects. As of May 2014 Cresswell told me that he had 61,996 images with at least basic data, with 3328 spe­ cies represented by “species pages” on his web­ site. Both Murray and Cresswell are amateurs in the finest sense of the word. Tom Murray is a sales representative, and Stephen Cresswell is a history professor. There are many comparable online collections posted by individual ama­ teurs in other parts of the world, such as Peter Chew’s extensive image library of Queensland insects, but perhaps the most significant digital collections are multiphotographer collections on the many websites dedicated to particular taxa. The European website DipteraInfo, for example, has galleries of Diptera images organ­ ized into a taxonomic hierarchy equivalent to the organization of a regular insect collection, and the North American website BugGuide.net is an extensive and superbly organized digital insect collection generated by tens of thousands of photographers and dozens of volunteer cura­ tors. BugGuide is emerging as such an impor­ tant phenomenon that it deserves further comment. BugGuide.net was established by Troy Bartlett in 2003 and has been maintained by John VanDyk at Iowa State University since 2006, at which time it consisted of a few thou­ sand images, generally posted by amateurs and mostly identified by volunteers (amateur and professional) visiting the site. BugGuide.net now includes content by around 28,000 con­ tributors who have posted over 800,000 photos to the site, now organized into more than 30,000 individual species pages. This already exceeds a quarter of all North American insect species and there is little doubt that most Nearctic insect species  –  including almost all field‐­ recognizable species  –  will be represented by BugGuide.net images within the decade. This is an enormous accomplishment with impor­ ­ tant  ramifications for the democratization of

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biodiversity information. Although the site was initially set up to serve a mostly amateur com­ munity who wanted help getting their images identified, it is organized along the lines of a museum collection with most contributions linked to at least basic data including locality, date, collector, and usually determiner. It has, thus, come to serve some of the important roles traditionally served by major insect collec­ tions  –  documenting distributions, exploring phenology and other natural history attributes, and checking identifications. BugGuide.net has opened up insect identification to the broad amateur community, but it is also now the first place many professional biologists turn to check North American insect identifications. And it is starting to significantly supplement traditional museum collections as a source of data for pri­ mary publications. Refereed publications in the Canadian Journal of Arthropod Identification, for example, routinely use data and images from BugGuide.net (with permission from, and credit to, the BugGuide.net contributors), and the unprecedented range of taxa imaged in these online collections has enabled other types  of publications that have also increased access to biodiversity information. The recently published photographic guide to beetles of eastern North America (Evans 2014), for exam­ ple, sets new standards for photographic guides by providing the naturalist community with the tools necessary to identify some 1500 beetle species. Most, of the images that made this book possible came from the new community of digital insect collectors. For example, the aforementioned Tom Murray contributed more than 300 images. I have so far dwelled on the astonishing explo­ sion of online digital insect collections, but online collections represent only the tip of the insect image iceberg. Most professional insect taxonomists now incorporate field digital pho­ tography into their entomological agenda, and many are accumulating broad collections of quality images just as insect collectors of previ­ ous generations accumulated broad collections of specimens. Only a small percentage of this

new generation of professional digital insect collectors posts images online. My own collec­ tion, for example is maintained on an array of hard drives, and I spend my evenings organiz­ ing it just as I would organize a specimen collec­ tion. Instead of drawers and unit trays full of specimens I have folders and subfolders full of high‐resolution TIFF files. Instead of paper data labels I have coordinates and other information in the metadata and basic collection data block‐ copied onto all the images from a single collec­ tion event. It is not a priority for me to put my collection online, because I am working with it constantly. But when I stop working on it, I hope someone will recognize the collection as a significant legacy, and I hope it will ultimately be available to generations of taxonomists to come, just as the current generation of taxono­ mists uses the Melander Collection, the Hull Collection, and other once‐private specimen collections now accessible at major museums. In the meantime, parts of my collection are rou­ tinely used in books, digital keys, and taxo­ nomic revisions. The use of digital images in interactive keys and taxonomic revisions is of particular significance, and is discussed fur­ ther below.

27.2 ­Digital Images in Interactive Keys Currently, insect diversity is organized around a taxonomic infrastructure that has been devel­ oped over centuries of descriptive and revision­ ary taxonomy, and it is largely inaccessible to the public. Even though primary taxonomic papers are increasingly freely available on the Web, most are useable only by other special­ ists  or by biologists with access to a reference collection. This situation is due, in part, to the taxonomic tradition of compact and jargon‐ rich text, and in part due to the historic diffi­ culty and cost of producing and publishing the extensive illustrations necessary to render taxo­ nomic works easily interpreted. As a result, much of the existing taxonomic infrastructure

27  Digital Photography

needed to access biodiversity information is essentially backlogged, waiting to be updated and translated into a more widely accessi­ ble form. Now that the tools and technologies are avail­ able to easily and cheaply translate existing tax­ onomies into user‐friendly and widely accessible reviews and keys, rapid gains in accessibility to  biodiversity information can be made by reviewing regional faunas to generate user‐ friendly and richly illustrated regional reviews and photographic keys. The appearance of such keys, especially open‐access Web keys, effec­ tively puts the reviewed taxa in the public domain, thus democratizing access to biodiver­ sity information. This is part of a process that has been going on in entomology since the first good field guides to butterflies appeared, and is continuing through the development of photo­ graphic field guides to dragonflies, tiger beetles, and other relatively conspicuous groups. I think of butterflies and dragonflies as “honorary birds” in the sense that they are now in the pub­ lic domain and available to naturalists, just as birds have been since the publication of the first Peterson Field Guides in the 1930s. But the democratization of other taxa is now occurring at an unprecedented pace because of the explo­ sion of digital field photography. This democra­ tization is in part because more and more naturalists are making digital image collections and, in part, because increasing numbers of professional entomologists are using digital images to Web‐publish photographic keys to more and more groups. The Canadian Journal of Arthropod Identification provides a case in point. The Canadian Journal of Arthropod Iden­ tification (CJAI) (http://www.biology.­ualberta. ca/bsc/ejournal/ejournal.html) is a peer‐reviewed, Web‐based journal devoted to the publication of works that contribute significantly to the recognition and documentation of Canada’s arthropod fauna. Although the digital keys at the heart of CJAI publications generally rely heavily on digital specimen images, most of the issues so far published also make extensive use

of digital field images. Live images are used to illustrate keys, to provide habitus illustrations for species pages, and to populate galleries used for rapid identification. Several CJAI publica­ tions also have used live images from public image libraries, such as BugGuide.net, as the sole basis for new records or revised distribu­ tions. For example, Buck et al. (2008) recorded a new species of potter wasp from Michigan, Illinois, and Nebraska based solely on images from BugGuide.net. Forty‐one of the 100‐plus field images in the same publication are cred­ ited to naturalists who posted their images on BugGuide.net. Buck et al. (2008), in turn, gener­ ated new interest in eastern North American Vespidae among photographers and naturalists, who are now able to identify the hundred or so species of northeastern Vespidae, using the exhaustively illustrated digital keys therein. There is, thus, a positive feedback loop between specialists writing open‐access digital keys, nat­ uralists providing images and data for the keys from their digital collections, and photogra­ phers and naturalists using the keys to identify their image collections.

27.3 ­Digital Photography and Taxonomic Revisions Primary taxonomic revisions are becoming much more widely accessible than in the past, thanks to online publication, free or low‐cost publication of color images, and readily availa­ ble tools for capture of high‐quality character and specimen images. Most taxonomists now collect field images as well as specimens, and revisions are now routinely enhanced with field  photographs that improve descriptions and diagnoses, making the work more accessi­ ble. For example, recent revisions in the Micro­ pezidae (Marshall 2011, 2013, 2014, 2015) incor­­porate field photographs for many of the species covered. These images are generally superior to images of pinned specimens, which are almost invariably faded and shriveled, and photos often include information not available

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from pinned specimens. This additional infor­ mation can include biological information (e.g., oviposition, adult feeding), behavioral informa­ tion (e.g., mating behavior), and information about features not generally visible  on dead specimens. For example, the soft abdominal pleuron of many Micropezidae has a species‐ specific pigmentation pattern clearly visible on field photos but usually altered or lost on dead specimens. Males of many species also have membranous pleural pouches clearly visible on field photographs, but difficult to discern on preserved specimens. These features not only render digital images of live material useful for diagnoses and descriptions, they also often allow reliable identification of micropezid images in digital insect collections made by naturalists. I predict that the data from these rapidly expand­ ing digital insect collections will become a criti­ cal source of distributional data for future revisions. Exhaustive regional collections made by dedicated photographers and naturalists based in areas of significance will be of particu­ lar importance. Research stations, educational institutions, and even eco‐lodges are likely

homes for these regional digital insect collec­ tions. The potential for research station and eco‐lodge digital image collections to contrib­ ute to taxonomy is immense, because there is an eco‐lodge or “birder lodge” close to almost every patch of attractive, high‐endemism habi­ tat on the planet. Many stations and lodges currently have partial image libraries of larger things such as moths or butterflies, but macro­ photography is increasingly becoming a core activity of the hundreds of thousands of people spending field time in these attractive places. I fully expect that insect taxonomists a decade from now will routinely curate parts of these regional digital image collections just as we now routinely curate our taxa in regional specimen museums. And, in turn, taxonomists will have access to data from a much wider range of local­ ities than are currently represented in tradi­ tional museums. The explosion of digital insect collections is still a new phenomenon, and its impact is just starting to be felt. In fact, it is still considered newsworthy when someone discovers a new species based on a digital insect collection.

Figure 27.1  Mating pair of Sapadrama (Tephritidae: Trypetinae). Photo by S. A. Marshall.

27  Digital Photography

For  example, the California‐based taxonomist Shaun Winterton recently discovered a new Malaysian species of green lacewing among images posted on Flickr, much the way taxo­ nomists routinely discover new species by perusing drawers of specimens in museums. Winterton contacted the photographer (Hock Ping Guek) in Malaysia, who was able to col­ lect  some specimens and later coauthored a formal description of the species along with Winterton and a colleague at the British Museum (Winterton et  al. 2012). Perhaps because of the catchy subtitle of the paper (“the confluence of citizen scientist, online image database and cybertaxonomy”) it received a great deal of publicity both online and offline. Curiously, some of the Web postings on the subject were critical of Winterton for taking first authorship and for naming a species first found by Guek. These postings were by indi­ viduals who failed to understand how much work and expertise goes into putting a speci­ men or image into the context necessary to rec­ ognize it as new and to formally describe it. Taxonomists working with museum collections routinely credit the collectors of those museum specimens within the text of published papers, and sometimes recognize them with patronyms, but including collectors as coauthors is unusual. Guek earned second authorship not just because there was an undescribed species in his online image collection, but because of his active involvement in looking for further specimens, illustrating the paper, and doing subsequent special‐purpose photographs of the new species and its habitat. Insect collectors are generally pleased at the prospect that their specimens might be used in a species description and that the associated data will form part of the back­ bone of taxonomy. Collections of digital images will inevitably come to be treated the same way, with the data recognized as public domain and the images themselves covered by copyright, just as specimens in some major museums are covered by copyright. For example, permission is needed to publish an image of a specimen from the British Museum of Natural History

(the Natural History Museum), and it could be argued that an image of a living specimen held in a museum’s digital collection of insect images is an equivalent entity. Long‐distance collaborations between pho­ tographers and taxonomic specialists are not new; what is new is the exploding volume of high‐quality, information‐rich images awaiting taxonomic assessment. I recently coauthored a  description of a new genus and species of Tephritidae in a long‐distance collaboration with an Australian tephritid specialist who has  yet to see a specimen of the taxon being described (Hancock and Marshall 2012). My role was as photographer‐naturalist answer­ ing  the specialist’s questions and sending him photographs (e.g., Fig. 27.1) until we were both satisfied that the description was complete. Admittedly, we were both professional dipter­ ists, but it is not that much of a stretch to see a similar collaboration between amateur digital specimen collectors and specialists. Nor is it a stretch to see the explosion of curated image collections and photographically rich descrip­ tions, keys, and reviews empowering a broad range of amateurs to recognize, and perhaps even describe, species in their own collections. The upshot will be bigger and better image col­ lections, which inevitably will become main­ stream components of taxonomic work by an increasingly broad community. A small foretaste of the impending impact of  image libraries appeared in the wake of some recent revisions (Marshall 2011, 2013) of Caribbean clades of Micropezidae with high island‐level endemism. Those revisions were based mostly on museum specimens of varying quality, and suffered from the inevitable gaps in museum material. These papers explicitly recognized a lack of study material from some islands, such as Guadeloupe and Martinique, and lamented that material from some other islands (such as Jamaica) was scarce or of low quality. Web searches at that time failed to turn up any images of use to fill gaps, but in the short time since those revisions were pub­ lished, several superb micropezid images from

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Martinique, Guadeloupe, and Jamaica have appeared in online digital insect collections. The Martinique and Guadeloupe images (Gral­ lipeza placidoides and Grallipeza spinuliger) represent new records for the islands, and the Jamaica images (http://www.americaninsects. net/f/micropezidae.­ html) are beautiful and informative shots of a species (Grallipeza cliffi) that I had named based on a couple of damaged, discolored museum specimens. In another decade, I fully expect to see every island, and every Caribbean micropezid species, repre­ sented in digital insect collections. Availability of such a resource a couple of years ago would have rendered those recent revisions much more complete and much more accessible.

27.4 ­Organization of Digital Insect Collections Collections of digital images of living specimens and collections of dead specimens have some obvious parallels. Both are accumulations of collection events, and their value depends on the accuracy and detail with which those collec­ tion events are documented, and the quality of the specimen or image collected. A poor speci­ men with little data is of little use; the same is true of a poor or inadequately documented image. Specimens in collections are organized hierarchically, typically with each family in a cabinet divided into genera in different drawers, with the drawers in turn divided into unit trays for species. Digital images can be filed physi­ cally the same way, while they can also be organ­ ized virtually by any number of parallel systems or keywords, online or offline. Specimen collec­ tions are usefully aggregated into a limited number of major museums where taxonomists and other users can identify specimens (curate) and extract data, either onsite or by loan; collections outside these major museums are not as accessible. Digital image collections are also most useful when aggregated into well‐­ organized major sites such as BugGuide.net, although any appropriately tagged online image

is theoretically searchable and thus accessible. There are parallels between useful large museum collections of specimens and useful aggrega­ tions of digital images, as both rely on a synergy between the taxonomic community (providing determinations) and a broader community of collectors that provide the specimens or images and associated data. These parallels suggest that major digital collections have the potential to evolve into museum‐like repositories of acces­ sible biodiversity information, more widely accessible and easier to maintain and grow than traditional museums. To my knowledge, no significant collections of digital images of living insects are managed and developed by a major museum in parallel with their collections of preserved specimens. The development and professional curation of a  digital insect collection in parallel with a respected museum collection would attract donations of digital image collections by seri­ ous digital insect collectors, including the many naturalists and scientists (myself included) with large image collections maintained only on local hard drives. A digital collection with the same policies for growth and curation as a major collection would provide a significant resource for taxonomists and users of taxon­ omy alike, but few of the available online image libraries approach this ideal. Flickr, for exam­ ple, houses enormous numbers of high‐quality images, but associated data are inconsistent and taxonomic organization is opportunistic and uneven. The many citizen‐science image collections such as iNaturalist and Project Noah (which stands for Networked Organisms and Habitats) also lack professional curation; iNatu­ ralist images are often gleaned from Flickr, and Noah was conceived as an app for sharing cell­ phone images. EOL hosts many images, but they are mostly gleaned from other sites such as Flickr. Probably the digital image collection that most closely approximates a museum collec­ tion in organization and function is BugGuide. net. BugGuide.net describes itself as “an online community of naturalists who enjoy learning about and sharing our observations of insects,

27  Digital Photography

spiders, and other related creatures”, but it has been tremendously successful in developing into a comprehensive virtual collection of liv­ ing  insect images from the United States and Canada. One of the limitations of BugGuide.net as a virtual collection of use to taxonomists is its scope, which is explicitly North American when the greatest need, at least from this insect taxonomist’s point of view, is for major, well‐ organized digital collections of tropical insect images. There is a great window of opportunity here for the great museums to fill this gap and to develop virtual collections to parallel their physical collections, without the limits to growth that currently constrain collections of dead specimens.

27.5 ­Conclusions Digital photography already has provided unprecedented access to biodiversity informa­ tion by allowing naturalists to use online photo­ graphs, often organized into user‐friendly interactive keys, to identify an ever‐growing proportion of the insects routinely seen and photographed. This achievement in itself is a democratization of biodiversity information, but it is only the start. The increasingly routine activity of capturing images as part of nature watching, followed by the pleasures of organiz­ ing and identifying those images, using printed or online reference photos, is already leading to the widespread development of huge collections of digital images of living insects. As technolo­ gies improve and significant groups of amateurs, photographers, and professional entomo­logists accumulate more, higher quality, and more var­ ied images, a sort of feedback loop is starting to develop. Digital insect collectors are educated and inspired by other digital insect collectors, and many are interacting with professional taxonomists by providing images and data and receiving identification help. The continued involvement of a rapidly expanding  –  and increasingly diverse – international community of digital insect collectors must be embraced by

the taxonomic community not only as a source of incredible amounts of useful data, but also as a welcome and inevitable democratization of the discipline.

­References Buck, M., S. A. Marshall and D. K. B. Cheung. 2008. Identification atlas of the Vespidae (Hymenoptera, Aculeata) of the northeastern Nearctic region. Canadian Journal of Arthropod Identification. Doi: 10.3752/ cjai.2008.05. Carver, E. 2009. U.S. Fish and Wildlife Service: Birding in the United States: A Demographic and Economic Analysis Addendum to the 2006 National Survey of Fishing, Hunting, and Wildlife‐Associated Recreation. USFWS report 2006‐4. 16 pp. Evans, A. 2014. Beetles of Eastern North America. Princeton University Press, Princeton, New Jersey. 560 pp. Hancock, D. L. and S. A. Marshall. 2012. New records of fruit flies from Northern Vietnam, with description of a new genus and species of Adramini (Diptera: Tephritidae: Trypetinae). Australian Entomologist 39: 55–64. Jeffries, A. 2013. The man behind Flickr on making the service ‘awesome again’. The Verge. http://www.theverge. com/2013/3/20/4121574/flickr‐chief‐markus‐ spiering‐talks‐photos‐and‐marissa‐mayer [Accessed April 2016]. Marshall, S. A. 2008. Field photography and the democratization of insect taxonomy. American Entomologist 54: 207–210. Marshall, S. A. 2011. A review of the genus Hoplocheiloma Cresson (Diptera: Micropezidae). Zootaxa 2806: 1–23. Marshall, S. A. 2013. Grallipeza Rondani (Diptera: Micropezidae: Taeniapterinae) of the Caribbean and North America. Zootaxa 3682: 45–84. Marshall, S. A. 2014. A review of the Afrotropical genus Aristobatina Verbeke (Diptera: Micropezidae: Taeniapterinae), with

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descriptions of three new species from the Eastern Arc Mountains of Tanzania. African Invertebrates 55: 143–155. Marshall, S. 2015. Mesoconius Enderlein (Diptera, Micropezidae, Taeniapterinae) of Central America. Zootaxa 3914: 525–540.

Winterton, S. L., H. P. Guek and S. J. Brooks. 2012. A charismatic new species of green lacewing discovered in Malaysia (Neuroptera: Chrysopidae): the confluence of citizen scientist, online image database and cybertaxonomy. Zookeys 214: 1–11.

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28 Bee (Hymenoptera: Apoidea: Anthophila) Diversity Through Time Sophie Cardinal Canadian National Collection of Insects, Agriculture and Agri‐Food Canada, Ottawa, Ontario, K1A 0C6, Canada

Bees are a diverse, worldwide group of ecologi­ cally and economically important pollinators. The most well‐known species, the domesticated western honeybee (Apis mellifera L.), is only one of more than 20,000 described species. Not only are bees diverse in number, they are also highly variable in appearance and behavior. The present diversity has evolved over the past ~125 million years (Cardinal and Danforth 2013, Peters et  al. 2017) throughout all major terrestrial biomes, and has led to our reliance on the pollination ser­ vices provided by bees for much of the food we eat. This chapter will focus mainly on the diversi­ fication of bees through time. Following a brief summary bee biology, given in more detail in Michener (2007), I will discuss which lineages are more species rich or poor relative to their age, as well as where and when bees experienced shifts in their diversification rates.

28.1 ­Morphological Diversity Bees are generally described as hairy vegetarian wasps, but many are actually quite smooth and shiny, especially if they do not carry pollen on their bodies. Some bees (e.g., Hylaeus species) instead carry pollen in their crops, and parasitic bees forgo collecting pollen altogether. Bees that

carry pollen externally tend to have specialized areas and structures on their bodies formed by hairs (scopae or corbiculae) to store pollen during transport. These specialized areas vary in  their position, but are most commonly on the hind tibia, hind femur, underside of the abdo­ men, or side of the propodeum. Some bees, such as species of Rediviva (Melittidae: Melittinae), have specialized hairs, and sometimes their front legs are longer than their bodies and adapted for collecting floral oils from flowers with long slen­ der oil spurs where oil is secreted. Bees vary greatly in size, with the smallest bees being only about 1.8 mm long and the largest bee, the leaf‐cutter Megachile pluto Smith (Megachilidae: Megachilinae), measuring 39 mm (Michener 2007). Some bees are entirely black, whereas others are bright metallic green, such as some orchid bees (Apidae: Euglossini) that occur in the Neotropical Region. Numerous parasitic species have bright red integument or color bands, as in many Nomada, or contrasting bands of hairs, which in some species of Thyreus (Apidae: Melectini), are a shocking electric blue. Bees, at times, can be difficult to differentiate from the wasps they evolved from, but despite their varia­ bility, bees will have at least some hairs that are branched, and usually they have a hind basitarsus that is broader than the succeeding tarsal segments.

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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28.2 ­Behavioral Diversity: Social, Nesting, and Floral Hosts Unlike the honeybee, most bees do not live in colonies or store honey, but are solitary, with each adult female making her own nest, provi­ sioning it with pollen, nectar, or floral oils, and laying her own eggs. Some solitary bees, espe­ cially those that are active in early spring, form large aggregations, with thousands of bees nest­ ing in one area. Other bees live in colonies con­ sisting of two or more females with varying degrees of cooperation, from none (i.e., com­ munal bees), to highly specialized division of labor (i.e., eusocial bees), as in honeybees, sting­ less bees, bumblebees, and some sweat bees and carpenter bees. Parasitic bees forgo nest con­ struction altogether and have their host bees provision their offspring. Social parasitic bees infiltrate their host’s colony and replace the queen so that host workers rear their eggs, as in some bumblebees, Halictini, and Allodapini. Brood parasites, commonly referred to as klep­ toparasitic bees, target solitary hosts. In these species, the adult female sneaks into the host nest, lays an egg in a cell and in some cases kills the host egg, and leaves her larvae to feed on the pollen ball provided by the host bee for its own offspring. If the adult parasitic female does not kill the host egg, a specialized larval instar  kills the host immature (Alves‐Dos‐ Santos et al. 2002). The majority of bees nest in the ground (e.g., Andrena, miner bees), but some species exca­ vate nests in wood (e.g., Xylocopa, large carpen­ ter bees), use existing burrows made by other insects (e.g., Tetrapedia), burrow in pithy stems (e.g., Ceratina, small carpenter bees), or nest in existing cavities such as rodent burrows (bum­ blebees) or tree hollows (honeybees) (Michener 2007). Typically, the nests of solitary bees that burrow in the ground consist of a main burrow that gives rise to numerous lateral burrows, each ending in a single cell that serves to protect the larva and the pollen ball on which it feeds while developing. The cell surface is usually tamped smooth, and some bees secrete a film

that they paint on the cell wall to make a cello­ phane‐like lining. Megachiline bees line their cells with foreign material such as disks of leaves, plant hairs, mud, resin, and small peb­ bles. These bees tend to nest in small cavities or existing burrows and even abandoned snail shells (Michener 2007). Some bees, such as the honeybee Apis mellifera, are polylectic; that is, they collect pollen and nectar from a diversity of floral hosts. Other bees, such as Proteriades bullifacies, are oligolectic, collecting resources from only a restricted subset of plants (Cane and Sipes 2006). Bees tend to have a wider breadth for their nectar hosts than for their pollen sources (Robertson 1925). For example, the oligolectic bee Macropis nuda collects pollen and floral oil only from its host plant, Lysimachia, but col­ lects nectar from other, unrelated plants. Early bees probably were host‐plant specialists, and generalists probably evolved from these special­ ist bees (Müller 1996).

28.3 ­Geographical Diversity Unlike many other insect groups, bees are not the most species rich in the tropics, but instead are most species rich in more arid environments (Michener 2007). The number of described spe­ cies known from each country (Fig. 28.1), based on political boundaries, can be misleading due to the size of the country and the diversity of habitats and amount of sampling in the country. Numerous countries around the Mediterranean Sea are, nonetheless, comparatively species rich, given their size. Within the species‐rich United States, there is especially high abundances of wild bees in chaparral and desert shrublands, and intermediate abundance in temperate for­ ests and grasslands and rangelands (Koh et  al. 2016). In Canada, grasslands support the high­ est bee diversity and have the highest level of endemism (Sheffield et  al. 2014). Although grasslands are the most extensive of the North American plant formations, in Canada they are restricted to its southern edge (Barbour and

28  Bee Diversity Through Time

No. of species 0 1–359 359–718 718–1077 1077–1436

1436–1796 1796–2155 2155–2514 2514–2873 2873–3232 3232–3591

Figure 28.1  World map with each country shaded according to the number of described species recorded by Ascher and Pickering (2016). Alaska does not have more than 3200 species, but is shaded, being part of the United States of America.

Christensen 1993). The range of climatic condi­ tions in grasslands, the diverse assemblage of associated forbs dominated by Fabaceae and Asteraceae, and the drastic change in species composition that takes place throughout the growing season (Graham 1999) probably con­ tribute to the high bee diversity in grasslands. Grasslands have been reduced in size more than any other major biome in North America (Gauthier et al. 2003), and declines continue in some areas.

28.4 ­Evolutionary History and Diversification This discussion on the ages and diversification rates of bee lineages will predominantly be based on the most comprehensive time‐­ calibrated

phylogeny published for bees (Cardinal et  al. 2018). This phylogeny was based on a Bayesian analysis using an uncorrelated relaxed molecular clock calibrated with 35 points based on the bee  fossil record, seven gene fragments,  and more than 55% of extant bee genera (Fig. 28.2). The mean crown age (i.e., age of the most recent common ancestor of all extant species) for line­ ages is given, unless otherwise noted. Crown group bees evolved an estimated 125 (114–137) million years ago (mya) and started diverging from their wasp ancestors 145 (127–156) mya. Previous time‐calibrated phylogenies based on much more sequence data but fewer taxa have estimated that bees originated 93–147  mya (Peters et al. 2017) and 92–107 mya (Branstetter et  al. 2017). Which sphecid wasps are most closely related to bees is uncertain, with mor­ phological hypotheses suggesting that bees most

853

Insect Biodiversity: Science and Society

C S eou ac

m e Co liss lle ina m op eli tina e a Ca sipha ttina e up e e Dip olic inae hag ani ni lo Dis so ssin Para glottin i colle i te Sten otritid s ae Halicti ni Thrinchost omini Sphecodini

125

ro

Ne

Ca

llo

Xe

tid ae

te La ceous ta re

Early C re t

S

100

75

50

25

0 Myo

Hes Sam perapin i Da bini Me sypo M gan dain M acro omi i e Eu litt pid inae he ini ini rb st iin i

i in ni en e lissi dr na i e An xae nom i O la sin * No lliop inae ini Ca nurg alict Pa nanth Co apini Pen lictini Xera itini Roph nae Nomii ae Nomioidin Augochlorini Caenohalictini

C

ol le

i i lin rin diel r a ol sen Ne wn ni o T asti Bi

cene Mio ene goc Oli cene Eo ene leoc Pa

Al M lod an ap Xy ue in Te loco liini i tr p Me aped ini lipo ia Bo nini Eug mbini loss in Ap i Centr ini idin Megach i ilini Noteriades Pseudoheriades Afroheriades Osmiini* ni Anthidii es d ia r Ochre xyini Dio iini m idos inae p s A rg ae u h t Li itin ae ph rho eliin ae a r d Fi ssin ae Pa lo rin e ryg te ina Eu crap lae S Hy

ni re d An

Me litti dae

An

Rhathymini es Coelioxoid olus Parepe rini opho Anth cerini Eu i ylin Anc rini* pho ini Em spid ini ota lops lis pi n Ta oma osce rini Ex cyl lect ini n op ati en er C Ct

Ericrocidini Osiris Isepeolin i Epeolo Prote ides Cae peolini n Me oproso pidi Am lectini ni Br mob a a c Ep hy tini A eol nom ad He mm ini ini N xe oba om p t o e i d o ad i in lini ni i

Apidae

Megachilidae

854

e da

ae Halictid

Figure 28.2  Simplified version of time‐calibrated phylogeny presented by Cardinal et al. (2018), with terminals pruned to tribe or subfamily level. A non‐monophyletic taxon is indicated by “*”. The dashed branch leading to the Andrenidae indicates that the placement of the Andrenidae, relative to the other families, is uncertain. Stars indicate the three core increases in diversification detected in bees. Abbreviations: mya, million years ago; S, Stenotritidae.

recently shared a common ancestor with the Crabronidae (Lomholdt 1982, Alexander 1992, Prentice 1998, Melo 1999), whereas more recent molecular analyses have recovered a para­ phyletic Crabronidae and support the crabronid subfamilies Pemphredoninae + Philanthinae as  being sister to bees (Debevec et  al. 2012,

Branstetter et  al. 2017, Peters et  al. 2017). Regardless of the exact sister group of bees, the evidence suggests that bees evolved from some kind of apoid wasp predator of other insects that they provided to their offspring. All bees, except for a few stingless bees (Camargo and Roubik 1991, Bänziger et al. 2009), collect plant

28  Bee Diversity Through Time

material to provision their offspring, making bees the most successful wasp group that turned vegetarian. The masarines, a lineage of vespid wasps, as well as the sphecid wasp Krombeinic­tus nordenae Leclercq (Krombein and Norden 1997), also provision their offspring with pollen and nectar. Given the strong dependence of bees on flowering plants and their coevolutionary rela­ tionships, especially with eudicots, it is not sur­ prising that crown group bees are estimated to have originated when tricolpate pollen (pollen with three furrows or pores characteristic of eudicots) first appears in the fossil record, which is often equated with the origin of eudicots (Magallon 2010), or their rise in abundance and geographical expansion (Smith et  al. 2010). Furthermore, bees are thought to have played an important role in the period of increased diversification of flowering plants during the late Cretaceous (Doyle 2012) when the crown group of all but one of our seven extant bee families are estimated to have originated: Melittidae  97 (79–125) mya, Andrenidae 67 (54–85)  mya, Halictidae 94 (82–106)  mya, Stenotritidae 20 (7–37)  mya, Colletidae 70 (60–81) mya, Megachilidae 103 (92–115) mya, and Apidae 104 (95–113) mya (Table 28.1). Relationships among bee families have been investigated in numerous studies, reviewed by Danforth et  al. (2013), and some controversy remains. Some morphological analyses support a topology with the Colletidae or Colletidae + Stenotritidae being sister to all other bees (Alexander and Michener 1995), in large part based on the wasp‐like bilobed tongue of colletid bees. Most recent molecular analyses have placed the Melittidae as sister to all the remaining bees (Hedtke et al. 2013, Branstetter et  al. 2017, Cardinal et  al. 2018, Peters et  al. 2017). Monophyly of the long‐tongued bees (Apidae + Megachilidae) is well supported, as is  Stenotritidae + Colletidae as sister to the Halictidae. Placement of the Andrenidae, how­ ever, is more uncertain but is likely to be sister to Halictidae + (Stenotritidae + Colletidae), although in the phylogeny using a relaxed

molecular clock of Cardinal et  al. (2018), the  Andrenidae are sister to the Melittidae (Fig. 28.2). Bees have been hypothesized to have origi­ nated in the Southern Hemisphere (Hedtke et al. 2013) and, more specifically, in the xeric interior of Gondwana (Michener 1979, 2007). Gondwanan fragmentation has been suggested to have influenced early bee evolution (Litman et  al. 2011, Hedtke et  al. 2013). The earliest diverging family, the Melittidae, is estimated to have had an ancestral range in Africa (Hedtke et al. 2013). Today, the Melittidae are most spe­ cies rich in southern Africa, and modest num­ bers are found, predominantly, in the warm xeric areas of other parts of the world (Michener 2007). Melittids are unknown in tropical America and Australia. Ancestral geographic range reconstructions have supported a South American ancestral range for the Colletidae, suggesting an ancient vicariance between them  and their sister Australian family, the Stenotritidae, followed by numerous inter­ changes between the two continents via Antarctica (Almeida et  al. 2012, Hedtke et  al. 2013). The ancestral ranges of the other fami­ lies remain unclear, but there is weak support for a New World ancestral distribution for both the Andrenidae and Halictidae (Hedtke et  al. 2013). The Megachilidae phylogeny of (Litman et al. 2011) supports a hypothesis of an ancient vicariance event between South America and Africa at the base of the Megachilidae, based on the Fideliinae being reconstructed as paraphyl­ etic. However, paraphyly of the Fideliinae is still uncertain; Cardinal et  al. (2018) recovered a monophyletic Fideliinae. Some families of bees have been much more evolutionarily successful than others. The Melittidae and Stenotritidae have only 201 and 21 species, respectively, whereas all other fami­ lies have thousands of species (Andrenidae 2974 spp., Halictidae 4427 spp., Colletidae 2632 spp., Megachilidae 4112 spp., and Apidae 5847 spp.) (Table 28.1). However, some families are older than others and, therefore, might have had more time to accumulate more species or for species

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Table 28.1  Numbers of species and mean and 95% highest posterior density interval (HPD) of crown ages and stem ages of bee lineages. No. of species in clade

Mean crown age

HPD crown age

Mean stem age

HPD stem age

Anthophila

20,214

125

127–156

140

137–174

 Melittidae

201

97

79–125

105

90–119

  Dasypodainae

94

61

43–82

97

79–125

   Hesperapini

46

40

24–58

61

43–82

   Dasypodaini

37

48

30–68

61

43–82

   Sambini

11

na

na

48

30–68

Clade

  Meganomiinae

10

na

na

78

63–98

  Melittinae

97

64

55–75

78

63–98

   Macropidini

18

12

10–14

65

55–75

   Melittini

79

42

35–49

65

55–75

 Andrenidae

2,974

67

54–85

105

90–119

  Andreninae

1,552

44

37–51

67

54–85

   Euherbstiini    Andrenini   Oxaeinae   Panurginae    Nolanomelissini

3

25

13–38

44

37–51

1,549

36

28–44

44

37–51

22

na

na

52

45–60

1,400

46

40–58

53

45–60

1

na

na

46

40–58

133

21

13–31

42

36–48

1,266

35

30–41

42

36–48

   Melitturgini*

44

—

—

—

—

   Panurgini*

137

—

—

—

—

   Neffapini

1

na

na

22

14–30

   Protandrenini

418

12

7–17

28

23–33

   Perditini

664

12

7–18

24

19–29

   Calliopsini    Melitturgini + Panurgini + Neffapini + Protandrenini + Perditini

   Protomeliturgini  Halictidae

2

—

—

—

—

4,427

94

82–106

109

99–120

  Rophitinae

261

64

48–80

94

82–106

   Conanthalictini

13

n/a

n/a

55

39–71

   Xeralictini

7

10

3–21

58

43–75

   Penapini

6

32

17–49

55

39–71

   Rophitini

235

41

28–56

58

43–75

 Nomioidinae

95

27

13–44

78

68–90

 Nomiinae

618

39

25–55

85

74–98

 Halictinae

3,453

65

58–73

78

68–90

28  Bee Diversity Through Time

Table 28.1  (Continued) No. of species in clade

Mean crown age

HPD crown age

Mean stem age

HPD stem age

   Augochlorini

608

44

34–55

65

58–73

   Caenohalictini

171

27

23–31

57

53–62

   Sphecodini

341

24

12–39

53

48–58

Clade

   Thrinchostomini    Halictini  Stenotritidae

42

19

9–31

48

41–55

2,291

37

31–44

48

41–55

21

20

7–37

85

71–101

2,632

70

60–81

85

71–101

  Diphaglossinae

130

48

37–62

61

47–73

   Dissoglottini

14

15

6–27

48

37–62

   Diphaglossini

5

17

9–28

38

27–49

   Caupolicanini

111

22

15–31

38

27–49

8

na

na

61

47–73

 Colletidae

?     Paracolletes   Callomelittinae

11

na

na

54

42–66

  Colletinae

517

23

13–34

54

42–66

  Xeromelissinae

137

36

27–47

56

47–65

  Hylaeinae

962

43

33–52

53

44–61

  Scrapterinae

59

20

12–29

47

37–57

  Euryglossinae

393

40

29–51

47

37–57

  Neopasiphaeinae

415

57

46–68

65

56–76

 Megachilidae

4,112

103

92–115

116

105–127

  Fideliinae

18

93

71–111

103

92–115

  Lithurginae

62

31

17–48

69

57–81

  Pararhophitinae

3

na

na

69

57–81

4,029

—

—

—

—

   Aspidosmiini

2

na

na

64

42–82

   Dioxyini

36

23

11–39

64

42–82

10

na

na

20

8–39

5

na

na

20

8–39

   Megachilini

2,014

25

14–41

62

60–66

   Osmiini*

1,073

—

—

—

—

16

na

na

62

60–66

1,074

61

50–71

75

68–81

  Megachilinae*

?     Pseudoheriades ?     Afroheriades

    Noteriades     Osmiini − Noteriades

(Continued)

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Insect Biodiversity: Science and Society

Table 28.1  (Continued)

Clade

No. of species in clade

Mean crown age

HPD crown age

Mean stem age

HPD stem age

2

na

na

70

60–80

?     Ochreriades    Anthidiini

887

52

43–63

70

60–80

 Apidae

5,847

104

95–113

116

105–127

   Nomadinae + Melectini

1,436

78

67–89

84

74–94

  Nomadinae*

1,241

—

—

—

—

   Caenoprosopidini

2

25

13–39

51

43–61

   Ammobatini

127

43

30–58

71

61–82

   Nomadini

726

20

9–34

46

32–59

   Ammobatoidini

33

42

28–58

57

46–68

   Hexepeolini

1

na

na

46

32–59

   Neolarrini

16

na

na

40

28–53

   Townsendiellini

4

na

na

35

23–48

   Biastini

13

15

7–26

35

23–48

   Brachynomadini

26

33

22–45

56

46–68

   Epeolini

293

40

29–51

56

46–68

3,612

—

—

—

—

   Anthophorini

  Apinae*

754

54

42–69

94

85–104

   Melectini

195

32

23–41

51

43–61

   Osirini*

51

—

—

—

—

    Parepeolus

5

na

na

60

40–76

    Osiris

32

na

na

62

50–74

    Epeoloides

2

10

4–19

48

35–60

   Tetrapediini*

28

—

—

—

—

    Coelioxoides

3

4

2–9

60

40–76

    Tetrapedia

25

10

5–17

49

41–59

   Rhathymini

19

26

12–40

61

49–72

   Ericrocidini

42

34

25–44

61

49–72

   Isepeolini

21

29

19–42

55

43–67

   Protepeolini

5

20

10–32

48

35–60

   Centridini

257

80

61–94

91

82–100

8

22

18–28

40

35–47

   Euglossini

246

26

23–30

40

35–47

   Bombini

263

19

10–32

75

70–82

   Meliponini

503

55

52–61

75

70–82

   Ctenoplectrini

19

26

14–42

89

79–99

   Apini

28  Bee Diversity Through Time

Table 28.1  (Continued) No. of species in clade

Mean crown age

HPD crown age

Mean stem age

HPD stem age

   Exomalopsini

155

47

30–63

72

59–85

   Emphorini*

117

—

—

—

—

    Ancyloscelis

20

27

10–47

72

59–85

    Emphorini − Ancyloscelis

97

37

26–49

58

47–70

   Tapinotaspidini

139

43

30–55

58

47–70

Clade

   Ancylini

16

4

1–9

57

48–67

   Eucerini

774

42

35–50

57

48–67

  Xolocopinae + Tetrapedia

1,019

83

68–95

94

85–104

  Xylocopinae*

994

—

—

—

—

   Xylocopini

372

29

21–36

49

41–59

   Manueliini

3

na

na

73

61–86

   Allodapini

253

41

32–50

56

49–62

   Ceratinini

366

37

28–45

56

49–62

Species numbers are derived from the Discover Life bee species guide and world checklist (Hymenoptera: Apoidea: Anthophila) (Ascher and Pickering 2016). The taxonomy follows that of Ascher and Pickering (2016), except here I recognize the clades Caenohalictini, Conanthalictini, Melitturgini, Nomioidinae, Perditini, Sphecodini, Thrinchostomini, and Xeralictini, following Danforth et al. (2013). Within the Megachilinae, I follow Danforth et al. (2013), and, following Michener (2007), I do not recognize the Teratognathini. I adjusted the numbers of Ascher and Pickering (2016) to reflect these changes. Clade ages were obtained from Cardinal et al. (2018). Clades that were not monophyletic in the time‐ calibrated phylogeny are indicated by “*”. These clades accordingly have “—” for clade ages. Some clades do not have an estimate for mean crown age because they were represented by a single exemplar in the phylogeny and are here indicated by “na”. When the tribe or subfamily of the subsequent genus is uncertain, “?” is used.

to have gone extinct. To see if some lineages have more species than expected given their age, each bee lineage above the genus level (except when not monophyletic in the phylog­ eny) was plotted (Fig. 28.3) according to its stem‐clade age, as estimated by Cardinal et  al. (2018) against its species numbers (Table 28.1), following Ascher and Pickering (2016). The 95% confidence intervals of the expected species richness of a clade through time, given no extinction (e = 0) and a high extinction rate (e = 0.9), were also plotted. The 95% confi­ dence  intervals were determined using the R package Geiger, (Harmon et al. 2008), which implements the Magallon and Sanderson (2000) method to calculate net diversification rate for a clade,  given extant diversity and age. For the

calculations, the background net diversification rate of 0.0899 was used. This rate was calculated using BAMMtools (Rabosky et al. 2014) and the BAMM output files from Cardinal et al. (2018). The only clade more species rich than expected is the Perditini (Andrenidae: Panur­ ginae) which diverged from its sister clade 24  mya and today contains 664 described species. The Perditini are a North American tribe and most of the species are oligolectic. However, as a group, they collect pollen and nectar from  diverse plant taxa such as the Liliaceae, Asteraceae, and Fabaceae (Michener 2007). The tribes Protandrenini, Andrenini, and Halictini fell just within the 95% confidence interval, assuming a high extinction rate, but were well above the expected number of species

859

Insect Biodiversity: Science and Society Clade with more species than expected

18

Perditini Clades with fewer species than expected . Neffapini . Nolanomelissini/Hexepeolini . Epeoloides . Caenoprosopidini Parepeolus Coelioxoides Aspidosmiini Ochreriades Pararhophitinae Manueliini Meganomiinae Stenotritidae Fideliinae Ctenoplectrini Dasypodainae Melittidae

16

14

Ln(no. of species in clade)

860

12

10

Clades with no. of species within 95% CIs

8

6

4

2

95% CI, e = 0.09

.

0 0

20

95% CI, e = 0.0

.. . 40

60

80

100

120

140

Stem clade age (millions of years)

Figure 28.3  Relationship between stem‐clade age and species richness as natural logarithm (Ln) in bees. The 95% confidence intervals (CIs) of the expected species richness of a clade through time, given no extinction (e = 0) and a high extinction rate (e = 0.09), were calculated using the Magallon and Sanderson (2000) method and the background net diversification rate for bees of 0.0899. The stem‐clade age and number of species in Table 28.1 are plotted. When the number of species in a lineage fell within the 95% CIs of the expected species richness, the lineage was plotted with a dot. When the number of species fell outside the CIs, the lineage was given a unique symbol and named in the inset.

when assuming no extinction. The only families with fewer species than expected were the Melittidae and Stenotritidae. Despite being 97 million years old (myo), the Melittidae presently have only 201 described species, many of which are rare and narrowly restricted geographically. The Melittidae are mostly distributed in the Old World and most of the species are oligolectic (Michez et  al. 2008). The Stenotritidae, esti­ mated to be 20 myo with 21 described species,

are restricted to Australia. The Melittidae sub­ families Meganomiinae and Dasypodainae, and the Megachilidae subfamilies Pararhophitinae and Fideliinae also contain fewer species than expected. The Fideliinae are sister to all other megachilid lineages and have a disjunct distri­ bution with a lineage in xeric areas of Africa (Fidelia) and South America (Neofidelia). All species might be oligolectic, as are the three species of the Pararhophitinae, which were once

28  Bee Diversity Through Time

considered part of the Fideliinae (Michener 2007). The Pararhophitinae are found from Morocco and Egypt to Kazakhstan, and south to  northwestern India (Michener 2007). All of the 11 other lineages with fewer species than expected are either kleptoparasitic or oli­ golectic, when known. In a study of diversifica­ tion rates in long‐tongued bees, the evolution of kleptoparasitism was associated with decreased rates of diversification in eight of 10 kleptopara­ sitic clades (Litman et  al. 2013). Many of the species‐poor lineages also tend to have rather restricted distributions (e.g., Manueliini is found only in southern Chile and Argentina, and Aspidosmiini are restricted to Namibia and South Africa), and all are from the families Andrenidae, Megachilidae, and Apidae (no colletid or halictid lineage was either species rich or species poor). Figure 28.3 gives the impression that the older a clade, the more species it contains. However, this can be misleading, as not all points on this graph are independent because some lineages are nested within others (e.g., Dasypodainae is a subfamily nested within Melittidae). Therefore, to test for a positive correlation between clade age and species richness, a phylogenetic generalized least‐ squares regression (PGLS) was performed using the time‐calibrated phylogeny of Cardinal et al. (2018), pruned so that all bee species could be assigned to a terminal taxon. The PGLS fit by maximum likelihood, using a Brownian model, found a positive relationship between clade age and number of species, with a slope of 1.590, but this correlation was not significant (p = 0.3250). Therefore, clade age alone does not account for the variation in lineage species richness. Clade age is also not a good predictor of species richness across the eukaryotic tree of life (Rabosky et al. 2012). Within angiosperms, a negative relationship was found  between clade age and diversity (Tank et al. 2015). If clade age is not the predominant factor in explaining the disparity in species richness among clades of bees, shifts in diversification rates might instead be a leading factor. To

reconstruct diversification rates across their bee phylogeny, Cardinal et  al. (2018) used a Bayesian Analysis of Macroevolutionary Mixtures (Rabosky 2014, Rabosky et  al. 2014) approach and found 18 significant rate shifts. The exact placement of these shifts was impos­ sible to summarize, as the 95% credible set of rate‐shift configurations sampled 29,498 dis­ tinct shift configurations. However, three core shifts were identified: one increase within the Anthidiini (a  tribe of large black, yellow, and sometimes red megachiline bees), one increase along the branch leading to the common ances­ tor of Xylocopa (large carpenter bees), and one increase within the Meliponini (stingless bees). All three increases are within long‐tongued bees and happened 28 to 55 mya (Fig. 28.2). Along the branch leading to the common ancestor of Xylocopa (Apidae: Xylocopinae), diversification increased from the background rate of 0.0899 to 0.1473. Xylocopa diverged from its sister lineage (Tetrapedia) 49 mya, and the 375 extant species shared a common ances­ tor  29 mya, indicating that the rate increase happened sometime between 29 and 49 mya. Xylocopa includes large‐bodied bees distributed worldwide, and most species burrow into hard plant material to make their nests. They are hypothesized to have originated in the Oriental– Palearctic Region and their present world distri­ bution most likely resulted from independent dispersal events (Leys et  al. 2002). A lineage through time (LTT) analysis of the Xylocopinae showed early diversification, followed by a long period of low diversification, followed by rapid diversification in at least some of the consti­ tuent four tribes of the Xylocopinae (Rehan et  al. 2013). The authors argued that this LTT plot pattern was most consistent with a massive extinction event close to the Cretaceous– Tertiary (K–T) boundary because they esti­ mated cladogenesis for the Xylocopinae to have started about 100 mya, which is much earlier than the chronograms of Cardinal et al. (2018) and Branstetter et  al. (2017) would indicate. Massive extinction events in angiosperms should have affected the bees that depend on

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them and there is evidence for extensive floral extinction at the K–T boundary (Johnson 1992) and for impacts of this event on plant‐insect interactions (Labandeira et al. 2002). However, the diversification rate analysis of Cardinal et al. (2018) did not detect any core diversifica­ tion shifts older than 49 myo. More detailed studies on diversification rates within lineages that originated before this K–T boundary event might uncover more evidence of the effect of this event on bee diversification. The core increase in diversification rate to 0.1672 along the branch leading to the common ancestor of New World stingless bees is esti­ mated to have happened 28 to 55 mya after the  New World lineage diverged from the Old World lineage. Rasmussen and Cameron (2010) also hypothesized that major diversifications happened approximately 30–40 mya in the New  World stingless bees, but they hypothe­ sized the split between Old World and New World stingless bees to have happened approx­ imately 71 mya. Stingless bees (Apidae: Apinae: Meliponini) are highly eusocial (Michener 1974), having perennial colonies of workers, division of labor with a queen and workers, and morphological differences between queens and workers. They have a pantropical distribu­ tion and their historical biogeography is exten­ sively discussed by Rasmussen and Cameron (2010). The third core shift in diversification rate occurred 36–43  mya, and was within the Anthidiini (Megachilidae: Megachilinae) where the rate increased to 0.1544. The tribe has a worldwide distribution and includes both klep­ toparasitic and nest‐making species (Michener 2007). They nest in preexisting burrows or cavities (e.g., wasp mud nests or empty snail shells) or simply attach their brood cells to rocks, stems, or leaves (except for a few burrow‐ digging taxa). Their cells are made with foreign materials including, resin, pebbles, soil, leaves, and plant hairs. An earlier diversification rate analysis of the Megachilidae recovered two rate  shifts, the largest being at the base of the higher megachilids, which was a few million

years after the advent of nest construction using foreign  materials in this group (Litman et  al. 2011). A subsequent diversification rate analysis of long‐tongued bees (Apidae and Megachilidae), detected two significant increases in diversifica­ tion: one at the base of Stelis (Megachilidae: Megachilinae: Anthidiini) and the other at the base of the Nomadinae (Apidae) (Litman et al. 2013). Stelis is one of the earlier diverging gen­ era in the anthidiine clade found to have a higher diversification rate in the analysis by Cardinal et al. (2018), indicating some consistency across different studies regarding an increase in the Anthidiini. The Nomadinae are a clade entirely of kleptoparasitic bees. Litman et  al. (2013) hypothesized that this increase in diversifica­ tion was due to the way that nomadines para­ sitize their hosts. Instead of finding a host cell that has already been fully provisioned with a pollen ball, a host egg laid, and the cell closed off, nomadines parasitize cells that are actively being provisioned and, therefore, still open. This transition to parasitizing open nest cells is hypothesized to have expanded the range of potential hosts, consequently opening new eco­ logical niches that allowed them to increase their diversification rate. All three core increases in diversification hap­ pened along branches whose mean ages are 39–41.5 myo. This would most likely place the three increases during the Eocene which, fol­ lowing the Early Eocene Climatic Optimum 53–50 mya, consisted of a cooling period, except for the brief 400–500 thousand‐year exception known as the Middle Eocene Climatic Optimum that occurred approximately 41 mya when sea surface temperatures increased due to increased CO2 (Galazzo et al. 2014) (Fig. 28.4). From the end of the Cretaceous through the Middle Eocene, warm‐temperate to tropical vegetation extended from equatorial regions to as far north as the Beringian and North Atlantic Land bridges (Graham 1999). However, the overall temperature decline and drying trend docu­ mented during the Eocene led to the displace­ ment of the tropical and subtropical forests to more seasonally dry and arid habitats to a point

28  Bee Diversity Through Time 16

Increases in bee diversification Ar

idi

Temperature (°C)

12

fic

ati

on

wi

Early Eocene Climatic Optimum

8

th

de

cre

as

ing

tem

pe

Middle Eocene Climatic Optimum

4

ra

tur

e

0 Paleocene 60

Oligocene

Eocene 50

40

30 Age (Ma)

Miocene 20

Plio. Plt. 10

0

Figure 28.4  Global Cenozoic climate change based on deep‐sea oxygen isotope ratio (δ18O) records. The curve is a running mean of approximate deep‐sea temperature with 500 ky resolution calculated from δ18O (from Hansen et al. (2008)). The gray shaded area indicates the most likely time period when all three core increases in bee diversification happened (mean ages of branches along which increases were detected ranged from 39 to 41.5 million years ago). All three increases occurred during a period of aridification. Plio., Pliocene; Plt., Pleistocene.

where woodland savanna might have become the predominant biome in North America (Graham 1999). In the Palearctic, woody plant groups ancestral to the modern Mediterranean dendroflora started to appear during the Middle Eocene (Palamarev 1989). Given that today a higher diversity of bees occurs in more xeric and Mediterranean environments than in tropical areas, it is perhaps not surprising that all three core increases in diversification occurred when tropical forests were receding. Likewise, a study of diversification of colletid bees hypothe­ sized increased diversification 25–30 mya dur­ ing a period of increasing aridification in the Southern Hemisphere where these bees are most diverse (Almeida et  al. 2012). In North America, the Middle Eocene was a transition period in the modernization of floras from ancient Cretaceous and Paleocene predecessors to several modern plant assemblages (Graham 1999). The Eocene is also when two primary shifts in diversification in angiosperms were found: one in the most recent common ancestor of the Capparaceae and Brassicaceae 35 mya, and the second in the Asteraceae 49 mya (Tank et al. 2015).

28.5 ­Conclusions Despite the evolutionary success of bees, some species today are experiencing sharp declines. The declines in wild bees have been docu­ mented in China (Williams et al. 2008), Europe (Biesmeijer et  al. 2006, Goulson and Lye 2008), and the Americas (Cameron et al. 2011, Bartomeus et al. 2013, Kerr et al. 2015). These declines might be due to a combination of factors including parasites, pathogens, pesti­ cides, and habitat losses that have led to reduced floral resources and nesting sites (Goulson et  al. 2015). Wild bee declines threaten the production of crops that depend on them for pollination (e.g., alfalfa, apple, blueberry, canola, cherry, cranberry, cucum­ ber, eggplant, peach, pear, raspberry, squash, strawberry, and tomato. In the United States, 39% of the pollinator‐dependent crop area has a mismatch between the supply of  wild bees and the pollination demand of cultivated areas (Koh et al. 2016). Furthermore, climate change may further increase this mismatch, as bum­ blebees have been found not to shift their ranges poleward or up in elevation in response

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to warmer climate, but instead to move the southern end of their ranges northward, result­ ing in smaller distributional ranges (Kerr et al. 2015). However, with the listing of a few bee species as endangered, thus drawing attention to them in various countries, and with the increase in research on factors negatively affecting bees, solutions might be forthcoming so that we can avoid a pollinator crisis.

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molecular data on our understanding of bee phylogeny and evolution. Annual Review of Entomology 58: 57–78. Debevec, A. H., S. Cardinal and B. N. Danforth. 2012. Identifying the sister group to the bees: a molecular phylogeny of Aculeata with an emphasis on the superfamily Apoidea. Zoologica Scripta 41: 527–535. Doyle, J. A. 2012. Molecular and fossil evidence on the origin of angiosperms. Annual Review of Earth and Planetary Sciences 40: 301–326. Galazzo, F. B., E. Thomas, M. Pagani, C. Warren, V. Luciani and L. Giusberti. 2014. The middle Eocene climatic optimum (MECO): a multiproxy record of paleoceanographic changes in the southeast Atlantic (ODP Site 1263, Walvis Ridge). Paleoceanography 29: 1143–1161. Gauthier, D. A., A. Lafon, T. P. Toombs, J. Hoth and E. Wiken. 2003. Grasslands: Toward a North American Conservation Strategy. Canadian Plains Research Center, University of Regina, Saskatchewan, and Commission of Environmental Cooperation, Montreal, Quebec. 99 pp. Goulson, D. and G. C. Lye. 2008. Decline and conservation of bumble bees. Annual Review of Entomology 53: 191–208. Goulson, D., E. Nicholls, C. Botias and E. L. Rotheray. 2015. Bee declines driven by combined stress from parasites, pesticides, and lack of flowers. Science 347: 1–17. Graham, A. 1999. Late Cretaceous and Cenozoic History of North American Vegetation. Oxford University Press, New York, New York. 370 pp. Hansen, J., M. Sato, P. Kharecha, D. Beerling, R. Berner, V. Masson‐Delmotte, M. Pagani, M. Raymo, D. L. Royer and J. C. Zachos. 2008. Target atmospheric CO2: where should humanity aim? Open Atmospheric Science Journal 2: 217–231. Harmon, L. J., J. T. Weir, C. D. Brock, R. E. Glor and W. Challenger. 2008. GEIGER: investigating evolutionary ratiations. Bioinformatics 24: 129–131.

Hedtke, S., S. Patiny and B. N. Danforth. 2013. The bee tree of life: A supermatrix approach to apoid phylogeny and biogeography. BMC Evolutionary Biology 13: 138. Johnson, K. R. 1992. Leaf‐fossil evidence for extensive floral extinction at the Cretaceous‐ Tertiary boundary, North Dakota. Cretaceous Research 13: 91–117. Kerr, J. T., A. Pindar, P. Galpern, L. Packer, S. G. Potts, S. M. Roberts, P. Rasmont, O. Schweiger, S. Colla, L. L. Richardson, D. L. Wagner, L. F. Gall, D. S. Sikes and A. Pantoja. 2015. Climate change impacts on bumblebees converge across continents. Science 349: 177–180. Koh, I., E. V. Lonsdorf, N. M. Williams, C. Brittain, R. Isaacs, J. Gibbs, and T. H. Ricketts. 2016. Modeling the status, trends, and impacts of wild bee abundance in the United States. Proceedings of the National Academy of Sciences USA 113: 140–145. Krombein, K. V. and B. B. Norden. 1997. Nesting behavior of Krombeinictus nordenae Leclercq, a sphecid wasp with vegetarian larvae (Hymenoptera, Sphecidae, Crabroninae). Proceedings of the Entomological Society of Washington 99: 42–49. Labandeira, C. C., K. R. Johnson and P. Wilf. 2002. Impact of the terminal Cretaceous event on plant‐insect associations. Proceedings of the National Academy of Sciences USA 99: 2061–2066. Leys, R., S. J. B. Cooper and M. P. Schwarz. 2002. Molecular phylogeny and historical biogeography of the large carpenter bees, genus Xylocopa (Hymenoptera: Apidae). Biological Journal of the Linnean Society 77: 249–266. Litman, J. R., B. N. Danforth, C. D. Eardley and C. J. Praz. 2011. Why do leafcutter bees cut leaves? New insights into the early evolution of bees. Proceedings of the Royal Society B 278: 3593–3600. Litman, J. R., C. J. Praz, B. N. Danforth, T. L. Griswold and S. Cardinal. 2013. Origins, evolution and diversification of cleptoparasitic lineages in long‐tongued bees. Evolution 67: 2982–2998.

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Lomholdt, O. 1982. On the origin of the bees (Hymenoptera: Apidae, Sphecidae). Entomologica Scandinavica 13: 185–190. Magallon, S. 2010. Using fossils to break long branches in molecular dating: a comparison of relaxed clocks applied to the origin of angiosperms. Systematic Biology 59: 384–399. Magallon, S. and M. J. Sanderson. 2000. Absolute diversification rates in angiosperm clades. Evolution 55: 1762–1780. Melo, G. A. R. 1999. Phylogenetic relationships and classification of the major lineages of Apoidea (Hymenoptera), with emphasis on the crabronid wasps. Scientific Papers, University of Kansas Natural History Museum 14: 1–55. Michener, C. D. 1974. The Social Behavior of the Bees: A Comparative Study. Harvard University Press, Cambridge, Massachusetts. 418 pp. Michener, C. D. 1979. Biogeography of the bees. Annals of the Missouri Botanical Garden 66: 277–347. Michener, C. D. 2007. The Bees of the World. Second edition. Johns Hopkins University Press, Baltimore, Maryland. 992 pp. Michez, D., S. Patiny, P. Rasmont, K. Timmermann and N. J. Vereecken. 2008. Phylogeny and host‐plant evolution in Melittidae s.l. (Hymenoptera: Apoidea). Apidologie 39: 146–162. Müller, A. 1996. Host‐plant specialization in Western Palearctic anthidiine bees (Hymenoptera: Apoidea: Megachilidae). Ecological Monographs 66: 235–257. Palamarev, E. 1989. Paleobotanical evidences of the Tertiary history and origin of the Mediterranean sclerophyll dendroflora. Plant Systematics and Evolution 162: 93–107. Peters, R. S., L. Krogmann, C. Mayer, A. Donath, S. Gunkel, K. Meusemann, A. Kozlov, L. Podsiadlowski, M. Petersen, R. Lanfear, P. A. Diez, J. Heraty, K. M. Kjer, S. Klopfstein, R. Meier, C. Polidori, T. Schmitt, S. Liu, X. Zhou, T. Wappler, J. Rust, B. Misof and

O. Niehuis. 2017. Evolutionary history of the Hymenoptera. Current Biology 27: 1–6. Prentice, M. A. 1998. The comparative morphology and phylogeny of apoid wasps (Hymenoptera: Apoidea) (2 volumes). Ph.D. Dissertation. University of California, Berkeley, California. xi + 1,439 pp. Rabosky, D. L. 2014. Automatic detection of key innovations, rate shifts, and diversity‐ dependence on phylogenetic trees. PLoS ONE 9: e89543. Rabosky, D. L., M. Grundler, C. Anderson, P. Title, J. J. Shi, J. W. Brown, H. Huang and J. G. Larson. 2014. BAMMtools: an R package for the analysis of evolutionary dynamics on phylogenetic trees. Methods in Ecology and Evolution 5: 701–707. Rabosky, D. L., G. J. Slater and M. E. Alfaro. 2012. Clade age and species richness are decoupled across the eukaryotic tree of life. PLoS Biology 10: e1001381. Rasmussen, C. and S. Cameron. 2010. Global stingless bee phylogeny supports ancient divergence, vicariance, and long distance dispersal. Biological Journal of the Linnean Society 99: 206–232. Rehan, S. M., R. Leys and M. P. Schwarz. 2013. First evidence for a massive extinction event affecting bees close to the K–T boundary. PLoS ONE 8: e76683. Robertson, C. 1925. Heterotropic bees. Ecology 6: 412–436. Sheffield, C. S., S. D. Frier and S. Dumesh. 2014. The bees (Hymenoptera: Apoidea, Apiformes) of the prairies ecozone with comparisons to other grasslands of Canada. Pp. 427–467. In D. J. Giberson and H. A. Carcamo (eds). Arthropods of Canadian Grasslands (Volume 4): Biodiversity and Systematics, Part 2. Biological Survey of Canada, Ottawa, Ontario, Canada. Smith, A. S., J. M. Beaulieu and M. J. Donoghue. 2010. An uncorrelated relaxed‐clock analysis suggests an earlier origin for flowering plants. Proceedings of the National Academy of Sciences USA 107: 5897–5902.

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Tank, D. C., J. M. Eastman, M. W. Pennell, P. S. Soltis, D. E. Soltis, C. E. Hinchliff, J. W. Brown, E. B. Sessa and L. J. Harmon. 2015. Nested radiations and the pulse of angiosperm diversification: Increased diversification rates often follow whole

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29 Insect Biodiversity in Culture and Art Gene Kritsky and Jessee J. Smith Department of Biology, Mount St. Joseph University, Cincinnati, Ohio 45233, USA

Insects have played a pivotal role in our evolu­ tion, and it could be argued that they have also  influenced the evolution of the human experience, which is reflected in our cultures. Throughout our evolutionary existence, we have been bitten by insects, and we have fed upon them in turn. Long before we were aware of insects, they were part of our lives. Molecular studies of lice show that they coevolved with our hominid ancestors. Thirteen million years ago, before the gorillas split from the common ancestor of humans and chimpanzees, ancestral Pediculus lice lived on our common ancestor. When the gorillas started on their own evo­ lutionary path 11 million years ago, their lice began their own evolutionary history. This was repeated when Ardipithicus marked the split of the hominin branch 5.8 million years ago, and as the Australopithecines and early Homo contin­ ued our evolution, so did the lice that evolved to live on us (Reed et al. 2007). These ectoparasites helped to cement the social interactions of early humans. Chimpan­ zees are often observed grooming each other to remove lice, a social ritual that fosters bonding among members of the group. Human moth­ ers have been observed doing the same to their young children, which was recorded in a seventeenth‐­century painting by Jan Siberechts,

Cour de Ferme (Weiss 2009). The association of humans with lice might have contributed to the loss of our dense covering of fur 1.2 million years ago as natural selection favored reducing our ectoparasitic load (Jablonski 2010). This reduction of suitable “habitat” contributed to the subsequent evolution of clothing, which in turn influenced the evolution of lice again. Clothing lice evolved from head lice, and the molecular record suggests that this divergence took place between 170,000 and 83,000 years ago (Toups et al. 2011, Kittler et al. 2003). The evolution of lice documents that insects were with us before we were aware of our own humanity, and they remained as we developed material culture. Thus, it is not surprising that lice and other insects should influence other aspects of our cultural evolution as well. The examination of insect biodiversity in cul­ ture is as broad as the definition of culture. Culture includes beliefs, symbols, values, meth­ ods of expression, and the accumulation and improvement of material objects by groups of people through time. In the end, these beliefs and objects help to define our cultures, and can even symbolize some cultures to others. From a historical perspective, the study of insect biodiversity in human culture reveals patterns that show why certain insects were

Insect Biodiversity: Science and Society, Volume 1, Second Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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important to the members of various cultures. Some insects have influenced our world from the earliest times, whereas other insects reveal a  geographic bias. In the end, this survey of insects in culture reflects our evolving interest in the practical, the meaningful, and the beauti­ ful insects that shaped our cultural world.

29.1 ­Prehistory Human culture in prehistory arose from utili­ tarian needs. Early stone tools that were used to butcher meat do not provide any evidence of an interest in insects, but the sort of mind that could fashion a hand‐axe from a piece of flint and conceive of spectacular paintings of animals on cave walls would not have overlooked the details of the insects that shared the world with early humans. The oldest representations of insects, dating from the Upper Paleolithic, were  created by people from the Late Magdalenian culture, possibly as early as 14,000 years ago. At the rock shelter Enléne in France, a fragment of bone was found, which was incised with a strikingly clear representation of a camel cricket (Trogophilus sp.) (Bégouen and Bégouen 1928). These crick­ ets were probably common inhabitants of the rock shelter (Guthrie 2005). Their jumping behavior would certainly have been noticed, and they might have been a source of food. This early association between humans and camel crickets continues today, as they are frequent residents in basements and cellars. The most common insect to be found in pre­ historic art was probably the warble fly or bot fly. Representations of the larvae of these flies date from between 14,000 and 11,000 years before present and were carved in the round or engraved on bone (Guthrie 2005). A detailed jet carving of a warble fly larva prominently dis­ played in the Vienna Museum of Natural History includes lines that reflect the insect’s characteristic segments and represents its tapered shape. These insects lay their eggs on reindeer and caribou, where they hatch into

larvae that burrow under the skin of their host inside a liquid‐filled shell. Animals infested with bot flies might have been a prized food, just as (in more recent times) the Inuit consid­ ered the larvae a delicacy. References to other prehistoric insect carv­ ings are based on equivocal identifications. “Butterfly” pendants carved from mammoth ivory are now more widely believed to be repre­ sentations of human breasts (Cook 2013), and “centipede” engravings found in caves are so geometric in style that a firm identification is impossible. A prominent cliff painting found in Bicorp in eastern Spain may date from 5,000 years ago and is among the oldest illustrations of humans inter­ acting with insects. This well‐known illustration shows a honey hunter suspended by ropes and carrying a bag. A natural hole in the rock wall on which it is painted is incorporated in the scene to represent a wild colony of bees. Flying around the honey hunter are several figures believed to represent honeybees (Kritsky 2010). Beeswax residues on pottery shards dating from between 9,000 to 8,000 years before present are the oldest direct evidence of humans interacting with hon­ eybees (Roffet‐Salque et al. 2015). These prehistoric carvings, engravings, and paintings share a common theme: they all involve insects that were living in  association with humans. The camel cricket was a com­ mon sight in their shelters and might have been consumed by the people dwelling there, and entomophagy might also have motivated the representations of warble fly larvae. Honey has been a prized food for millions of years; chimpanzees not only tear into honeybee nests, but also fashion special tools to use as they rob the bees. These tools are so valued that the chimpanzees carry them around rather than simply discarding them when they have finished robbing the nest (Boesch et al. 2009). It is not unreasonable to assume that our ancestors might have done the same. This util­ itarian recognition of insects in prehistory is the prelude to their increasing importance in the ancient world.

29  Insect Biodiversity in Culture and Art

29.2 ­Insects in the Ancient World Ancient history brings to mind great monu­ ments: the pyramids of Egypt, the Parthenon of Greece, the Colosseum of Rome, and the Great Wall of China. Increasingly complex material culture, along with the development of writing, architecture, and complex mythologies, brought with it a more complex interest in insects. This  ancient entomology included honeybees, acknowledging their continued importance, but also involved representations of other insects as religious symbols. Honeybees were among the first insects that caught the attention of the Egyptian pyra­ mid builders. Beekeeping was a state‐controlled occupation, as documented in King Neuserre Any’s Sun Temple, built in the century after the construction of the Great Pyramid and the Sphinx. The honeybee commonly appeared as a hieroglyph that represented the Nile’s delta region in the royal titulary, which was carved in association with the cartouches of the pharaohs (Kritsky 2015). Several beetles attained important symbolic status in ancient Egyptian culture. Click bee­ tles (family Elateridae) were associated with the goddess Neith, who dates back to the beginning of Egyptian civilization. These bee­ tles were represented in offering bowls, and the shape of their elytra was incorporated into stylistic representations of Neith. The bupres­ tid beetle Steraspis squamosa was associated with the Osiris mythology. According to the myth, Osiris was lured into a special chest by his brother Seth, who sealed Osiris inside the chest and dumped it into the Nile. It floated into the Mediterranean, where it eventually came ashore at Byblos in the branches of a tamarisk tree. The tree grew to envelop the coffin, with Osiris inside. The king of Byblos, after seeing the magnificent tree, ordered it to be cut down and taken to his palace as a col­ umn. Osiris’ wife, Isis, searched for her hus­ band and found him at Byblos. After healing the king’s son, who was critically ill, Isis was offered a gift, and she asked for the pillar. She

instructed carpenters to split the tree open to reveal the chest, releasing Osiris. She then bound the tree together and returned it to the king. This bound tree became the djed pillar (Kritsky 1991), an important and ubiquitous symbol for the Egyptians. The Egyptians believed that the actions of animals, including insects, reflected their myth­ ological importance. Buprestid beetles bored into trees, and when the trees were split and these insects found inside, the Egyptians saw this as a symbol of Isis splitting the tamarisk to release Osiris. The Egyptians also carved amu­ lets in the form of tenebrionid beetles, which are commonly found wandering on the sand around the pyramids and tombs. The symbolic importance of these beetles probably related to their feigning death when touched, drawing the limbs close to the body in a way that resembles a bound mummy. Then, after a few moments, the beetle resumes its activities, seeming to come back to life. Given the Egyptian fixation with death and the afterlife, this behavior would have made the tenebrionid beetle a symbol of resurrection (Kritsky 1991). Other insects played an important role in the Book of the Dead, the lengthy set of spells and instructions intended to guide the deceased into the afterlife. There are references to praying mantids using their raptorial forelegs to carry the body of the deceased to the Weighing of the Heart, where his or her good and evil deeds were measured. This might explain the discov­ ery of a mantid mummy, which was wrapped in linen, carefully preserved, and placed in a small sarcophagus (Kritsky 1993). Carved amulets in the form of flies have been found on necklaces placed on mummies. These amulets were, in many instances, highly pol­ ished and made of carnelian, faience, gold, sil­ ver, or wood. Their importance is related to the mummification process, which could take up to 70 days. This process provided ample time for flies to lay eggs on the body of the deceased, complete their development, and emerge from the mummy before it was properly prepared. These flies might have been associated with the

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ba or the personality of the dead individual, generally represented as a human‐headed bird. By placing these amulets on the body, the Egyptians were symbolically returning to the body what the flies took away. The symbolic association of the dead with calliphorid blow­ flies continues to this day in Egypt, where folk wisdom holds that blowflies that enter a house should not be harmed because they might con­ tain the spirit of someone who once lived there (Kritsky 1985). Flies also formed the basis of a military award: large golden flies were awarded to soldiers who demonstrated tenacity in battle. The most spec­ tacular of these are the three golden flies that were awarded to Ahhotep II, which were given to her by her sons Khamos and Ahmose for her help in driving out the Hyksos, foreign occupi­ ers of Egypt. The flies were based on the vicious and common biting flies that would return to bite again in spite of the victim’s attempts to slap them away. Soldiers who continued to attack despite the enemy’s efforts to repel them mir­ rored this behavior and warranted this award (Kritsky 1993). Grasshoppers or locusts figured prominently in Egyptian jewelry and hieroglyphs. The locust as a hieroglyph represented a number of sol­ diers. Small amulets in the form of grasshoppers have been found on necklaces interred with mummies. Locusts are often depicted in tomb reliefs of marshes and gardens, where they are shown resting on plants, being eaten by pre­ dators, or being caught by a boy (Kritsky 1993, Hansen 2001). Butterflies also figured in scenes of the after­ life. Numerous scenes show detailed paintings of butterflies flitting around the idyllic land­ scapes. Some of these were so detailed that it is possible to determine the species that was illus­ trated. Butterflies were represented in jewelry. The ritualistic importance of these butterflies is not well understood. The simplest explanation is that, as in many cultures, they were symbols of beauty or possibly a reference to the afterlife (Kritsky 1993).

Probably the best‐known example of an insect in Egyptian art is the scarab beetle, Scarabaeus sacer L. Scarabs, or dung beetles, are often found rolling a small ball of dung on the hot sands of Egypt. This action was thought to be symbolic of the sun moving across the sky, a process that the Egyptians attributed to the beetle god, Kephre. In Egyptian mythology, the sun was swallowed at sunset by the goddess Nut, who would give birth to it at sunrise. The  sun was then pushed across the sky by Kephre until it was again consumed at dusk (Kritsky 1993). By the 11th Dynasty, scarabs were commonly used as seals that were worn on rings. These scarabs were carved on the underside with names, titles, or decorative symbols. When an amphora or some other container was tied shut, the knot would be encased with clay that was stamped with the seal of the owner or inspec­ tor. Scarabs made of semiprecious stones were incorporated in some of the spectacular gold pectorals found in the tomb of Tutankhamen. The pharaoh Amenhotep III issued commemo­ rative scarabs that were carved of steatite, with the underside carefully inscribed with details of important events of his reign (Kritsky 1993). Heart scarabs, generally the size of the palm of a hand, were inscribed on the underside with spells from the Book of the Dead and interred with mummified bodies. After the 15th Dynasty, these heart scarabs were not inscribed, but had  holes where outstretched wings could be attached. Eventually, the scarabs were carved more realistically, with six legs and clearly depicted elytra. Although they were still placed on mummies, these were smaller than other scarabs and functioned as an amulet represent­ ing new life. The grandest of all of the scarabs were the colossal granite scarabs found at Karnak Temple at ancient Thebes (now Luxor) and at Abu Simbel in southern Egypt. These large scarabs were more than a meter in diameter and were the largest insect sculptures of the ancient world (Kritsky 1993).

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Several insects now regarded as pests lived with the ancient Egyptians, but we do not know their ancient names. The Egyptians shared their environment with bed bugs, fleas, lice, mos­ quitoes, and a number of stored grain pests (Panagiotakopulu et al. 2010). Although none of these common nuisances have been found in ancient Egyptian art or writing, that does not mean that the Egyptians viewed insects only in positive ways. A spell in the Book of the Dead is intended to protect the deceased from a feared, monstrous water beetle, and papyri show a man standing in a boat, aiming a spear at a beetle in the water. In this case (and for unknown rea­ sons), the water beetle was considered an evil animal of the river, as dangerous as crocodiles, hippopotami, and snakes (Taylor 2010). Interest in insects extended beyond Egypt in the ancient world. Insects also appear in the culture of ancient Israel, as documented in the Old and New Testaments. Biblical verses men­ tion bees, flies, and grasshoppers, as well as insects that were not part of the Egyptian lan­ guage, including ants, fleas, grubs, maggots, mole crickets, and moths. Deuteronomy 8:15 might be among the oldest references to mole crickets: “All of your trees and the fruit of the ground will be infested with the mole cricket” (Kritsky 1998). Locusts make up a large proportion of the insects mentioned in the Bible. Many of the ref­ erences deal with locusts as plagues or pests. However, Mark 1:6 describes John the Baptist eating these insects: “John was dressed in a rough coat of camel’s hair, with a leather belt round his waist, and he fed on locusts and wild honey.” John’s consumption of locusts was meant to make him seem more of an ascetic, but locusts graced many tables throughout the region (Kritsky and Cherry 2000). In the British Museum is a bas‐relief from Sennacherib’s pal­ ace in Nineveh (now called Kouyunjik, in north­ ern Iraq) that shows two men carrying skewers of grasshoppers in each hand. These locust kebabs dating from between 704 to 681 bce reveal that entomophagy was fit for a king.

During the time of Solomon, the people of  Israel were keeping bees in large apiaries associated with settlements. Bees were impor­ tant  not only for honey, much of which was exported to Egypt, but also for beeswax. Beeswax was a critical component in the lost wax‐casting process used to produce cast ves­ sels and other metal objects. The beehives at Tel Rehov might have been associated with an ambitious metalsmithing industry (Mazar and Panitz‐Cohen 2008). The Epic of Gilgamesh, a Sumerian story about the king of Uruk, is generally considered the oldest known example of great literature. It first existed as five poems that were merged into a single narrative dating from ancient Babylon, around 1800 bce. Notably, one passage used insects as a metaphor to illustrate the brevity of human life. The insect mentioned has been interpreted as a dragonfly, but other transla­ tions suggest that it was actually a mayfly. The doubt involves the meaning of an ancient Sumerian term that directly translates as “water locust” (Parrella 2013). The passage has been translated as, “There is no permanence. Do we build a house to stand forever, do we seal a con­ tract to hold for all time? Do brothers divide an inheritance to keep forever, does the flood‐time of rivers endure? It is only the nymph of the water locust who sheds her larva and sees the sun in his glory.” Flies were carved on some Babylonian and Assyrian cylinder seals. On one Babylonian cyl­ inder seal, a large fly was carved next to the god Nergal. The association with a god helped to deify the fly, and the Epic of Gilgamesh describes the gods gathering “like flies.” Another passage reads, “The gods of strong‐walled Uruk are changed into flies and buzz around the streets.” The mother goddess, Belit‐ili or Aruru, wore a necklace of lapis lazuli fly heads (Greenberg and Kunich 2005). Many of the flies on cylinder seals are carefully carved with a head, thorax, and abdomen, whereas the flies made of gold were more abstract. Like the fly amulets of Egypt, these are thought to depict blowflies.

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The ancient Greeks had a different view of flies; they considered them both pests and objects of curiosity. Apollo protected herds and functioned as myiagros or “fly‐chaser,” which was an important duty because flies were thought to bring plague. However, the Greeks were the first to make the transition from using flies as symbols to studying flies as biological organisms. Aristotle studied their reproduc­ tion  and development, making detailed obser­ vations  on their immature stages (Davies and Kathirithamby 1986). Thus, the ancient Greeks  are widely considered the first ento­ mologists, and much of that reputation comes from similar observations by Aristotle and oth­ ers (Bodson 1983). Insects that captured the Greek imagination went beyond the few that interested the Egyptians. The Greeks wrote jokes about fleas and carved ants, bees, butterflies, flies, and grasshoppers into gemstone intaglios. Indeed, one intaglio shows the caterpillar, pupa, and adult butterfly, documenting that the Greeks had witnessed this progression through the stages of the holometabolous development of butterflies (Davies and Kathirithamby 1986). The Greek word psyche means both “but­ terfly” and “soul” or “conscious self.” The god Hermes was also linked to the soul and the phallus. Heraclitus believed the soul was liquid and these beliefs might be behind the Greek images that show a flute‐playing man with semen dripping onto a butterfly (Davies and Kathiri­thamby 1986). The Greeks were skilled beekeepers who kept their bees in cylindrical horizontal hives made of Earthenware. Aristotle wrote more about bees than about any other insect, and his detailed observations suggest that he must have studied the interior of a hive (Davies and Kathirithamby 1986). The ancient Greeks’ inter­ est in bees might not be surprising, but they also represented wasps in painted motifs and sculp­ ture. Some of the wasp intaglios are so detailed that they can be identified to the family Vespidae and to a genus. In the Iliad, Homer wrote that

wasps constructed their nests along roads, which attracted the attention of children, who would disturb the wasps to the extent that they would attack individuals walking near the nests (Davies and Kathirithamby 1986). The ordered way of life and industrious nature  of ants attracted attention in Greek art and literature. Aristotle, observing these social insects, considered ants to be “political.” The Myrmidons, fierce soldiers described in the Iliad, were said to have originated from ants that Zeus transformed into men. In another fable, Zeus does just the opposite: he punishes men who stole their neighbors’ fruit by turning them into ants. Ants were also thought to have the ability to divine the future, and even to predict the weather (Davies and Kathirithamby 1986). Cicadas were popular with ancient Greeks. Plato, in his Phaedrus dialogue, discussed love as cicadas sang in the background. According to Greek mythology, cicadas were once men who were so enamored by the music that the Muses played that they sang without ceasing until they perished. The Muses were moved to transform the men into cicadas and endowed them with the ability to live without food or drink. This myth was probably linked to the popular belief that cicadas subsist only on dew (Davies and Kathirithamby 1986).

29.3 ­The Cult of Artemis: A Case Study Bees were so important during the time of Greek influence that several cities, including Ephesus, featured bees on their coinage as a symbol of the city (Ransome 1986, Stoneman 2013). Artemis, the patron goddess of Ephesus, was worshipped as a mother‐goddess from as early as 400 bce, and her cult lasted until 300 CE. The “Greater Statue” (as it is known) from Ephesus shows her standing upright and wear­ ing a tall crown and a gown covered with a range of animals, including honeybees (Fig. 29.1). The “Beautiful Statue” shows the goddess wearing a

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much shorter headdress and a pectoral com­ posed of the signs of the zodiac. Both of these statues date back to the second century of the Common Era and were found in the ruins of the ancient city (Rogers 2012). Entomological interest in Artemis is sparked by the honeybees that adorn the sides of these statues. Strangest of the ornamentations that festoon Artemis are the rows of egg‐shaped or ovoid objects across her upper torso, whose

identity has provoked considerable debate. They have been thought to be breasts, hilltops, a type of leather bag favored by the Hittites, bulls’ testicles, honeybee “cocoons,” or figs (Rogers 2012, Stoneman 2013). The common theme of these suggestions is their shared sym­ bolism of fertility. More certain is the connec­ tion of Artemis with bees and honey. The priestesses of Artemis were called melissae, which translates to “bee,” and other names such as “beekeeper” (Elderkin 1939). A more recent suggestion is that the insects on the statues and on the Ephesian coins were not honeybees at all, but fig wasps. Support for this notion requires the identification of the ovoid objects as figs (Eisenberg 2009). This view, however, has not received wide support because the fig wasp (only about 1–2 mm) would have been difficult to observe in an age before magnification, and even more difficult to represent on a statue. The fig wasp hypothesis also ignores the several titles that link Artemis with bees, honey, and beekeeping. Some statues of Artemis pair the bee motif with an open flower, casting more doubt on the fig wasp sug­ gestion, as fig trees do not produce highly visi­ ble, open flowers. We might never know for certain what the ovoid objects were, but it is clear that bees played a central role in this complex cult. Although our modern minds are inculcated with values and associations that differ from those of the creators of these objects, the example of the symbols of Artemis underscores the  value of entomologists working alongside archaeologists, mythologists, and classicists to help identify the insects on these ancient works and suggest interpretations based on the insects’ biology.

29.4 ­Roman Insect Art Figure 29.1  A cast of the Greater Statue of Artemis, showing the ovoid objects and the bees on the sides of the dress, on display at the Ephesos Museum in Vienna. Photograph by Gene Kritsky.

Ancient Rome’s cultural insect biodiversity was heavily influenced by the Greeks, and we find intaglios of ants, butterflies, and grasshoppers

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from Rome. Intaglios carved with scorpionflies constitute a new addition to the diversity of insect motifs. In the British Museum’s collec­ tion are two rings (British Museum numbers 1812,0704.1453 and 1812,0704.1451) from Imperial Rome (first to third centuries CE), each of which is set with an intaglio carved from sard (a brownish‐red variety of chalcedony) and engraved with a stylized scorpionfly. On one intaglio, the scorpionfly is holding a bow and drawing an arrow. The images cannot be attrib­ uted to any particular species, as the insects look like flies with scorpion pincers and their abdomens end in long tails; in short, these crea­ tures resemble winged scorpions rather than realistically depicted mecopterans. The concept of these “scorpion‐flies” might have originated with the writings of the ancient Greeks; the Roman naturalist Pliny the Elder claimed that the Greek author Apollodorus stated that there were scorpions with wings. Actual mecopterans were not described in literature until the 17th century (Carpenter 1931). The Romans also used insects as a form of decoration and created fibulae in the form of flies and cicadas (British Museum number 1772,0309.86). These devices, now familiar as  shawl pins or brooches, were a decora­ tive  way to fasten clothing. The Romans also incor­p orated grasshoppers as a decoration on pot­ tery oil lamps (British Museum number AN1129978001). The cultural insect diversity of the world increased over three millennia. The inclusion of insects of economic importance, such as honey­ bees and locusts, is not surprising. However, tenebrionid beetles in Egyptian culture, flies in Assyria, and the representation of butterfly metamorphosis in ancient Greece document an interest in insects as mythological symbols. Gems inscribed with insects, or carved in their shape, show that people of the ancient world considered insects interesting and beautiful enough to use for personal adornment, and sug­ gest that they had special symbolic meaning to the wearers.

29.5 ­Ancient China Examples of insect diversity in the ancient world were not limited to Western civilizations, as insects also featured largely in the art of ancient China. Possibly as early as the Shang Dynasty (1500–1050 bce), craftsmen were carving intri­ cate jade cicadas (British Museum number 1945,1017.14). The practice continued through the Western Zhou Dynasty and into the Han Dynasty (202 bce –220 ce). Some of these cica­ das were pierced and used as toggles or counter­ balancing weights for small pouches that were worn at the waist; the ornamental toggle would secure the cord attached to the pouch to the belt of the wearer. Cicadas were considered symbols of rebirth in China, based on the observation that the nymphs emerge from the ground and transform into winged adults (Riegel 1981). Following this belief, cicada amulets carved from jade were placed on the tongue of the deceased at burial. The ancient Chinese first carved the picto­ graph for the honeybee around 1000 bce, as recorded on an inscribed bone, and the word “honey” first appeared in 300 bce. The Chinese developed beekeeping by 200 ce, using Apis cerana rather than Apis mellifera, which occurred further west. The honeybee figured prominently in poetry; the earliest examples are found in the Hymns of Zhou, which appeared sometime before 200 bce. The poet Guo Puin wrote about the daily duties of bees in his poem composed during the Jin Dynasty (265–420 ce). The poem describes honeybees foraging in the wilderness, the transformation of nectar into honey, the activities of the queen, and swarming (Lau 2013). Cicadas also figure prominently in Chinese folklore as a Buddhist symbol. Lafcadio Hearn noted that according to Buddhism, the adult cicada, which is the result of the molt of the last nymphal instar into the adult stage, is symbolic of how “reincarnation obscures the memory of the previous [life]; we remember our former existence no more than the [cicada] remembers

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the shell from which it has emerged.” The empty  cicada exuviae are a symbol of “Earthly pomp  –  the hollow show of human greatness” (Riegel 1981).

29.6 ­Religions of India The complex religious traditions of India extend back to the ancient world. Hinduism, consid­ ered by some to be the oldest surviving religion, includes several insect references, the honey­ bee  being the most common. The Hindu god of  love, Kama, uses a bow made of sugarcane equipped with a bowstring made of bees. The mother goddess, Bhramari Devi, is the goddess of the black bees. Honey is used in Hindu rituals as a food offering to bring rain and to aid fertil­ ity (Cherry and Sandhu 2013). Ants are consid­ ered the “first‐born” of the world, which is represented by an anthill (Hogue 1987). Ants also decapitated the god Vishnu. Other insects that figure into Hinduism include a beetle, the cochineal bug, and the silkworm. Silk is still worn on the wrists to ward off the evil eye. Other major religions in India include Bud­ dhism, Jainism, and, more recently, Sikhism. Buddhism holds that if an individual’s life has been filled with bad deeds, that person will be reincarnated as a lower form of life, such as an insect. Jainism includes six “great vows,” one of which is to do no harm to other living creatures, including insects. Jains will not drink water dur­ ing the night to avoid accidentally swallowing an insect, and they wear masks over the mouth and nose to prevent inhaling insects. Finally, Sikhs believe that God is present in all life, including insects (Cherry and Sandhu 2013).

29.7 ­Post‐Classical Era The post‐classical era, also Europe’s Middle Ages, includes the period from 500 to 1500 ce, after the closure of the last Egyptian temple and the fall of the Roman Empire. Records are sparse

during this period, but new sources are coming to light as museums and libraries digitize their medieval illuminated manuscripts – early books that are highly decorated with lavish hand‐ drawn and painted illustrations. The oldest such manuscripts, dating back to between 400 and 600 ce, incorporate illustrations of insects and entomological subjects. These works were probably influenced by Physiologus, a book written during the early Christian period (between 200 and 400 ce) by an unknown author. Physiologus included many references to insects and their symbolism. Flies were thought to represent sorrow and the devil; ants represented work; bees represented virtue and wisdom; scarabs represented sinners; grasshop­ pers represented Christ; and moths symbolized physical temptations (Morge 1973). Many of these insects were commonly used as symbols during the ancient world and their importance continued during the Middle Ages; some of these insect symbols endured for more than a millennium. For example, towards the end of the post‐classical era, Carlo Crivelli’s vivid 1480 painting Madonna and Child (Fig. 29.2) shows a fly next to the Christ child, symbolizing sin and  evil as well as the brevity of Christ’s life (Zeri 1973). Many of these illustrated manuscripts were created by monks, so it is not surprising that they should include biblical themes. Among the oldest are the Exultet Rolls, which date from 1000 ce and celebrate the contribution of the bees in providing the beeswax that was used to make the Easter or Paschal candle. Tradition required that the Paschal candle be made of pure beeswax, and chants and eulogies to the bees were recited at Easter. The ending of one such eulogy read “O truly happy and marve­ lous  bee…who is productive and yet chaste” (Freeman 1945). The straw skep beehive appears on many illustrated manuscripts as a symbol of the ordered society of the Catholic Church (Wilson 2007). Insect biodiversity represented on these medieval manuscripts increased after 1000 ce.

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Figure 29.2  Carlo Crivelli’s 1480 painting, Madonna and Child. Note the fly at the lower left.

In addition to the butterflies, cicadas, and grass­ hoppers, all holdovers from the ancient world, several new insects make their appearance. In the Hours of Joanna I, one border includes moths as well as flies (Biggs 2014). The 14th‐ century treatise Seven Vices features a praying mantis in the corner of one page and rhinoceros beetles and stink bugs (including the nymphal and adult stages) on other pages. This might be one of the earliest illustrations of a heteropter­ ous insect (Cocharelli 1330–1340). Bestiaries – books that describe various ani­ mals and often provide a morality lesson with each animal  –  became popular in the Middle

Ages, and ants on anthills appear in several bes­ tiaries. Some describe the ants’ behavior: walk­ ing in orderly lines and storing up food, working in groups, and carrying items that clearly exceed their own weight. The illustrations in bestiaries were not always totally accurate, and some were wildly fantastical. Six‐legged and eight‐legged spiders in spider webs were illustrated in medieval bestiaries. The 12th‐century Gerald of Wales’ Topographic Hiberniae even has a ten‐ legged spider. Other insects illustrated in the margins of medieval books include biting flies, butterflies, caterpillars, dragonflies, and grass­ hoppers (Biggs 2014).

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Antlions (family Myrmeleontidae) first appear during the sixth to seventh centuries. These lar­ val neuropterans, known for living in sand or dusty soil and making conical pits to trap ants, were noted insects in medieval times, recurring in illustrations from the 8th, 12th, and 13th cen­ turies. In Guillaume le Clerc’s 13th‐century bes­ tiary, he writes, “There is still another ant, Not of those which I have told you, Which has the name ant‐lion. Of the ants this is the lion, It is  the smallest of all, The boldest and wisest. Other  ants it hates bitterly; In the dust quite deftly, It hides; so clever it is. When the others come laden, It jumps out of the dust upon them, It attacks and kills them” (Druce 1923). It is clear from this manuscript that the life cycle and winged adult stage of the antlion were not well understood at the time. The transition from simply using insects as a metaphor to actively collecting and working with insects can be observed in some illu­ minated manuscripts. Apicultural illustrations show a swarm of bees being captured with a bag, as well as detailing how the comb was cut out of a beehive. A 13th‐century French parch­ ment shows a woman in the margin carrying an  insect net, presumably to catch butterflies, and a 14th‐century French marginal illustration shows an ape aiming at a butterfly with a bow and a blunt arrow. During the Middle Ages, apes or monkeys were thought to be symbolic of mischief (Nazari 2014). At the same time as European monks were creating illuminated books, a new religion was beginning in the Middle East. In 570 or 571 ce, Muhammad was born in Mecca to an influential tribe in what is now known as Saudi Arabia. Muhammad was illiterate, so it is believed that he dictated the Qur’an to his believers between 610 and 632 ce (el‐Mallakh and el‐Mallakh 1994). The Qur’an does not contain as many insect references as are found in the Bible, but it does refer to insects that are found in the Bible. The chapters in the Qur’an are called Sûrahs, and Sûrah II includes a reference to a gnat or mosquito: “Lo, Allah disdaineth not to coin the similitude even of a gnat.” The quotation can

be interpreted to mean that Allah’s lessons may be gleaned from observing even the smallest and lowest of creatures. Locusts are discussed in Sûrah VII, which relates the story of the plagues that God set upon Egypt during the time of Moses. Verse 133 reads, “So we sent upon them the flood and locusts and vermin and frogs and blood as dis­ tinct signs, but they were arrogant and were a criminal people.” Some authorities have inter­ preted the word “vermin” as “lice” to bring the passage more in line with the biblical account. Generally, the Arabic word could mean any ectoparasitic insect. The Qur’an’s passage on bees is rather inter­ esting in what it implies. The section is found in Sûrah XVI, verses 68 and 69: “And the Lord inspired the bee, saying: Choose thou habita­ tions in the hills and in the trees and in that which they thatch; Then eat of all fruits and follow the ways of thy Lord, made smooth [for thee]. There cometh forth from their bellies a  drink diverse of hues. Wherein is healing for mankind. Lo! herein is indeed a portent for  people who reflect” (el‐Mallakh and el‐ Mallakh 1994). Muhammad enjoyed honey, and in Sûrah XLVII, verse 15, paradise is described as flow­ ing with rivers of purified honey. The passage in Sûrah XVI indicates that Muhammad knew that bees made honey from material they gath­ ered from trees, and that honey possessed medicinal value. Some writers have gone so far as to suggest the phrase “in that which they thatch” shows that Arabs at the time were keeping bees in hives (el‐Mallakh and el‐ Mallakh 1994). Sûrah XXVII is titled “The Ants,” as they are the focus of verses 18 and 19: “Till, when they reached the Valley of the Ants, an ant exclaimed; O ants! Enter your dwellings lest Solomon and  his armies crush you, unperceiving. And [Solomon] smiled, laughing at her speech, and said: My Lord, arouse me to be thankful for Thy favor wherewith Thou hast favored me and my parents, and to do good that shall be pleasing unto Thee, and include me in [the number of ]

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Thy righteous slaves.” These passages have two interpretations. One suggests that there was a valley of the ants and that Solomon’s conversa­ tion with them was a miracle. Others suggest that the valley in question was inhabited by humans who belonged to the tribe known as “the ants.” Sûrah XXII, verse 73, mentions flies: “Indeed, those you invoke besides Allah will never create [as much as] a fly, even if they gathered together for that purpose. And if the fly should steal away from them a [tiny] thing, they could not recover it from him. Weak are the pursuer and pursued.” Sûrah XXIX, verse 41, uses a spider to further show the failings of other deities: “The likeness of those who choose other patrons than Allah is as the spider when she taketh unto herself a house, and lo! the frailest of all houses is the spi­ der’s house, if they but knew” (el‐Mallakh and el‐Mallakh 1994).

29.8 ­The Americas On the opposite side of the world, other cultures were also weaving insects into their mytho­ logies. Of the many cultures that arose in Meso­ america, the Maya, Olmecs, Toltecs, and Aztecs left behind the best‐known material legacies, including sprawling temple complexes, intricate stone carvings, and works of gemstones and precious metals. Honeybees figured promi­ nently in the mythology of the Maya, who believed that “bacabs” were at the four corners of the Earth, where they held up the sky. These “bacabs” were gods of bees and the apiary (Ciaramella 2002). Beekeeping developed independently in Cen­tral America using the stingless bee, and honey and wax were prized commodities. The bees were kept in hollow logs that were placed on a rack under a canopy or beneath the eaves of a hut to protect them from the elements (Kritsky 2010). Aspects of stingless beekeeping have also been recorded in an ancient Maya codex that shows beekeepers tending hives (Jones 2013).

Butterflies were prominent motifs of the Toltec culture, which dates from 800–1200 ce. The insects appear at Tula on stone columns carved in the shape of elaborately dressed war­ riors, whose costume includes a prominent pec­ toral ornament in the form of a stylized butterfly. Butterflies also adorn the walls of the Patio of Quetzalpapalotl at Teotihuacán. Two Central American gods are linked to butterflies. Beutelspacher (1988) suggested that Xochiquetzal, the goddess of love, flowers, fire, and plants, was represented by Papilio multi­ caudatus, the western tiger swallowtail, which is often seen in Central Mexico. The other butterfly goddess is Itzpapalotl, or Obsidian Butterfly. She was represented as a fierce deity with claws and butterfly wings studded with stone knives; Beutelspacher (1988) proposed that this imagery might have been inspired by a  saturniid moth. An altar in the National Archaeological Museum in Mexico City shows  Itzpapalotl as a skull‐headed butterfly with obsidian knives on the wings, surrounded on one side by skulls (Fig. 29.3). Butterflies can also be found on an outer ring of the famous Aztec calendar stone. The large circular stone was carved in 1479 and found in 1790, where it had been buried in what is now downtown Mexico City. The butterflies on the outer ring have been linked to the goddess Itzpapalotl, but also as a symbol of transforma­ tion, given the understanding of the butter­ fly  life cycle at the time. Butterflies were also thought to be associated with fire and transfor­ mation (Beutelspacher 1988). The Aztecs also fashioned stone carvings representing fleas, insects not often found in other cultures. Although the stone carvings are relatively small (30 to 60 cm in length), the art­ ists magnified the flea many times its natural size, and the attention to anatomical detail is remarkable (Fig. 29.4). The Aztecs also pro­ duced highly finished stone grasshoppers. Many Aztec carvings had symbolic meanings; for example, dogs were thought to help guide the deceased in the afterlife, cacti served as  boundary markers, and locusts signified

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Figure 29.3  The Aztec goddess Itzpapalotl on display at the Museo Nacional de Antropología in Mexico City. Photograph by Gene Kritsky.

swarms. However, the symbolic meaning of the flea is unknown (Morge 1973). Insects also permeated the cultures of other native peoples of North America. Petroglyphs,

or rock art symbols, are common in what is now the American Southwest, and they include sev­ eral examples of insects. Dragonflies, as drawn by the Hopi, were represented as a cross with Figure 29.4  An Aztec flea carving at the Museo Nacional de Antropología in Mexico City. Photograph by Gene Kritsky.

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two horizontal lines. Dragonflies often deco­ rated baskets and pottery, and were inscribed as petroglyphs. These aquatic insects represented water, abundance, and fertility (Patterson 1992). A popular petroglyph in the Southwest is Kokopelli, or the flute player. Kokopelli dates back to 800 ce, and was generally drawn as a  humpbacked stick figure playing a flute. Kokopelli is based on the cicada, which was common during the summer months and prized as a food source. Cicadas are sucking insects, and the proboscis that extends between their front legs could easily have been interpreted as the flute that is drawn on the glyph, while their rounded thorax gives them a humpbacked pro­ file. Adding to their association with the flute player, male cicadas make a loud call in an effort to attract the female, which is silent. The Hopi also created a glyph that repre­ sented ants. Ants hold special meaning for the native peoples of the Southwest. The figure rep­ resented the Red Ant Clan, a group of extended and interrelated families within the Hopi tribe. A myth describes how the ant people saved the  Hopi ancestors by inviting them into their subterranean homes. The ant people fed the Hopi to the extent that the ants had to tighten their  belts, which (the myth explains) is why ants have a constricted “waist” to this day (Waters 1977). The mythology of the Native Americans of eastern North America also features insect characters. One myth, recorded by James Mooney (1900), an ethnographer who lived with the Cherokee for several years, relates how a water beetle created land: The Earth is a great island floating in a sea of water, and suspended at each of the four car­ dinal points by a cord hanging down from the sky vault, which is made of solid rock. When the world grows old and worn out, the cords will break and let the Earth sink down into the ocean, and all will be water again. The Indians are afraid of this. When all was water, the animals were above in Galun’lati, beyond the arch; but it was very much

crowded, and they were wanting more room. They wondered what was below the water, and at last Dayunisi, “Beaver’s Grandchild,” the little Water‐beetle, offered to go and see if it could learn. It darted in every direction over the surface of the water, but could find no firm place to rest. Then it dived to the bottom and came up with some soft mud, which began to grow and spread on every side until it became the island which we call the Earth. It was afterward fastened to the sky with four cords, but no one remembers who did this. Other insects that are part of Cherokee mythology include the cicada (or “jar‐fly”), crickets, fireflies, katydids, mole crickets, and  scarab beetles. All have their own sto­ ries  as part of the interpretive lore through which  the  Cherokee understood their world (Mooney 1900).

29.9 ­Modern History Modern history, which began around 1500 ce, is marked by interest in scientific discovery, the  artistic and intellectual revolutions of the Renaissance, and increased exploration of the world. It was a time when knowledge led to power, and an increased knowledge of the insect world, shared through printed books, led to the increased inclusion of insects in human culture. The work of Jacob Hoefnagel (1573–1632) exemplifies the changing interest in insects. He was born in Antwerp, the son of a self‐taught painter. However, Jacob was formally trained, and he became an accomplished painter of miniatures as well as a diplomat, art dealer, and printmaker. He was associated with several royal courts, which might have given him access to insect collections that were part of  cabinets of  curiosities. He produced 302 illustrations of insects, including the first examples of mag­ nified insects. His draw­ ings  included 78  dipterans, 72 lepidopterans,

29  Insect Biodiversity in Culture and Art

37  beetles, 35 hymenopterans, 22 orthopter­ ans, 21 hemipterans, 16 neuropterans, and 14 odonates. These were organized into 16 plates and published as  his Diversae Insectarum Volatilium Icones (Fig. 29.5). Likely the most prolific of the early entomo­ logical artists was Maria Sibylla Merian (1647– 1717), who is considered by some to be the mother of entomological illustration. Her major work was Metamorphosis Insectorum Surina­ men­sium, an exquisitely illustrated book that pictured the insects of Suriname on plants, though not all of the plants were actually hosts of the insects she depicted. Merian was fasci­ nated by butterflies and their metamorphosis, and she bred and reared out many of the insects she illustrated. She produced hundreds of paint­ ings of insects; many include several insects in each illustration (Dicke 2000). Balthasar van der Ast (1593/4–1657), a con­ temporary of Maria Sibylla Merian, was a Dutch still‐life artist whose paintings include numer­ ous insects crawling or perching on flowers, fruits, and shells. His paintings illustrate several

orders, including Coleoptera, Diptera, Ephemer­ optera, Hymenoptera, Lepidoptera (caterpillars and adults), Odonata, and Orthoptera. Some are detailed enough to make species‐level iden­ tifications. The record number of insects in one painting goes to a Flemish painter, Jan van Kessel, whose 1664 painting The Continents fea­ tured more than 100 insects (Dicke 2000). Some of the most famous figures in art history also considered insects worthy of representa­ tion. Leonardo da Vinci produced gestural drawings of dragonflies and a cerambycid bee­ tle. He also sketched butterflies and flies as part of his study of flight, and ants and cicadas as part of a puzzle (Kritsky and Mader 2010). Pieter Brueghel the Elder incorporated insect themes in some of his most famous paintings, such as The Combat between Carnival and Lent and Children’s Games. Brueghel might have used his engraving The Beekeepers as a subtle criticism of the Inquisition; The Beekeepers might have represented Catholics trying to restore the Church (Kritsky and Mader 2011). One of Albrecht Dürer’s most famous works is a

Figure 29.5  A plate from Jacob Hoefnagel’s 1630 Diversae Insectarum Volatilium Icones showing several insects.

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1505 painting of a stag beetle. Dürer wrote, “It is indeed true, that art is omnipresent in nature, and the true artist is he who can bring it out” (Anonymous 2008). Several insects were used as heraldic sym­ bols. Ants, butterflies, crickets, fleas, flies, and grasshoppers found their way onto the coats of arms of important families from England, Flanders, France, Genoa, Venice, and Verona. Honeybees, which represented industry and thrift, were used in heraldic achievements for several British, Dutch, Flemish, French, German, Greek, and Spanish families. Bees adorn the escutcheon of the Barberini family of Florence, including Maffeo Barberini, who became Pope Urban VIII and was involved in the prosecution of Galileo. Bees were even engraved on the suit of  armor of the general of the Papal armies. Originally, the Barberini arms showed three horse flies on a red field (a symbol of the fami­ ly’s original name, Tafani da Barberino), but they were later changed to the more appealing

bees on a blue field to reflect the family’s climb up the social ladder (Fig. 29.6). Napoleon also used bees as a heraldic symbol, and ordered that families needed an Imperial grant if they  wished to use bees in their achievement (Velde 2002).

29.10 ­Japanese Art While insects were regarded as pests in many Western cultures, they were celebrated for their beauty in Japanese culture from its earliest ori­ gins. Insects appeared as decorative elements in  textiles and added realism to the scenes of nature often painted on hanging scrolls and folding screens. In contrast to Western art tra­ ditions, “art” and “craft” were not perceived as separate domains by Japanese artists, and utili­ tarian items were often adorned with beautiful motifs requiring highly developed technical skills. Figure 29.6  Barberini’s bees (15th century) at the Vatican. Photograph by Gene Kritsky.

29  Insect Biodiversity in Culture and Art

Almost every item of traditional garb pro­ vided an opportunity for creative adornment, and personal items that were used daily received particular attention. The sagemono (a small container that hung from the wearer’s belt and essentially functioned as a wearable pocket) was often highly decorated, but even more interest­ ing was the toggle that held it in place, the netsuke. These netsuke became a sculptural for­ mat in themselves, and were carved into myriad fanciful shapes that frequently embodied an ele­ ment of humor. Insects were common motifs for delicately carved netsuke, many of which depicted cicadas, dragonflies, mantids, wasps and their nests, and water striders. The arms and equipment of the high‐ranking warrior class known as the samurai reflected their elite status and, as such, were exquisitely wrought and highly decorated. The tsuba, or hand guards, of a samurai’s katana and waki­ zashi (long and short swords) were often graced with delicate renderings of insects. These beau­ tiful art objects, frequently seeming at odds with their militaristic context, balanced the weight of the blade and prevented the wearer’s hand from being cut during a sword thrust. Earlier tsuba were generally made from iron or steel, but examples made in later, peaceful peri­ ods could be of brass, copper, or shakudo (an alloy of gold and copper). Tsuba were made by  generations of artists who dedicated their entire  careers to their craft, and their skill is reflected in the decorative motifs that were often pierced  in openwork designs, engraved, carved, or inlaid  with gold and other metals (Tsuda 1976). Sometimes the decoration would form a miniature scene of nature, including highly accurate or stylized representations of ants, butterflies, cicadas, crickets, dragonflies, slant‐faced grasshoppers, and spiders and their webs (Fig. 29.7). Japanese woodblock prints, particularly the genre known as ukiyo‐e (“pictures of the floating world”) also reveal a fascination with insects in the context of natural beauty, as well as provid­ ing an insight into human interactions with insects. The British Museum houses several

Figure 29.7  A Japanese tsuba with a gold dragonfly in the upper right. The original is in the Walters Art Museum; photograph in the public domain.

examples that show butterflies, crickets, drag­ onflies, katydids, and wasp nests, as well as people collecting cicadas and fireflies, throwing rocks at cicadas, and hanging lamps filled with fireflies. As documented by these prints, insect collecting has long been a traditional Japanese pastime, and children often care for crickets that are kept in special cages. Even today, the rearing and care of beetles and other insects remains a popular hobby in Japan. After the period of isolation ended in 1853, Japanese art powerfully influenced the work of European artists and designers, who were capti­ vated by the restrained and elegant use of space and the balance of composition found in wood­ block prints, as well as by the exotic imagery. Japanese insect motifs were not overlooked by designers such as William Morris and René Lalique, and throughout the Arts and Crafts Movement and the rise of the Art Nouveau style, jewelry, textiles, and wallpaper featured

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insects ranging from scarabs to grasshoppers to yellowjackets, often depicted with a realism that could be as unsettling as it was attractive.

29.11 ­Language and Literature It is possible to study increasing insect diversity in Western culture through an examination of the first recorded use of words for particular insects, as documented by the Oxford English Dictionary. The oldest English words for spe­ cific insects date back to the 8th century and apply to insects that are bothersome to peo­ ple: fleas, lice, and wasps or hornets. All three insects are present in early English translations of Greek literature. Aristophanes and others wrote of fleas in a comic sense, and Aristotle, whose writings had become increasingly better known by this time, gathered detailed informa­ tion about both lice and wasps (Davis and Kathirithamby 1986). By the end of the Middle Ages, the insects that had attracted the attention of the Egyp­ tians, Greeks, Romans, and Chinese  –  ants, bees, beetles, cicadas, crickets, flies, locusts, and moths  –  had all appeared in the English language (Table 29.1). Two insects not com­ monly known to earlier periods – earwigs and stoneflies  – appeared later in the English vocabulary. The word “earwig” appeared circa 1000 in a leechdom (a collection of medical remedies) that included a treatment for ear­ wigs in the ears. The word “stonefly” was used around 1450, not surprisingly, in a book on angling. After 1500, the number and variety of insect words in the English language increased dramatically. Many of these insects had come to the  attention of collectors and naturalists; thus, the interest was similar to the illustrations by Hoefnagel, Merian, and van der Ast, who were painting the newly discovered diversity of insects at the same time. The first entomological book printed in England was Thomas Moffett’s Insectorum sive Minimorum Animalium Theatrum, or The

Theatre of Insects. The work included the unpublished writings of C. Gesner, E. Wooton, and T. Penny. Moffett finished the book in 1589, but it was not published until 1634, thirty years after his death. The book was written in Latin and is illustrated with several insects including aquatic Heteroptera (including back‐ swimmers and nepids), beetles (including aquatic beetles, blister beetles, a darkling bee­ tle, longhorned beetles, scarabs, and stag bee­ tles), butterflies (including swallowtails and several caterpillars), caddisflies, cicadas, cock­ roaches, crane flies, crickets, damselflies, drag­ onflies, grasshoppers, a mole cricket, more than 100 moths, praying mantids, and wasps. The book also described a few of the insects’ life histories. The development of the printing press around 1440 and the resulting availability of printed books probably contributed to the increased inclusion of insects in creative writing  –  the body of work that falls under the category of literature. Literature includes poems, plays, fables, novels, and stories that reflect the imagi­ nation of the author. The term “literature” can also document the collected writings of a peo­ ple, such as English literature, or it may reflect the body of work of an individual, such as the plays of Shakespeare. Humans have been creat­ ing literature for more than four thousand years, and insects and insect references are found in its earliest examples, as well as in the poems, novels, and plays of the great writers of more recent times. Beowulf, which dates back to 700 to 1000 ce, is considered the oldest example of Anglo‐Saxon literature, and it is filled with mead references. Even the name of the lead character, Beowulf, directly translates as “bee‐wolf” or “bee‐hunter” and, less directly, means “bear,” referring to the bear’s hive‐raiding behavior. This Medieval story has inspired a modern retelling that incor­ porates bees into the Beowulf legend (Nye 2004). One exciting passage reads: Beowulf halted his men when they came to the crack that led to the Firedrake’s

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Table 29.1  The earliest recorded English use of insect nouns as recorded in the Oxford English Dictionary Online (http://www.oed.com/). The year of first use precedes the English word. Year

Insect

Year

Insect

700

flea, translation of Pliny

1601

insect

725

louse

1622

bed bug

725

wasp/hornet

1624

cockroach

800

blattis or beetle

1626

dragonfly

893

gnat

1640

mayfly

950

fly (any insect)

1642

bug (general)

1000

beetle

1646

mantis

1000

earwig

1658

thrips (leafhopper)

1000

house fly

1658

firefly

1000

butterfly

1668

scorpionfly

1000

bee

1668

watermoth

1000

chafer, any destructive beetle

1686

mole cricket

1200

locust

1760

walkingstick

1325

ant

1781

termite

1325

cricket

1791

agrion

1340

spider

1817

damselfly

1382

moth

1826

brown lacewing

1400

grasshopper

1841

bark louse

1400

honeybee

1855

silverfish

1425

cicada

1863

lacewing

1450

stonefly

1884

aphid

1572

mosquito

1889

larval dobson(fly)

1579

scarab

1923

webspinner

den.  He had them set the hives down in the entrance. Then he sat for a while, muttering to the bees in each hive. No one could make out what he said. It sounded like nonsense…The others were too puz­ zled to protest. They noticed that the bees in each hive buzzed busily as Wiglaf wriggled past them. Beowulf stooped and  murmured soothingly and the noise subsided. “Call yourself a dragon?” shouted Beowulf. “You look more like a glowworm!” … Beowulf made a high‐pitched buzzing sound.

The Firedrake took a deep breath… … And swallowed a big Queen Bee that emerged from the glove as if in answer to Beowulf ’s call! “They follow the Queen Bee anywhere!” This, whispered to Wiglaf on the way up the moun­ tain, was the essence of Beowulf ’s plan. Now, in response to another noise he made, sawing at his lips with his square‐tipped fingers, all the twelve hives came alive. The bees poured out, a singing angry stream, orange, brown, black, yellow. They buzzed into the crack in the mountain.

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They whirled past Beowulf. And on into the brightness of the treasure‐chamber. The Firedrake saw them coming. Its gold eyes bulged with fright. It tried to shut its mouth, but the stake between its jaws pre­ vented this. The bees poured down the monster’s throat like a stream of honey, in pursuit of their queen. But when they reached the Firedrake’s stomach their effect was like no honey in the world. They began to sting! Hundreds of bees, stinging it from the inside! The Firedrake roared with pain and fury. It tried to spit out bees. But there were too many. Much later in the development of English literature, insects made several appearances in the plays of William Shakespeare. Beetles, but­ terflies, crickets, fleas, flies, glow‐worms, gnats, grasshoppers, grubs, honeybees, humble bees (now called bumblebees in the United States), and mayflies served to underscore scenes of love, tragedy, and mirth (Eddy 1931). Several insects are mentioned in Act 3, scene 1 of A Midsummer Night’s Dream: The honey bags steal from the humble‐ bees, And, for night‐tapers, crop their waxen thighs, And light them at the fiery glow‐worm’s eyes, To have my love to bed and to arise; And pluck the wings from painted butterflies, To fan the moonbeams from his sleep­ ing eyes. Another insect‐related passage in Act 3, scene 1 of Measure for Measure uses a lowly beetle to convey the terror and suffering felt by all living beings at the moment of death. …Darest thou die? The sense of death is most in apprehension; And the poor beetle, that we tread upon, In corporal sufferance finds a pang as great As when a giant dies.

As insect biology became better understood, later authors composed poems that included details of insect life cycles and behaviors. Sometimes these descriptions were used as a metaphor or to impart a moral lesson, whereas at other times they served a simpler, didactic purpose. For example, a poem by Thomas Barker published in 1659 concerns mayflies as a lure for fishing, and was meant to instruct and inspire fellow anglers. Mary Alcock, on the other hand, describes the activities of a hive as a fable in her poem The Hive of Bees, written in 1792. Poetry based on observation of insects was common during the late 18th century. Even Charles Darwin’s grandfather, Erasmus Darwin (1791), who was a contemporary of Alcock, wrote about insects in prose. The romantic poet Percy Bysshe Shelley, who lived two and a half centuries after Shakespeare’s time, peppered his poems with insect imagery, as did the great Scottish poet Robert Burns. The spectacular metamorphosis of a dragonfly into its adult form did not escape the notice of Alfred, Lord Tennyson, who captured the moment in his 1833 poem, “The Dragon‐fly”: Today I saw the dragon‐fly Come from the wells where he did lie. An inner impulse rent the veil Of his old husk: from head to tail Came out clear plates of sapphire mail. He dried his wings: like gauze they grew; Thro’ crofts and pastures wet with dew A living flash of light he flew. Shortly thereafter, across the Atlantic, the American poet Emily Dickinson, who drew much of her inspiration from nature, mentioned insects in 180 of her poems. Seventy‐eight of these dedicate at least one stanza to half of the poem to an insect; the other 102 involve an insect metaphor or simile (Rutledge 2003). In the realm of fiction, many novels mention insects, but others involve insects as characters or explore the world from an insect’s point of view. The best‐known insectan novella is Franz

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Kafka’s Die Verwandlung, which was published in 1915. Known to English‐speaking readers as The Metamorphosis, the book tells the story of Gregor Samsa, who wakes up one morning to find that he has metamorphosed into some kind of insect. The term that Kafka uses to describe the insect is Ungeziefer, which means “vermin.” A cleaning woman in the book refers to him as a Mistkäfer, which translates to “dung beetle,” but some readers have suggested that the term could also mean a cockroach, or any kind of beetle. This compelling novella can capture the reader in its opening paragraph (Kafka 2003): One morning, as Gregor Samsa was waking up from anxious dreams, he discovered that in bed he had been changed into a monstrous verminous bug. He lay on his armour‐hard back and saw, as he lifted his head up a little, his brown, arched abdomen divided up into rigid bow‐like sections. From this height the  blanket, just about ready to slide off completely, could hardly stay in place. His numerous legs, pitifully thin in comparison to the rest of his circumference, flickered helplessly before his eyes. The insects that occur with the greatest fre­ quency in literature are the same insects that have attracted notice throughout time: bees, bee­ tles, butterflies, crickets and grasshoppers, fleas, and flies. The mystery of insect metamorpho­ sis,  which influenced the mythology of many cultures, is also pervasive in fiction. However, increasing awareness of insects and understand­ ing of their biology has been reflected in more recent creative works. Insects appear with par­ ticular frequency in science fiction novels, often as antagonists or  as inspiration for alien races. Robert A. Heinlein’s 1959 novel Starship Troop­ ers, an enduringly popular example of the genre, pits humans against an arachnoid race known simply as “the Bugs.” Other novels delve more deeply into the intricate lives of eusocial insects. Bernard Werber’s Les Fourmis (The Ants) trilogy

involves a myrmecologist who invents a machine that converts typed words into pheromone chemicals, enabling him to communicate with colonies of intelligent and power‐hungry ants (Shelomi 2013).

29.12 ­Insects in Music Just as cicadas inspired the imagery of the flute player Kokopelli, other insects have inspired musical compositions that convey, through music, the actions of the insect. This inspira­ tion may be best heard in the 1899–1900 com­ position The Flight of the Bumblebee, written by Nikolai Rimsky‐Korsakov for his opera The Tale of Tsar Saltan. The composition accompa­ nies the opera as the tsar’s son transforms into a bumblebee to enable him to fly to his father. The music captures the zigzagging flight and hovering that bumblebees are capable of per­ forming, and this imagery led to its use as the theme for the Green Hornet radio program. It has also been adapted by Freddie Martin into a jazz piece, The Bumble Boogie. Thirty‐three years before Rimsky‐Korsakov put the flight of a bee to sheet music, Joseph Strauss composed Die Libelle (The Dragonfly) Polka Mazurka Op. 204. Although the flight of a dragonfly is silent, the graceful, undulating pro­ gression of the piece conveys the impression of its motion: now lazily gliding, then ascending or descending, suddenly darting, and every now and again alighting. Looking farther back into the his­ tory of insects as musical inspiration, the late 15th‐century composition El Grillo (“The Cricket”), a song for four voices written by Josquin Desprez, makes a lyrical observation: “The cricket is a good singer / He can sing very long / He sings all the time / But he isn’t like the other birds.” Insect diversity in music increased during the 20th century, both with songs about bugs and music groups named for insects. Of the former, the Boll Weevil Blues was a favorite of blues musicians. It is attributed to Charley Patton, who wrote it in 1908 about the boll weevil,

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Anthonomus grandis, a curculionid that feeds on cotton buds and flowers (Peterson 2007). The lyrics changed slightly as different artists (ranging from the poet Carl Sandburg to Woody Guthrie, Burl Ives, The Weavers, Pete Seeger, Jimmy Page, and most recently the Punch Brothers) recorded it, but the version recorded by Lead Belly is among the best known: (Wah‐hoo) Well the boll weevil and the little black bug Come from a‐Mexico they say Came all the way to Texas Just a‐lookin’ for a place to stay Just a‐lookin’ for a home, just a‐lookin’ for a home (Doo‐doo‐wop‐wop) Well the first time that I seen the boll weevil He was a‐sittin’ on the square Well the next time that I seen him He had his a‐family there Just a‐lookin’ for a home, just a‐lookin’ for a home (Doo‐doo‐wop‐wop) Well the farmer took the boll weevil And he put him on the red‐hot sand Well the weevil said this is a‐mighty hot But I take it like a man This will be my home, this will be my home Well the farmer took the boll weevil And he put him on a keg of ice Well the weevil said to the farmer This is mighty cool and nice This will be my home, this will be my home (Doo‐doo‐wop‐wop) Well if anybody should ask you Who it was who sang this song Say a guitar picker from a‐Oklahoma city With a pair of blue jeans on Just a‐lookin’ for a home, just a‐lookin’ for a home (Doo‐doo‐wop‐wop) (Ledbetter 2014) As the blues made its way into the main­ stream of popular music, the insects frequently

referenced in blues lyrics continued to appear in what we now know as rock and roll. Coelho (2000) analyzed insect references in popular music by surveying a music database using a variety of insect keywords. His survey found 912 tracks that referred to insects from 18 dif­ ferent orders. The frequency is noteworthy: 23.1% of the songs involve hymenopterans, 20.4% lepidopterans, and 18.4% dipterans. Within these represented orders, the hyme­ nopteran references focused on several bees, with a smaller number of wasps and ants. Common lepidopteran references related to butterflies. The dipterans represented were overwhelmingly flies, mosquitoes, and mag­ gots. Bed bugs were the most common hemip­ teran, but like the coleopterans, odonates, and orthopterans, they each accounted for only 5% or less of the insect references. The increase in diversity of insects in popular songs is also notable. While just over 61% of the songs involved the same three orders of insects that have been part of human culture for millennia, the remaining songs (more than 300 of them) encompassed 15 other orders. Generally, the insects that appear in the lyrics of popular songs fulfill the same metaphorical functions that they had in earlier poetry. Bees often connote sweetness and most frequently appear in reference to an affectionate love inter­ est, but their stinging ability is also noted. Bed bugs, flies, and fleas are often stand‐ins for trou­ blesome rivals. Ants tend to be associated with either their insignificant size or their industri­ ous nature, and butterflies are most often sym­ bols of beauty. Insects inspired names for many music groups, especially during the early days of rock and roll. Given the success of groups like Buddy Holly and the Crickets and the overwhelming popularity of the Beatles (originally the Silver Beetles), one might think that band members would choose more diverse insect names in an effort to stand out. However, Hymenoptera, Diptera, and Lepidoptera accounted for approx­ imately 50% of groups with insectan names,

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likely reflecting the limited entomological expe­ rience of the musicians. Even so, several rather unusual orders are represented among music group names: Coleoptera, Dermaptera, Ephe­ mer­ optera, Isoptera, Mantodea, Neuroptera, Odonata, Phthiraptera, Plecoptera, Siphonap­ tera, and Thysanura.

29.13 ­Insects in Cinema The invention of motion pictures in the early 1900s created an entirely new form of entertain­ ment, and it was not long before insects infested early films. The 1910 silent film The Acrobatic Fly, made by the British naturalist F. Percy Smith, was an insect debut on the big screen. Smith secured a live fly to a matchstick in such a way that the insect was lying on its back with its legs in the air. He found that the fly would ­“juggle” almost any small, lightweight object, so the film consists of brief clips of the fly manipu­ lating twigs, corks, a miniature dumbbell, and (perhaps most entertainingly) a ball with another fly walking on top (Bevir 2013). From their earliest beginnings, animated films also featured insects. Though he was better known as the creator of Gertie the Dinosaur, Winsor McCay produced a short film titled How a Mosquito Operates, in which a mosquito repeatedly attacks a sleeping man (Dirks 2016). The mosquito wears a top hat and carries a tiny briefcase, setting a precedent for the highly anthropomorphized insects that would con­ tinue to star in cartoons as the popularity of animated films grew. Walt Disney, Warner Brothers, and other animation studios produced numerous cartoons featuring insects in a variety of situations, though they were often minor characters without dialogue or even names. Yet, one of Disney’s most popular feature‐length films, Pinocchio (1940), introduced an insect as a major character: the charming Jiminy Cricket, who serves as Pinocchio’s conscience and even sings the movie’s Academy Award‐winning song, “When You Wish Upon a Star.”

Insects figured less prominently in live‐action films, but when the beginning of the nuclear age after the Second World War resulted in a boom in the popularity of science fiction movies, the  “Big Bug” genre was spawned and theater screens began to crawl with insects of mam­ moth proportions. Reflecting both the fear and fascination with which the 1950s public regarded science, the “Big Bugs” usually were created as the result of nuclear accidents, irra­ diation, or scientific curiosity gone awry. Them! (1954), the first “Big Bug” movie, is one of the best examples. It featured giant ants that arose as a result of atomic bomb testing. The “Big Bug” movies pushed the boundaries of the cinematic special effects of the time, and stop‐ motion animation, matting techniques, minia­ ture sets, and giant puppets were all brought into play as filmmakers tried (and sometimes failed) to create convincing stories of enormous arthropods (Cloyd 2013). However awkward many of the “Big Bug” films might have been, they were produced in the United States, Japan, Mexico, Finland, and Italy, documenting global interest. As was the case with Winsor McCay’s early cartoon, insects were favorite subjects for the pioneers of computer animation, and a two‐ minute 1984 short film titled The Adventures of André and Wally B. starred a stylized bee with a striped black‐and‐yellow egg‐shaped body, a round black nose, two wings, and four over­ sized, dangling feet (Berenbaum and Leskosky 2009). In the mid‐1990s, advanced computer technology made it possible to produce feature‐ length computer‐animated movies. Although computer animation had undergone vast improve­ ments from its simple beginnings, it was still best suited to rendering geometric vol­ umes and shiny surfaces – ideal for construct­ ing arthropod characters. Following the success of its first full‐length computer‐animated movie, Toy Story (1995), Pixar released A Bug’s Life in 1998. The film was well received by audi­ ences and critics for its entertaining story and beautifully rendered imagery. Like the cartoons

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of the 1930s, the insect characters were highly anthropomorphized; notably, the ant protago­ nists of the story have human‐like, expressive faces and only four limbs, while the antagonistic grasshoppers are rendered with the full comple­ ment of six limbs. Antz (1998), which was released just a few weeks before A Bug’s Life by DreamWorks Animation following a public feud between the two studios, featured more realistically detailed ant characters with six legs and elongated, prominently segmented bodies. The cynical, more adult‐oriented humor and storyline of Antz (a burned‐out worker ant voiced by Woody Allen tries to escape the daily grind by joining the soldier caste) provided more oppor­ tunities to include aspects of actual insect biol­ ogy, although (as in A Bug’s Life) the main character and many other worker and soldier ants are male. This error was repeated in Bee Movie (2007), in which Jerry Seinfeld voices a male worker bee dissatisfied with his job prospects. Whether animated or live action, insect biodi­ versity in film history reflects the same major orders that dominated popular music. Almost half of the movies incorporating insects involved Hymenoptera, with bees and ants most com­ monly represented. Flies represented the sec­ ond most common insect inspiration for movies, and nearly one in five insect movies focused on Diptera. Roaches displaced Lepidoptera as the third most common movie insect, appearing in just over 10% of movies. In many instances, the insects were intended to “creep out” the viewer, and flies and roaches appeared primarily as indicators of filth. A few movies, however, por­ tray roaches almost affectionately; the Japanese film Twilight of the Cockroaches (1987) and the 1996 American cult film Joe’s Apartment sym­ pathize with the poor treatment roaches have received at the hands of their human cohabit­ ants, while Pixar’s far more profitable WALL‐E (2008) celebrates the resilience of roaches with a lovable and unkillable cockroach sidekick – the last living creature on a terminally polluted planet Earth.

29.14 ­Akihabara Culture: Toys, Video Games, and Anime from Modern Japan Given the unusually high regard for insects in traditional Japanese artwork, it is not surprising that insects continue to provide a source of inspiration for creativity in modern Japan. Insects and other arthropods feature promi­ nently in numerous anime and manga series (animation and graphic novels, respectively) and in the toys and video games that are often spin‐offs of such series. Anime, video games, and related toys or games can be referred to col­ lectively as “Akihabara culture” or Akiba kêi, referring to a trendy shopping district of Tokyo (Hoshina and Takada 2012). In Japan (and increasingly in the rest of the world), toys, video games, manga, and anime series are as popular with adults as with children, and often involve adult themes. Although anime has undergone a recent explosion of popularity with Western audiences, the history of Japanese animation parallels that of the United States; the earliest surviving exam­ ples of Japanese animated films were created in 1917 (Litten 2014). American audiences were introduced to Japanese animation when Astro Boy (Mighty Atom) aired on NBC stations in  1963 (Hoffman 1995). During the 1980s, American writers worked with Japanese anima­ tors to produce cartoon series that were often linked with lines of toys and action figures. The Transformers, a particularly popular series in the United States, involved alien robots that could take on various forms. The Insecticons, allied with the villainous Decepticons, could shift between humanoid and insect modes. The original Insecticons included stag beetle, grass­ hopper, and boll weevil characters, and a later series (the “Deluxe Insecticons”) added a Japanese rhinoceros beetle and a cicada (who was, oddly, named Venom). The action figures that accompanied the animated characters, made by the Japanese company Takara‐Tomy for Hasbro, are detailed, cleverly constructed

29  Insect Biodiversity in Culture and Art

models that incorporate many accurate details of insect morphology. In the same vein, arthro­ pod action‐figure tie‐ins are currently made for several different anime and television series in Japan. The Zoids series, also made by Takara‐ Tomy, is perhaps the most diverse, including battle vehicles modeled after cicadas, stag bee­ tles, rhinoceros beetles, dragonflies, wasps, mantids, and other arthropods (Hoshina and Takada 2012). The contrast between Western and Japanese attitudes towards insects can be seen in video games; while they are often villains or simply targets in games from Western countries, Japanese games often feature heroic insect char­ acters, or insect sidekicks who accompany a sprightly heroine. In Mushihimesama (literally, “insect princess”), a fast‐paced shooting game, insects find themselves on both sides of the bat­ tle as a beautiful young woman teams up with a giant rhinoceros beetle to save her village from enemy insects (Hoshina and Takada 2012). Several role‐playing games, such as Dragon Quest and Monster Farm, also include insect characters. Perhaps the best known and most complex example of this genre is the Pokémon franchise, which began as a video game launched in Japan in 1996 and has since spawned an anime series, card games, toys, clothing, and manga. Like many Japanese children (and chil­ dren in general), Satoshi Tajiri, the creator of Pokémon, enjoyed collecting insects and other small creatures (Bulbapedia 2015a). Inspired by his childhood hobby, he created a game in which the player roams a virtual world full of various habitats to seek, capture, and raise creatures known as Pokémon (a contraction of “pocket monsters”). Pokémon fall into categories known as types, which characterize their attributes and behaviors and come into play when they are paired up with other Pokémon in “battles” that test their particular skills. “Bug” is one of the 18 types, and there are currently 69 Bug‐type Pokémon  –  9.4% of all known “species” (Bulbapedia 2015b). Evoking Tajiri’s own expe­ rience, Bug Pokémon are usually among the first that a player encounters at the beginning of his

or her journey, and they grow (“level up”) quickly. Numerous insect orders are represented among the Bug Pokémon, and insect inspiration is obvious even among a few Pokémon of other types. Most Pokémon “evolve” as they gain more experience points within the game, but in the case of the Bug types, the evolution closely par­ allels the real‐life metamorphosis of the insects on which they are based. Several lepidopteran Pokémon go through the stages of caterpillar, cocoon or chrysalis, and adult (e.g., Caterpie, Metapod, and Butterfree). In the example of Wurmple, the player does not know whether the caterpillar‐like first stage will become a Silcoon or Cascoon (two cocoon‐like Pokémon) and emerge as a butterfly‐like Beautifly or a moth‐ like Dustox, capturing a sense of mystery and anticipation that will be familiar to any ento­ mologist who has ever reared out an unidentifi­ able insect larva. Among hymenopteran‐inspired Pokémon, the transformation from grub to pupa to adult wasp is well illustrated in Weedle, Kakuna, and Beedrill, and social insect biology is recognized in Combee: the player must find and capture a rare female Combee and raise her to produce a Vespiquen, the more powerful “queen bee” Pokémon. A particularly clever twist on Bug‐ type evolution occurs in Nincada, which is based on a last‐instar cicada nymph. The player must have an empty space available in his or her party (which can consist of only six Pokémon at a time) because when Nincada evolves, the evo­ lution results in two new Pokémon: the hand­ some winged adult cicada, Ninjask, and the shed skin, Shedinja, a rather strange Bug/Ghost‐ type Pokémon that evokes the eeriness of empty cicada exuviae (Ballard et al. 2010). Insect behavior is also echoed in the moves that Pokémon learn and use in battle. Bug Bite, Bug Buzz, and Infestation are typical Bug‐type moves. Cricket‐like Bug types may use the move Sing, while wasp‐like Pokémon are more likely to resort to Poison Sting or even Fell Stinger. Pokémon inspired by stag and rhinoc­ eros beetles can often use Fighting‐type moves.

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Intriguingly, many lepidopteran Pokémon are a combination of Poison and Bug types, and they can often use a move known as Poison Powder, in which they shake a toxic powder from their wings or bodies onto an opponent. This detail harkens back to Mushi Mezuru Himegimi (The Lady Who Loved Insects), the 12th‐­century Japanese story of a young woman who defies the social expectations of Heian court life by collecting and raising caterpillars. In the story, the protagonist states that she prefers caterpil­ lars to butterflies because the latter can leave a golden powder on your hand that is dangerous (Keene 1955). The idea of toxic wing scales also came into play in the 1961 daikaiju (“giant monster”) movie Mothra, in which a giant sat­ urniid moth terrorizes Japan. In Japan, where collecting and caring for insects is an industry‐driving hobby, large lucanid bee­ tles and dynastine scarabs are mass‐reared and sold in vending machines, and elements of insect‐ related folk tales have remained intact for 900 years. In our rapidly changing and increasingly globalized society, this persistence of insect‐ related traditions is remarkable, as is its impact on other cultures as Akihabara culture grows in worldwide popularity.

29.15 ­Present and Future Trends in Cultural Entomology An exploration of insects in human culture throughout time documents the striking extent to which they have been our constant compan­ ions and our unyielding adversaries. From the earliest representations of insects carved into bison bone by Cro‐Magnon hunters to the latest video games from Tokyo, we see clear patterns of prominence among several orders of insects, even as our understanding of insect diversity grows. Insects that are bothersome, useful, beautiful, and even tasty have dominated human cultural expression (Hogue 1987). Crickets and grasshoppers were among the first insects to capture our imagination, but later civilizations valued their gentle songs, just

as cicadas were celebrated for the constant background music they offered during ancient Greek summers. The flamboyant coloration of butterflies made them a natural decorative motif for cultures the world over; their spectac­ ular metamorphosis caught the attention of early observers, and might have helped to encourage the first truly scientific explorations of nature. Humans perceived the ordered lives of social insects such as ants, bees, and wasps as a reflection of their own increasingly complex societies, and were captivated by that most mys­ terious and useful insect, the honeybee, whose care and management has driven industry, innovation, and scientific curiosity for thou­ sands of years. Fleas, lice, bed bugs, and flies have pestered humans to the extent that some cultures made them recurring symbols of evil and decay. Even though a few beetles were elevated to divine status in ancient Egypt, their large diver­ sity escaped notice until the period of explora­ tion during the Enlightenment made them favorite subjects of scientific illustrators. (It is noteworthy that although beetles have accom­ panied humans wherever there is stored grain, no artistic, mythological, or poetic reference to stored‐grain pests has come to our attention.) Some insect orders, such as dragonflies, man­ tids, and scorpionflies, were rarely represented, but their outlandish beauty did not escape the artist’s eye, especially in the East. Mayflies appear in some poetic references as symbols of ephemeral life, but like caddisflies and stone­ flies, they have largely been known only to anglers and fly‐tyers. Among the less popular insects, one stands out as a relatively recent cultural addition: the cock­ roach. Although the word “cockroach” has been in English usage since 1624, roaches did not make frequent appearances in creative works until humans began to move into large, urban environ­ ments and shared apartment buildings. Roaches are almost universally unloved, but humorous and often sympathetic portrayals, such as Don Marquis’ 1916 character Archy (Marquis 1970), suggest our resignation to the fact that they are

29  Insect Biodiversity in Culture and Art

here to stay, as well as a grudging admiration for their toughness. Roaches are the insect under­ dogs of our modern, urban world.

29.16 ­The Internet Age Even though insects have provided such abun­ dant inspiration throughout human history, and in spite of the great advances we have made in cataloging and understanding insect diversity and biology, the majority of our population remains relatively ignorant of the smaller crea­ tures that surround us. Thanks to the Internet and other means of global communication, information is more readily available than ever before, but increasing dependence on technol­ ogy and a tendency toward a more “indoor” life­ style have created a growing divide between modern humans and our natural context. This lacking connection is evident in the frequent scares that surface on the Internet, often spread through social media, claiming that some new, hideous insect menace has been discovered. Without fail, the “mutant bug” turns out to be an ordinary and even common insect that the originator of the meme had simply never noticed before, as in the example of robber flies and crane flies being mistaken for “giant mutated mosquitoes” (American Mosquito Control Association 2014; Lennon 2013). On the other hand, insects are gaining popu­ larity with a growing segment of the population that is concerned about environmental issues. Insects have figured prominently in caution­ ary  tales of human environmental blunders in movies such as Mothra and Hayao Miyazaki’s Nausicaä of the Valley of the Wind (1984), but a few real‐world insects are increasingly perceived by the public as “coal‐mine canaries” that can warn us of far‐reaching ecological damage. In keeping with the pattern of prominent orders seen in historical examples, the most successful of these insect mascots are ones that have been traditionally viewed as beautiful or friendly  – particularly the monarch butterfly and the hon­ eybee. The monarch, already beloved as the

“state insect” of seven U.S. states, gained enor­ mous status with anti‐GMO activists after being dubbed “the Bambi of the insect world” by Iowa  State professor Marlin Rice (Weiss 1999). Although the safety of GM crops is still hotly debated, the concern for monarchs brought to attention the destruction of their habitat in North and Central America, and today it would be almost impossible to find a gardening maga­ zine that did not contain an article on butterfly gardening. Some entomologists and ecologists have questioned the actual importance of these efforts to aid monarchs, but at the very least, the monarch has brought insects to the attention of the public and, in this case, the term “non‐target organisms” can apply to the myriad insects that will benefit from gardens planted for a single species. In a similar fashion, the phenomenon of col­ ony collapse disorder (CCD), which decimated honeybee colonies and began to attract media attention around 2006, stirred public con­ cern for these ever‐popular insect companions. The  decline of bee populations encouraged a boom in backyard and rooftop beekeeping and resulted in a dramatic increase in public aware­ ness of the importance of pollinators in general. Concern for honeybees is displayed front and center on websites and Facebook pages, and other bees, such as blue orchard bees, bumble­ bees, and mason bees, also have become polli­ nator mascots. Even in the case of insect mascots for envi­ ronmental causes, modern society is biased toward insect orders that have always enjoyed the cultural limelight; only a small segment of the US population likely would recognize the Coral Pink Sand Dunes tiger beetle. It is likely, however, that the influence of better‐known, more charismatic mascot species such as the monarch and honeybee will encourage greater insect‐awareness in general, and may spur increased investigation and appreciation of the natural world. As our understanding of the diversity of insects and their role in our com­ plex environment continues to improve, that understanding will be reflected in the growing

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panoply of insects that fuel our creativity and are represented in our cultural expressions.

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Index of Arthropod Taxa Arranged by Order and Family. Non‐insect classes and other supraordinal taxa and informal clades are listed as primary entries coordinate with insect orders. Taxa and informal clades between order and family level are treated as secondary entries coordinate with family within the relevant order (or class). Family‐group names below family level are treated as tertiary entries coordinate with genus under the appropriate family. Phthiraptera and Psocoptera are listed separately following their separate treatment in this volume. The family level classification of Phasmatodea is unsettled; the family structure of Phasmida Species File is used here for convenience. Page numbers in bold indicate table entries, and numbers in italic face indicate entries on figures and in figure captions. incertae sedis (extinct) Sinonele (extinct)  351 Srokalarva berthei (extinct)  758

a

Arachnida–Acari  167, 733, 752 Eriophyidae 486 Erythraeidae  69 Orobatida 737 Arachnida–Araneae (= Araneida)  651, 733, 878, 885 Deinopidae Menneus unifasciatus 376 Lycosidae Lycosa hawaiiensis  141 Lycosa howarthi  141 Pisauridae Pisaurina mira 109 Salticidae 168 Habronattus pugillis  82, 89 Phidippus rimator 109 Scitodidae 168 Tetragnathidae Tetragnatha 88

Arachnida–Pseudoscorpionida Chernetidae Cordylochernes scorpioides 820 Arachnida–Scorpiones 733 Archeognatha. See Microcoryphia

b

Blattodea (= Blattaria, Blattoptera)  61, 63, 70, 130, 144, 199, 209, 219, 359–377 (chapter 14), 389, 602, 673, 684, 705, 805, 886 Anaplectidae 362, 363, 364, 371 Archotermopsidae  67, 363, 372 Blaberidae  63, 360, 361, 362, 363, 364, 366, 368, 373, 374, 376 Aptera fusca  360 Blaberinae  364 Blaberus 374 Blaberus craniifer 376 Blaberus discoidalis 374 Calolampra elegans 375 Calolampra solida 375 Diploptera punctata  361, 362 Diplopterinae  364

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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Index

Blattodea: Blaberidae (contd.) Epilampra 369 Epilampra abdomennigrum 369 Epilampra involucris 369 Epilampra irmleri 376 Epilampra maya 369 Epilampra rothi 369 Epilamprinae  364, 369 Eublaberus posticus 369 Geoscapheinae 368 Geoscapheini  364 Geoscapheinae  364 Gromphadorhina portentosa 376 Gyna henrardi 368 Gyninae  364 Macropanesthia rhinoceros  362, 368 Monastria biguttata 369 Nauphoeta cinerea  374, 376 Opisthoplatia orientalis 375 Oxyhaloinae  364 Panchlora 368 Panchlorinae  364 Panesthia  361, 369 Panesthia cribata 362 Panesthiinae  364 Paranauphoeta  364 Paranauphoeta formosana 359, 360 Paranauphoetinae  364 Parasphaeria boleiriana  361, 369 Perisphaeria 368 Perisphaeriinae  364 Pycnoscelinae  364 Pycnoscelus surinamensis  374, 375 Rhyparobia maderae 374 Thorax porcellana  360, 368 Zetoborinae  364 Zuluia 360 Blaberoidea 366–369 Blattellidae. See Ectobiidae Blattidae  63, 360, 362, 363, 364, 369, 374, 675, 684, 692 Archiblattinae  364, 369, 370 Blatta orientalis  362, 369, 370, 374, 375, 376 Blattinae  364, 370 Cartoblatta pulchra 370 Cosmozosteria 370

Duchailluia  364 Duchailluiinae  364, 370 Macrocercinae  364, 370 Melanozosteria 369 Neostylopyga rhombifolia  374, 375 Pelmatosilpha lenti 370 Periplaneta 362 Periplaneta aboriginea 370 Periplaneta americana  369, 370, 373, 374, 375, 376 Periplaneta australasiae  374, 375 Periplaneta brunnea  374, 375 Periplaneta fuliginosa  361, 370, 374, 375 Platyzosteria 369 Polyzosteria flavomaculosa 370 Polyzosteria mitchelli 359 Polyzosteria obscuroviridis 370 Polyzosteria pubescens 370 Polyzosteriinae  364, 370 Pseudoderopeltis albilatera 359 Pseudoderopeltis cf. albilatera  360 Scabina antipoda 370 Shelfordella lateralis 374 Blattoidea 369–373 Corydiidae  63, 360, 362, 363, 363, 365, 370, 374, 376 Anisogamia tamerlana 376 Arenivaga 365 Arenivaga bolliana 376 Arenivaga investigata 365 Corydiinae 363, 365, 365 Ergaula capensis 365 Eupolyphaga sinensis 375 Euthyrraphinae  365 Euthyrrhaphinae 363 Heterogamisca chorpardi 365 Heterogamisca marmorata 365 Holocompsinae 363, 365 Latindiinae 363, 365 Melyroidea magnifica 359 Polyphaga saussurei 374 Polyphagoides 365 Therea petiveriana  361, 362, 365 Therea petiverana 376 Tivia  365 Tiviinae 363, 365 Corydioidea 363–366

Index

Cryptocercidae  63, 359, 360, 362, 363, 364, 371, 372 Cryptocercinae  364 Cryptocercus 360, 361, 362, 371, 373 Cryptocercus clevelandi 362 Ectobiidae (= Blattellidae)  63, 360, 362, 363, 364, 366, 374, 675, 684, 692 Anallacta 360 Anaplectinae (see Anaplectidae) Attaphila  364, 366 Attaphilinae (see Blattellinae) Balta 367 Blattella  361, 362 Blattella germanica  362, 366, 373, 374 Blattellinae  363, 364, 366 Cariblatta imitans 367 Ectobiinae  363, 363, 364, 366, 367 Ectobius  362, 363, 367 Ectobius lapponicus 375 Ectobius pallidus 367 Ectobius panzeri 362 Ellipsidion 367 Euphyllodromia 367 Hemithyrsocera 366 Hemithyrsocera histrio 366 Imblattella 367 Imblattella impar 367 Latiblattella lucifrons 367 Luridiblatta trivittata 367 Margattea nimbata 367 Margatteoidea amoena (extinct)  376 Megaloblatta 367 Nyctibora acaciana 368 Nyctibora brunnea 368 Nyctibora sericea 368 Nyctiborinae  363, 364, 366, 367 Paramuzoa alsopi  361, 367 Paratropes bilunata  368, 376 Parcoblatta  361, 362, 366, 376 Phyllodromica  363, 367 Phyllodromica marginata 367 Phyllodromica megerlei 367 Plectopterinae (see Pseudophyllodromiinae) Prosoplecta 367 Pseudoanaplectinia yumotoi 366 Pseudophyllodromiinae  363, 364, 366

Saltoblattella montistabularis 366 Shelfordina orchidae 367 Supella longipalpa  362, 366, 367, 374 Temnopteryx phalerata 366 Xestoblatta cantralli 366 Xestoblatta hamata 366 Hodotermitidae  67, 180, 183, 363, 372 Kalotermitidae  67, 363, 372 Coptotermes 373 Mastotermes 373 Mastotermes darwiniensis  372, 373, 389 Lamproblattidae 362, 363, 364, 370, 371 Eurycanthablatta 370 Lamproblatta 370 Lamproblatta albipalpus 370 Lamproblattinae  364 Lamproglandifera 370 Mastotermitidae  363, 372 Nocticolidae 362, 363, 365, 365 Nocticola australiensis 366 Panesthiidae  63 Polyphagidae (see Corydiidae) Raphidiomimidae  63 Rhinotermitidae  67, 363 Serritermitidae  363 Stolotermitidae  363, 372 Stylotermitidae  363 Termitidae  67, 363, 371, 372, 373 Amitermes laurensis 373 Macrotermitinae 372 Termitoidae  61, 67, 70, 72, 362, 363, 364, 371, 372, 805, 891 Termopsidae (see Archotermopsidae and Stolotermitidae) Tryonicidae 362, 363, 364, 371 Anaplecta 371 Anaplecta brachyptera 371 Lauraesilpha  364, 371 Lauraesilpha mearetoi 371 Tryonicus  364, 371 Tryonicinae  364

c

Carbotriplurida (extinct) Carbotripluridae (extinct) Carbotriplura (extinct)  159 Chilopoda 168

901

902

Index

Coleoptera  5, 18, 20, 23, 23, 27, 28, 30, 31, 35, 36, 37, 59, 61, 63, 70, 71, 75, 88, 130, 144, 146, 168, 318, 457, 628, 651, 659, 674, 726, 752, 769, 796, 804, 805, 813, 883, 891 Bupestridae  20, 871 Steraspis squamosa 871 Byrrhidae  20, 31 Morychus viridis 31 Simplocaria metallica 39 Cantharidae  20 Carabidae  20, 23, 25, 27, 29, 31, 39, 134, 146, 840 Amara alpina 36 Amara quenseli 28 Anophthalmus schmidti 128 Cicindela albissima 895 Duvalius 146 Notiophilus aquaticus 37 Patrobus assimilis 37 Pseudanophthalmus 146 Pterostichus 39 Pterostichus brevicornis 92 Pterostichus empetricola 92 Pterostichus melanarius 111 Trechiama 146 Trechini 142 Trechus pertyi 30 Cerambycidae  20, 70, 883, 886 Cholevidae (see Leiodidae–Cholevinae) Chrysomelidae  20, 25, 26, 27, 32, 33, 808, 820 Cephaloleia 820 Chelobasis 820 Chrysolina 36 Chrysolina arctica 31 Chrysolina brunnicornis bermani 31 Chrysolina subsulcata 39 Chrysomela septentrionalis  33, 39 Galerucella interrupta 31 Leptinotarsa decemlineata 109 Oulema melanopus 808 Phaedon amoraciae 31 Coccinellidae  20 Coccinella trasversoguttata 42 Coleomegilla maculata 110 Cucujidae  20, 38 Cucujus 71

Cucujus clavipes 38 Curculionidae 4, 20, 25, 31, 37, 88, 821 Anthonomus grandis 890 Hypothenemus hampei 821 Isochnus flagellum 39 Lepidophorus lineaticollis 31 Miocalles 88 Otiorhynchus sulcatus 812 Rhyncophorus ferrugineus 326 Stephanocleonus 31 Trigonoscuta 90 Vitavitus thulius 31 Dermestidae  20 Hydroporini  142 Dytiscidae  20, 25, 29, 35, 36, 129, 134 Bidessini  142 Hydroporus morio 28 Hydroporus polaris 28 Ilybus angustior 36 Elateridae  20, 871 Gyrinus opacus 35 Gyrinidae Halipliidae  20 Hydrophilidae  20 Lampyridae  882, 885 Languriidae 457 Lathridiidae (see Latridiidae) Latridiidae  20, 25, 32, 33 Dienerella elegans 33 Dienerella filum 39 Leiodidae  138, 139, 146, 457 Leptodirus hochenwartii 123, 124, 124, 125, 133, 134, 135 Bathysciola 146 Bathysciotes khevenhuelleri 128 Ptomaphagus 137, 138, 139 Ptomaphagus brevior  138, 139 Ptomaphagus cavernicola  138, 139 Ptomaphagus hirtus 137, 138, 138, 139 Ptomaphagus loedingi 137, 138, 139 Ptomaphagus longicornis 137, 138, 139 Ptomaphagus valentinei  138, 138, 139 Speonomus  128, 146 Stagobius troglodytes 123 Leptinidae 457 Platypsyllinae 457

Index

Lucanidae  318, 884, 886 Meloidae 886 Platypsyllidae (see Leiodidae–Platypsyllinae) Pselaphidae 146 Scarabaeidae  109, 318, 457, 804, 882, 886 Aphodius pseudolividus 108 Aphodius rufipes 111 Coprini 110 Dynastinae  878, 893 Oniticellini 110 Onitini 110 Onthophagini 110 Onthophagus hecate 108 Onthophagus pennsylvanicus 108 Onthophagus taurus 108 Pinotini 110 Scarabaeus sacer 872 Silphidae  20 Staphylinidae 4, 20, 25, 26, 29, 31, 32, 33, 457 Micralymma brevilingua  23, 33 Atheta graminicola  29, 39 Boreophila subplana 29 Micralymma brevilingua 39 Omalium caesum 28 Tenebrionidae  146, 813, 871 Morica hybrida 813 Tribolium 753, 758, 795 Tribolium castaneum  758 Pseudosinella 135 Pseudosinella violenta 134 Sinella 135 Collembola 38, 130, 143, 144, 728, 733 Entomobryidae Bessoniella 146 Pseudosinella 144 Crustacea  127, 733 Crustacea–Isopoda Philosciidae Littorophiloscia  141 Littorophiloscia hawaiiensis  141 Crustacea–Amphipoda Niphargidae Niphargus 137 Crustacea–Bathynellacea Bathynellidae Bathynella 129

d

Dermaptera 61, 63, 71, 144, 210, 315–327 (chapter 12), 457, 886, 891 Anisolabididae  63, 316, 317, 325, 326 Anisolabidae  320 Anisolabis howarthi  141 Anisolabis maritima  141, 317, 323, 326, 327 Ctenisolabis 320 Euborellia 326 Euborellia annulipes 327 Gonolabis electa 324 Paralabis 324 Anisolabis hawaiiensis  141 Apachyidae  63, 316, 325, 326 Arexeniidae 457 Arixeniidae 315, 316, 316, 317, 318, 325 Arixenia  323, 325 Arixenia esau  317 Carcinophoridae  63 Chelisochidae  63, 316, 317, 318, 325, 326 Chelisoches morio  317, 327 Schizoproreus volcanus  317 Diplatyidae  63, 316, 317, 318, 323, 325, 326 Diplatys  323, 325 Diplatys flavicolla 325 Forficulidae  63, 71, 316, 318, 320, 321, 325, 326 Allodahlia scabriuscula  320 Anechura harmandi 323 Chelidurella acanthopygia 327 Doru lineare 324 Forficula auricularia 315, 317, 319, 322, 323, 324, 326, 327 Forficula senegalensis  317, 324 Forficulina  316, 318 Timomenus lugens  321 Hemimeridae 315, 316, 316, 318, 325, 457 Hemimerus  315, 318, 323, 325 Karschiellidae  316, 318, 323, 324, 325 Labiduridae  63, 316, 320, 325, 326 Allostethus indicum 325 Labidura herculeana 92, 320, 323, 327 Labidura riparia  317, 320, 322, 323, 324, 326, 327 Labidura truncata 327 Nala tenuicornis  320

903

904

Index

Dermaptera (contd.) Labiidae  63 Pygidicranidae  63, 316, 317, 320, 323, 325, 326 Challia 325 Challia fletcheri 327 Cranopygia marmoricrura  320, 323 Echinosoma 317 Pyragropsis 325 Tagalina  318, 323 Tagalina papua 325 Spongiphoridae  63, 316, 318, 325, 326 Chaetospania borneensis 323 Geracinae 318 Labia minor  319, 323, 327 Marava arachidis 323 Pseudomarava prominens 323 Vostox brunneipenis 324 Dictyoptera  210, 246, 389, 398 See also Blattodea and Mantodea Diplopoda 821 Arthropleuridae Eoarthropleura (extinct)  765 Diplura 130, 144 Campodeidae 146 Diptera 18, 20, 23, 23, 24, 25, 25, 27, 31, 32, 33, 33, 35, 36, 37, 39, 40, 41, 42, 61, 63, 72, 130, 144, 146, 457, 651, 673, 684, 692, 706, 707, 708, 715, 726, 743, 756, 756, 758, 759, 766, 767, 804, 805, 813, 843, 876, 878, 880, 882, 883, 884, 890, 892 Acartophthalidae  21 Acroceridae  63 Agromyzidae  21, 63, 743, 769 Liriomyza 810 Liriomyza trifolii 810 Apioceridae 754 Anisopodidae  63 Anthomyiidae  21, 25, 27, 28, 31, 41, 63 Asilidae  63 Glaphyropyga dryas 206 Asteiidae  63 Athericidae  63 Aulacigastridae  63 Bibionidae  20, 63 Blephariceridae  63 Bolitophilidae  24

Bombyliidae  63 Braulidae  63 Calliphoridae  21, 27, 33, 41, 63, 872 Boreellus atriceps  33 Cynomya mortuorum  33 Protophormia terraenovae  33, 37 Canaceidae  63 Carnidae 457 Cecidomyiidae 17, 20, 63, 808 Sitodiplosis mosellana 808 Celyphidae  63 Protoculicoides (extinct)  775 Ceratopogonidae  17, 18, 20, 24, 42, 63 Chamaemyiidae  63 Chaoboridae  20, 24, 35, 63 Chironomidae  17, 18, 20, 23, 24, 25, 27, 29, 31, 32, 33, 37, 38, 41, 42, 63 Chaetocladius perennis  32 Diamesia 34 Diamesia aberrata 34 Diamesia arctica 34 Diamesia bertrami 34 Diamesia bohemani 34 Hydrobaenus 42 Limnophyes 36 Limnophyes brachytomus  32 Limnophyes eltoni  32 Limnophyes pumilio  32 Metriocnemus ursinus  32 Oliveridia 42 Orthocladiinae 34 Paraphaenocladius impensus  32 Pseudodiamesa 42 Pseudosmittia  32 Smittia  29, 36, 37, 41 Smittia brevipennis  32 Smittia extrema  32, 37 Chloropidae  21, 63 Clusiidae  63 Coelopidae  33, 41 Conopidae  63 Corethrellidae  63 Ctenostylidae  63 Culicidae  20, 27, 41, 42, 63, 775, 873 Aedes 3 Anopheles  3, 775

Index

Anopheles gambiae 3 Culex quinquefasciatus 818 Ochlerotatus nigripes  34, 37 Curtonotidae  63 Cypselosomatidae  63 Diadocidiidae  24, 63 Diastatidae  63 Diopsidae  63 Dixidae  20, 63 Dolichopodidae  20, 25, 63 Drosophilidae  21, 63 Drosophila  4, 88, 327, 605, 753, 795, 820, 842 Drosophila melanogaster 5 Ramphomyia 41 Empididae  20, 25, 38, 41, 63 Empidoidea  24 Ephydridae  21, 63 Fanniidae  63 Gasterophilidae (see Oestridae) Glossinidae Glossina 774 Heleomyzidae  21, 33, 38, 63 Calacanthia trybomi  33 Heleomyza 133 Heleomyza borealis  33, 38 Neoleria prominens  33 Heteromyzidae  63 Hippoboscidae  63, 457 Hybotidae  63 Keroplatidae  63, 124 Arachnocampa luminosa 124 Lauxaniidae  63 Limoniidae 37, 63 Symplecta scotica 37 Liphistiidae  63 Lonchopteridae  63 Megamerinidae  63 Micropezidae  21, 63, 840, 841, 842, 845, 846, 847, 848 Grallipeza cliffi 848 Grallipeza placidoides 848 Grallipeza spinuliger 848 Taeniapterinae 840 Milichiidae  21, 63 Muscidae 18, 21, 25, 28, 41, 64 Spilogona  41, 42

Mycetophilidae  17, 18, 20, 24, 25, 31, 33, 41, 64 Mydidae  64 Mystacinobiidae 457 Mythicomyiidae  64 Nemestrinidae  64, 754, 756 Florinemestrius pulcherrimus (extinct)  756 Protonemestrius jurassicus (extinct)  756 Neriidae  64 Neurochaetidae  64 Nothybidae  64 Nycteribiidae 457 Odiniidae  64 Oestridae  21, 42, 64 Hypoderma  42, 870 Pediciidae  64 Pedicia hannai  36, 39 Periscelididae  64 Phoridae  21, 64 Piophilidae  21, 41, 64 Pipunculidae  21, 64 Platypezidae  21, 64 Platystomatidae  64 Psilidae  64 Lutzomyia 775 Psychodidae  24, 64, 775 Ptychopteridae  64 Pyrgotidae  64 Rhagionidae  20, 64 Rhiniidae  64 Sarcophagidae  21, 64 Scathophagidae  21, 25, 33, 64 Scathophaga furcata  33 Scathophaga litorea  33 Scatopsidae  20, 33, 64 Scenopinidae  64 Sciaridae  20, 33, 41, 64 Lyoriella 29 Sciomyzidae  21, 64 Sepsidae  64 Simuliidae  2, 5, 20, 25, 42, 64 Sphaeroceridae  21, 33, 64 Stratiomyidae  64, 804 Streblidae  64, 457 Syrphidae  21, 25, 41, 64, 804 Melangyna 40 Syrphus 40

905

906

Index

Diptera (contd.) Tabanidae  20, 64, 884 Tachinidae  21, 41, 64, 324, 840 Ocytata pallipes 324 Triarthria setipennis 324 Tanypezidae  64 Tephritidae  64, 675, 684, 692, 693, 846, 847 Bactocera 693 Sapadrama 846 Trypetinae  846 Thaumaleidae  64 Therevidae  64 Tipulidae  20, 28, 39, 42, 64 Limonia lindrothi 39 Tipula carinifrons  35, 36 Tipuloidea  25, 31 Trichoceridae  20, 27, 33 Trichocera  33 Trichocera borealis 38 Ulidiidae  64 Vermileonidae  64 Xylomyidae  64 Xylophagidae  64

e Embiidina. See Embiodea Embiodea 61, 64, 72, 210, 219–240 (chapter 9), 246, 286, 288 Andesembiidae  220, 231, 233, 236 Andesembia 231 Bryoembia 231 Anisembiidae  220, 222, 232, 233, 236, 238 Anisembia texana  222, 229, 233 Chelicerca  232, 233 Chelicerca galapagensis 233 Dactylocerca 233 Dactylocerca rubra 233 Glyphembia 233 Microembia 238 Saussurembia calypso 229 Saussurembia davisi 233 Archembiidae  220, 231, 233, 234, 235, 236, 238 Archembia  233, 234 Archembiinae 233 Calamoclostes  233, 234

Australembiidae  220, 224, 230, 231, 232, 234, 235 Australembia  222, 234 Metoligotoma  229, 231, 234 Metoligotoma brevispina  230 Metoligotoma incompta 222, 224, 226 Metoligotoma rileyi  232 Clothodidae  220, 221, 222, 224, 228, 230, 234, 235, 239 Antipaluria 235 Antipaluria aequicercata  222 Antipaluria urichi 221, 221, 224, 228, 228, 229, 230, 235 Chromatoclothoda 235 Clothoda  229, 235 Clothoda longicauda 235 Cryptoclothoda 235 Cryptoclothoda spinula 235 Embiidae  64, 219, 220, 224, 230, 231, 233, 235, 238 Dinembia 236 Embia 236 Embia major 236 Embia nuragica  224 Embia ramburi  229, 236, 238 Embolyntha batesi 219 Macrembia 224, 230 Parthenembia reclusa 236 Embonychidae  64, 220, 220, 222, 231, 236 Embonycha interrupta  222, 236 Notoligotomidae  220, 224, 231, 233, 234, 236, 238, 239 Burmitembia (extinct)  236 Burmitembiinae (extinct)  239 Notoligotoma 236 Notoligotoma hardyi  224, 226, 229, 234, 236 Notoligotoma nitens 236 Oligotomidae  64, 219, 220, 222, 223, 224, 236, 239 Aposthonia  236, 237, 238 Aposthonia ceylonica  228, 237 Aposthonia gurneyi 237 Aposthonia japonica 219 Bulbosembia 237 Eosembia 237

Index

Eosembia auripecta  223, 226 Haploembia 237 Haploembia solieri  223, 227, 237 Haploembia tarsalis  223, 224, 227, 228, 237 Lobosembia 237 Oligotoma  237, 238 Oligotoma nigra 221, 222, 223, 229, 237 Oligotoma saundersii 237 Paedembiidae  220, 220, 231, 238 Ptilocerembiidae  220, 238 Ptilocerembia  231, 236, 238 Ptilocerembia catherinae 238 Ptilocerembia rossi 238 Scelembia (see Rhagadochir) Scelembiidae  220, 235, 238 Gibocercus napoe  235, 239 Neorhagadochir moreliensis 238 Rhagadochir virgo  229, 238 Sorellembiidae (extinct)  239 Teratembiidae  64, 220, 239 Dachtylembia 239 Dachtylembia siamensis 239 Diradius 239 Oligembia 239 Paroligembia 239 Teratembia 239 Embioptera. See Embiodea Ephemeroptera 18, 19, 23, 24, 27, 31, 34, 37, 61, 64, 72, 90, 164, 766, 883, 888, 891 Ephemerellidae  19 Baetidae  19, 64 Baetis bundyae 37 Behningiidae  64 Caenidae  64 Ephemerellidae  64 Ephemeridae  64 Heptageniidae  19, 64 Isonychiidae  64 Leptophlebiidae  19, 64 Metretopodidae  19 Neoephemeridae  64 Palingeniidae  64 Polymitarcyidae  64 Potamanthidae  64 Prosopistomatidae  64 Teloganodidae  64

Triassomachilidae (extinct)  164 Triassomachilis uralensis (extinct)  159 Tricorythidae  64 Vietnamellidae  64

g

Glosselytrodea (extinct)  627 Grylloblattodea Blattogryllidae (extinct)  351 Grylloblattidae  73, 199, 210, 246, 335, 336, 337, 338, 338, 339, 341, 342, 351 Galloisiana 340, 341, 343 Galloisiana chujoi  342 Galloisiana kiyosawai  342 Galloisiana kosuensis  342 Galloisiana nipponensis  342, 343 Galloisiana notabilis  342 Galloisiana odaesanensis  342 Galloisiana olgae  342 Galloisiana sinensis  342 Galloisiana sofiae  342 Galloisiana ussuriensis  342 Galloisiana yezoensis 341, 342 Galloisiana yuasai  340, 342 Grylloblatta  338, 341 Grylloblatta barberi  342 Grylloblatta bifratrilecta  342 Grylloblatta campodeiformis  340, 342 Grylloblatta chandleri  342 Grylloblatta chintimini  342 Grylloblatta chirurgica  342 Grylloblatta gurneyi  342 Grylloblatta marmoreus  342 Grylloblatta newberryensis  342 Grylloblatta oregonensis  342 Grylloblatta rothi  342 Grylloblatta scudderi  342 Grylloblatta sculleni  342 Grylloblatta siskiyouensis  343 Grylloblatta washoa  343 Grylloblattella 340, 343 Grylloblattella cheni  343 Grylloblattella pravdini  343 Grylloblattella sayanensis  343 Grylloblattina 343 Grylloblattina djakonovi  343

907

908

Index

Grylloblattodea: Grylloblattidae (contd.) Namkungia  341, 343 Namkungia biryongensis  343 Namkungia magnus  343, 343 Grylloptera. See Orthoptera–Ensifera

h

Hemiptera 19, 23, 27, 30, 31, 32, 33, 33, 35, 36, 37, 61, 64, 65, 66, 71, 72, 75, 77, 130, 144, 210, 457, 484, 487, 501–551 (chapter 19), 591–616 (chapter 20), 658, 673, 682, 684, 691, 692, 693, 715, 730, 752, 804, 805, 883 Acanaloniidae  64, 504, 531, 534, 535, 536, 537, 547, 548 Acanalonia 537 Acanalonia conica  531, 537 Philatis 536 Achilidae  64, 504, 531, 534, 535, 537, 540 Achilinae  504, 537 Apateson 537 Apatesoninae  504, 537 Cixidia 537 Cixidia colorata  531 Ilvia 537 Myconinae  504, 537 Sevia 537 Tropiphlepsia 537 Uniptera 537 Achilixiidae  504, 531, 534, 535, 537 Achilixiinae  504 Achilixius 537 Bebaiotes 531, 537 Bebaiotinae  504 Aclerdidae  612 Adelgidae  599, 602, 608, 612 Adelges 598 Pineus 598 Aetalionidae  502, 514, 516, 521, 522, 525, 526 Aetalion 527 Aetalion reticulatum  514 Aetalioninae  502, 514, 526, 527 Aetalionini 527 Biturritiinae  502, 514, 527 Darthula 526 Darthulini 526 Tropidaspis 514 Aleyrodidae  66, 592, 594, 599, 604, 612, 613

Aleurodicinae 594 Aleyrodinae 594 Bemisia tabaci 591, 592, 604 Udamoselinae 594 Alydidae  65 Anthocoridae  19, 65 Orius 495 Aphalaridae  66, 595 Ctenarytaina eucalypti 603 Pachypsylla venusta 603 Aphelocheiridae  65 Aphididae  19, 66, 108, 108, 110, 592, 594, 598, 599, 602, 604, 608, 612, 769, 805 Acyrthosiphon pisum  604, 605 Acyrthosiphon svalbardicum  31, 36, 39, 42 Aphidinae  594, 608, 609 Aphis glycines 593 Aphis gossypii 111 Aphis nerii  71 Aphis sambuci  592 Cerataphidini  599, 615 Chaitophorinae 594 Cinara 592 Cinara confinis (= Cinara abieticola) 40 Colophina arma 615 Diuraphis noxia 593 Drepanosiphinae 594 Hormaphidinae 608 Lachninae 594 Macrosiphum euphorbiae 604 Myzus persicae 604 Nipponaphis monzeni 609 Pemphiginae 608 Pemphigus obesinymphae 609 Pseudoregma bambucicola 609 Rhopalosiphum padi 614 Sitobion calvulus  31, 36, 39 Therioaphis trifolii (f. maculata)  614 Tuberaphis styraci 609 Aphidoidea  23, 25, 31, 592, 594, 599, 602, 608 Aphrophoridae  64, 502, 510, 512, 518, 519, 520, 521, 675, 684, 692 Aphrophora cribrata 520 Aphrophora maculata 519 Cephisus siccifolia 520 Neophilaenus lineatus  512 Philaenus spumarius  509, 519, 520

Index

Philagra 512 Philagra cf. parva  512 Ptyelus goudoti 520 Aradidae  65 Beesoniidae 595, 612, 613 Belostomatidae  65 Berytidae  65 Caliscelidae  65, 504, 531, 534, 535, 537, 538, 545 Adenissini  535, 538 Asarcopus palmarum 538 Augilini 538 Bruchomorpha jocosa  531 Caliscelinae  504, 538 Caliscelini 538 Ommatidiotinae  504, 538 Peltonotellini 538 Callipappidae  612 Calophyidae  66, 595 Carsidaridae  66, 595 Cercopidae  65, 502, 510, 512, 518, 519, 520, 521, 675, 684, 692 Aeneolamia 519 Amberana 520 Aphrophorinae (see Aphrophoridae) Bourgoinrana 520 Callitettix versicolor 519 Cercopinae  518, 520 Cercopis vulnerata  512 Cosmoscartinae 520 Deois 519 Eocercopidium maculata 518 Ischnorhininae 520 Mahanarva 519 Mahanarva fimbriolata 519 Notozulia 519 Prosapia 519 Zulia 519 Asterolecaniidae  612 Cercopoidea 518–521 Cerococcidae  612 Cicadellidae  2, 4, 19, 65, 503, 508, 509, 513, 514, 516, 521, 522, 523, 524, 525, 675, 684, 692, 693 Agalliinae 523 Aphrodinae  503 Austroagalloidinae  503

Bathysmatophorinae  503 Boundarus 513 Cicadellinae  503, 513, 516, 522, 523, 524 Circulifer tenellus 509 Coelidiinae  503, 522, 523, 524, 530 Cofana spectra 694 Dalbulus elimatus 693 Dalbulus gelbus 693 Dalbulus maidis  524, 693 Deltocephalinae  503, 513, 523, 524 Empoasca fabae 509 Errhomeninae  503 Errhomenus brachypterus 509 Erythroneura palimpsesta  513 Erythroneurini  523, 524 Euacanthellinae 525 Eupelicinae 523 Eupelix cuspidata 509 Eurymelinae  503, 522, 523 Evacanthinae  503, 513 Evansiola 525 Flexamia pict  513 Giustina 524 Hespenedra 513 Hylicinae  503, 522 Hymetta balteata  513 Iassinae  503, 513, 523, 524 Idiocerinae  503, 513, 523 Idiocerus 513 Idiocerus atkinsoni 693 Idiocerus clypealis 693 Idiocerus niveosparsus 693 Idioscopus clypealis 693 Ledra aurita 509 Ledrinae  503, 513, 523 Macropsinae  503, 523 Macrosteles fascifrons 509 Megophthalminae  503, 525 Mileewinae  503 Molopoterus theae 693 Myerslopella 525 Neoaliturus haematoceps 524 Neoaliturus tenellus 524 Neobalinae  503 Neocoelidiinae  503 Neopsinae  503 Nephotettix 693

909

910

Index

Hemiptera: Cicadellidae (contd.) Nioniinae  503, 523 Nirvaninae 523 Paulianiana 525 Pawiloma 513 Phereurhininae  503 Proconiini 522 Rugosana querci  513 Sagmation 525 Scaphytopius 524 Scaphytopius nitrides 524 Scenergates viridis 523 Signoretiinae  504 Tartessinae  504, 524 Typhlocybinae  504, 513, 523, 524 Ulopa carneae 509 Ulopa reticulata  509, 692 Ulopinae  504, 522, 523 Xestocephalinae 523 Cicadidae  65, 502, 508, 509, 511, 516, 517, 874, 876, 878, 882, 883, 885, 886 Cicadettinae  502 Cicadinae  502 Drymopsalta daemeli  511 Hamza ciliaris 517 Magicicada 517 Megapomponia imperatoria  511 Platypleura hirtipennis  511 Quesada gigas 517 Tibicininae  502 Cicadoidea 516–518 Cicadomorpha 511–530 Cimicidae  65, 457, 873 Cixiidae  65, 504, 508, 516, 530, 531, 534, 535, 536, 537, 538, 539, 545, 546 Andes 538 Bennini 537 Borystheninae  504, 538 Bothriocerinae  504, 538 Brixia 538 Cixiinae  504, 538 Cixius 538 Haplaxius crudus  536, 539 Hyalesthes obsoletus  536, 539 Melanoliarus 538 Oliarus  141, 538 Oliarus koanoa  141

Oliarus lorettae  141 Oliarus makaiki  141 Oliarus polyphemus  141 Rhamphixius cf. championi  531 Clastopteridae  502, 510, 512, 516, 518, 519, 520, 521 Beesoniella 521 Clastoptera 512, 520, 521 Clastoptera theobromae 521 Clastoptera undulata 521 Clastopterini 521 Grellaphia 521 Iba  518, 520, 521 Parahindoloides  518, 520, 521 Parahindoloides lumuana 520 Sepullia 521 Sepulliini 521 Taphrotylus 521 Tremapterus 521 Coccidae  19, 66, 598, 601, 602, 612, 615 Coccus hesperidum 596 Parthenolecanium 611 Pulvinaria urbicola 817 Coccoidea 31, 592, 594, 595, 600 Coelidiidae  65 Coelostomidiidae  600, 612 Ultracoelostoma 593 Coleoscytidae 530 Coleoscytoidea 530 Colobathristidae  65 Conchaspidae  612, 613 Coreidae  65, 675, 684, 692 Corixidae  19, 35, 65 Cydnidae  65, 682, 685 Dactylopiidae 595, 612, 658 Dactylopius  601, 602, 658 Delphacidae  19, 65, 504, 509, 531, 534, 535, 538, 539, 540, 674, 675, 684, 691, 692 Asiracinae  504, 539 Copicerus irroratus  531 Delphacinae  504, 539 Delphacini  539, 540 Eumetopina flavipes 540 Javesella dubia 691 Kelisiinae  504, 539 Laodelphax striatellus  536, 540, 691 Nilaparvata lugens  509, 535, 536, 540, 691

Index

Peregrinus maidis  536, 540 Perkinsiella saccharicida  536, 540 Pissonotus 540 Plesiodelphacinae  505, 539 Saccharosydne saccharivora 540 Saccharosydnini 539 Sogatella furcifera  536, 540, 680, 691 Stenocraninae  505, 539 Stobaera 540 Tropidocephalini 539 Ugyops 539 Vizcayinae  505, 539 Derbidae  65, 505, 531, 534, 535, 540, 541, 675, 684, 692, 693 Cedusa 541 Cedusinae  505, 540 Derbinae  505, 541 Diostrombus mkurangai 541 Mysidia 531, 541 Omolicna joi 541 Otiocerinae  505, 541 Pintaliini 540 Proutista moesta  541, 693 Rhotanini 541 Diaspididae  66, 592, 595, 596, 600, 612, 613 Comstockiella  611, 615 Hemiberlesia lataniae 596 Melanaspis obscura  592 Dictyopharidae  65, 505, 532, 534, 535, 541, 543, 675, 684, 691, 692, 693 Dictyopharinae  505, 541, 542 Orgeriinae  505, 541 Paralappida 532 Phylloscelis rubra 541 Rhynchomitra 541 Dinidoridae  65 Dysmorphoptilidae (extinct)  516 Epipygidae  502, 510, 512, 518, 519, 521 Eicissus 512 Epipyga 521 Eriococcidae (sensu lato)  595, 601, 602, 612, 613 Apiomorpha 610 Apiomorpha macqueeni 610 Cystococcus 608 Lachnodius  611, 615

Eurybrachidae  505, 531, 534, 542, 544, 545, 675, 684, 691, 692, 693 Ancyra 531, 542 Eurybrachinae  505 Eurybrachys tomentosa 542 Parancyra bivulnerata 542 Platybrachinae  505 Platybrachus 692 Eurymelidae (see Cicadellidae—Eurymelinae) Flatidae  65, 505, 532, 534, 537, 542, 543, 675, 684, 691, 692, 693 Adexia erminia  532 Flatinae  505, 542 Flatoidinae  505, 542 Metcalfa pruinosa 542 Selizini 542 Zanna 543 Zanninae  506, 543 Fulgoridae  65, 505, 508, 533, 534, 541, 542, 543, 691 Cladodipterinae (see Cladyphinae) Amyclinae  505 Aphaeninae  505 Cladodipterinae 543 Cladyphinae  505, 543 Dichopterinae  505, 543 Fulgorinae  505 Lycorma delicatula 543 Lyncidinae  505, 543 Lystrinae  505 Phenacinae  505 Phrictus quinquepartitus  533 Sclerodepsa granulosa  533 Strongylodematinae  505, 543 Xosopharinae  505 Fulgoroidea  84, 144, 146, 530–548, 693, 730 Fulgoroidea incertae sedis Agenia  507 Aylaella  507 Buca  507 Chondroptera  507 Gastererion  507 Hesticus  507 Hiracia  507 Karna  507 Mijas  507 Ziartissus  507

911

912

Index

Hemiptera (contd.) Gelastocoridae  65 Gengidae  506, 533, 534, 542, 544 Acrometopum 544 Acrometopum panoplites 544 Gengis (see Acrometopum) Microeurybrachys 544 Microeurybrachys vitrifrons  533 Gerridae  65, 885 Hebridae  65 Helotrephidae  65 Henicocephalidae  65 Heteroptera  24, 65, 72, 144, 168, 501, 508, 510, 682, 685, 691, 692, 886 Homotomidae  66, 595 Hydrometridae  65 Hylicellidae 516 Hylicidae (see Cicadellidae) Hypochthonellidae  506, 532, 534, 535, 544 Hypochthonella caeca  532, 544 Hypsipterygidae  65 Icaniidae  65 Issidae  65, 506, 532, 534, 535, 536, 537, 538, 544, 545, 547, 548 Agalmatium bilobum 545 Hemisphaeriini  506, 544 Issini  506, 544 Parahiraciini  506, 544 Sarima nigroclypeata 545 Thabena brunnifrons 545 Thionia 532 Thionia simplex 545 Karajassidae 516 Kermesidae  612 Kerriidae  66, 601, 612 Kinnaridae  65, 506, 532, 534, 535, 545, 546 Kinnara 545 Kinnarinae  506 Oeclidius fraternus  532 Prosotropinae  506, 545 Kuwaniidae  612 Largidae  65 Lecanodiaspididae  601, 612 Ledridae (see Cicadellidae—Ledrinae) Leptopodidae  65 Liviidae  66 Diaphorina citri  591, 603, 604, 607

Lophopidae  65, 506, 533, 534, 542, 544, 545, 546, 547 Acarna 546 Acarnini 546 Acothrura 546 Aluma 546 Apia 546 Asantorga 546 Binaluana 546 Bisma 546 Buxtoniella 546 Carrionia  545, 546 Carrioniini 546 Clonaspes 546 Corethrura 546 Elasmoscelini 546 Elasmoscelis 546 Jugoda 546 Kasserota 546 Lapithasa 546 Lophopinae  506, 546 Lophops 533, 546 Lophops saccharicida  533 Maana 546 Magia 546 Magia subocellata 545 Makota 546 Megacarna 546 Menosca 546 Menoscinae  506, 546 Onycta 546 Painella 546 Painella simmondsi 546 Paracorethrura 546 Podoschtroumpfa 546 Pseudocorethrura 546 Pseudotyxis 546 Pyrilla perpusilla 546 Sarebasa 546 Venisiella 546 Virgilia 546 Virgilia luzonensis 546 Virgiliini 546 Zeleja 546 Zophiuma 546 Zophiuma butawengi 546 Zophiuma lobulata 546

Index

Lygaeidae  19, 27, 65, 682, 685 Nysius groenlandicus  33, 35, 36, 37, 40, 41 Machaerotidae  65, 502, 510, 512, 518, 519, 521 Aecalus 521 Ambonga 521 Aphrosiphon 521 Conditor 521 Enderleinia 521 Enderleiniinae 521 Hemizygon 521 Hindoloides 521 Hindoloidini 521 Labramachaerota 521 Machaerota 521 Machaerota coomani  512 Machaerotini 521 Maxudeini 521 Neuromachaerota 521 Pectinariophyes 521 Pseudoclastoptera 521 Pseudomachaerota 521 Tapinacaena 521 Zygon 521 Zygonini 521 Malcidae  65 Marchalinidae  612 Margarodidae  66, 607, 612 Matsucoccidae 607, 612 Meenoplidae  65, 66, 506, 532, 534, 535, 545, 546 Kermesiinae  506, 547 Meenoplinae  506, 547 Nisia 532 Nisia carolinensis 547 Nisia nervosa 547 Melizoderidae  502, 514, 521, 522, 525, 526 Melizoderes 514 Membracidae  65, 265, 502, 508, 514, 515, 521, 522, 525, 526, 527, 528, 675, 684, 692 Aconophora 526, 528 Aconophorini 527, 528 Acutalini  528 Amastrini  528 Anchistrotus 527, 528 Antianthe 526, 528 Beaufortianini 530

Boccharini 530 Boocerini 529 Campylenchia  528 Centrocharesini  528, 530 Centronodinae  502, 515, 527, 528, 529 Centrotinae  502, 515, 527, 528, 529, 530 Centrotini 530 Centrotus cornutus 509 Centrotypini 530 Ceresa 526 Ceresini  528 Chelyoidea  528 Choucentrini 530 Cladonota  528 Cymbomorphini 529 Darninae  502, 515, 527, 528, 529 Ebhuloidesini  528, 530 Endoiastinae  502, 514, 526, 527, 528, 529 Endoiastus 514 Erechtia  528 Flexocentrus  528 Gargara 526 Gargarini  529, 530 Guayaquila 526, 528 Harmonides  528 Hemikyptha 515 Hemikypthini  515, 527, 528, 529 Heranice miltoglypta  514 Heteronotinae  503, 514, 527, 528, 529 Heteronotus 514 Holdgateilla chepuensis  515 Hoplophorionini  515, 527, 528 Hyphinoini 529 Hypsaucheniini  515, 528, 530 Hypsoprora  528 Hypsoprorini  528 Ischnocentrus  528 Jingkara hyalipunctata  515 Leptobelini 530 Leptocentrini 530 Leptocentrus 526 Lobocentrini 530 Maarbarini 530 Membracinae  503, 515, 527, 528, 529 Membracini  528 Membracis 526 Metcalfiella 526

913

914

Index

Hemiptera: Membracidae (contd.) Metcalfiella vicina  515 Micrutalini  528 Micrutalis 526 Monobelini 529 Nessorhinini 529 Nicomiinae  503, 515, 527, 528, 529 Otinotus 526 Oxyrhachini  528, 530 Oxyrhachis 526 Paracentronodus 515 Philya  528 Pieltainellini 529 Platycentrini 529 Platycentrus  528 Polyglyptini  514, 528 Potnia  528 Quadrinareini 527, 528, 529 Smiliinae  503, 514, 527, 528, 529 Spissistilus 526 Stegaspidinae  503, 515, 527, 528, 529 Stegaspis  528 Stictocephala 526 Talipedini 527, 528, 529 Telamonini  528 Terentiini  528, 530 Thuridini 527, 528, 529 Tragopini  528, 529 Tricentrus 526 Umbelligerus woldai  515 Vanduzea 526 Xiphopoeini 530 Membracoidea 521–530 Mesoveliidae  65 Micronectidae  65 Miridae 4, 19, 65 Chlamydatus acanthioides  33 Chlamydatus pullus  35, 36, 41 Lygus 3 Metriocnemus 36 Monophlebidae  592, 595, 600, 610, 612 Icerya purchasi  592 Iceryini  610, 611 Myerslopiidae  503, 508, 512, 516, 521, 522, 524, 525 Myerslopia rakiuraensis  512 Nabidae  65, 168

Naucoridae  65 Neococcoidea 611 Nepidae  65, 886 Nerthridae  65 Nogodinidae  506, 533, 534, 535, 536, 547 Biolleyana 533, 547 Bladina molorchus 547 Bumerangum deckerti 547 Colpopterinae  506, 547 Colpopterini 536 Gastriniinae  506, 547 Nogodina 547 Nogodininae  506, 547 Tonginae  506, 535, 547 Notonectidae  65, 886 Ochteridae  65 Ortheziidae  19, 600, 611, 612 Palaeontinidae (extinct)  516 Peloridiidae 525 Pentatomidae  65, 675, 682, 684, 685, 692, 818, 878 Nezara viridula 818 Phacopteronidae  66, 595 Phoenicococcidae 595, 612, 613 Phylloxeridae  599, 602, 608 Piesmatidae  65 Pityococcidae  595, 612, 613 Phenacoleachiidae  612 Plataspidae  65 Megacopta cribaria 605 Megacopta punctatissima 605 Pleidae  65 Plokiophilidae  65 Polyctenidae  65, 457 Procercopidae  516, 519 Prosboloidea (extinct)  516 Pseudococcidae  19, 66, 595, 600, 612 Phenacoccinae  600 Planococcus citri 603 Pseudococcus obscurus 613 Pseudococcinae  600 Psyllidae  19, 66, 592, 595, 675, 684, 692 Arytaininae 595 Cacopsylla groenlandica 26 Cacopsylla myrtilli 37 Chamaepsylla hartigii  592 Diaphorina citri  603, 607

Index

Psylloidea  31, 36, 592, 594, 595, 599, 612, 613 Putoidae  600, 612 Pyrrhocoridae  65 Reduviidae  65, 168 Nesidiolestes ana  141 Nesidiolestes selium  141 Rhodnius 327 Zelus 111 Zelus renardii 111 Rhizoecidae  600 Rhopalidae  65 Rhyparochromidae  65 Ricaniidae  65, 507, 533, 534, 535, 542, 544, 547, 548 Kruegeria 547 Pharsalinae  507, 547 Pharsalus 547 Ricania 548 Ricania speculum 548 Ricaniinae  507 Scolypopa australis  533, 548 Silvanana 547 Saldidae  19, 27, 65 Chiloxanthus arcticus  33 Chiloxanthus stellatus  33 Salda littoralis  33 Schizopteridae  65 Scutelleridae  65, 675, 684, 692, 693 Steingeliidae  612 Stictococcidae 595, 612, 613 Stictococcus  611, 615 Stigmacoccidae  612 Stigmacoccus garmilleri 593 Surijokocixiidae 530 Tessaratomidae  65 Tettigarctidae  502, 508, 511, 516, 517 Tettigarcta 517 Tettigarcta crinita  511 Tettigometridae  507, 533, 534, 535, 548, 675, 684, 692 Egropinae  507, 548 Hilda patruelis  533, 548 Hildinae  507, 548 Phalixinae  507, 548 Tettigometrinae  507, 548 Tingidae  65 Triozidae  66, 595

Bactericerca cockerelli 604 Tropiduchidae  65, 507, 533, 534, 535, 537, 545, 548, 549, 675, 684, 692 Elicinae  507, 537, 545, 548 Epora subtilis 691 Gaetuliina 535 Gaetuliini 548 Kallitaxila granulata 548 Lavora 549 Numicia viridis 548 Ommatissus 549 Ommatissus lybicus 548 Pelitropis rotulata  533 Tambinia 549 Trienopinae 535 Trienopini 548 Tropiduchinae  507, 548 Ulopidae (see Cicadellidae—Ulopinae) Urostylididae  65 Veliidae  65 Velocipedidae  65 Xylococcidae  612 Hydropalaeoptera (extinct) Bojophlebiidae (extinct) Bojophlebia (extinct)  159 Hymenoptera 18, 22, 23, 25, 26, 28, 30, 31, 33, 36, 37, 39, 41, 42, 61, 66, 67, 71, 72, 77, 130, 144, 229, 596, 611, 636, 651, 673, 675, 677, 683, 686, 688, 715, 726, 752, 758, 759, 767, 768, 789, 791, 792, 792, 793, 796, 804, 805, 815, 851–864 (chapter 28), 883, 885, 886, 890, 892 Agaonidae  66, 820 Alloxystidae  22, 66 Ampulicidae  66 Andrenidae  683, 854, 855, 856, 859, 861 Andrena 40, 677, 688, 689, 693, 852 Andrena complexa 688 Andrena minutula 688 Andrena stromella 688 Andrena suavis 688 Andrena vaga 688 Andreninae  856 Andrenini  856, 859 Calliopsini  856 Euherbstiini  856 Melitturga clavicornis 688

915

916

Index

Hymenoptera: Andrenidae (contd.) Melitturgini  856 Neffapini  856 Nolanomelissini  856 Oxaeinae  856 Panurginae  856, 859 Panurgini  856 Perditini  856, 859 Protandrenini  856, 859 Protomeliturgini  856 Aphelinidae  66, 686 Apidae  22, 23, 28, 66, 72, 109, 636, 684, 764, 814, 815, 851, 855, 858, 861, 862, 870, 873, 874, 884, 884 Allodapini 852, 859 Ammobatini  858 Ammobatoidini  858 Ancylini  859 Ancyloscelis  859 Anthophorini  858 Apinae  858, 862 Apini  858 Apis 40 Apis cerana  108, 876 Apis dorsata  71 Apis mellifera  107, 804, 851, 852, 871, 876, 888 Biastini  858 Bombini  858, 888, 889 Bombus  40, 107 Bombus bellicosus 811 Brachynomadini  858 Caenoprosopidini  858 Centridini  858 Ceratina 852 Ceratina cognata 108 Ceratinini  859 Coelioxoides  858 Ctenoplectrini  858 Emphorini  859 Epeolini  858 Epeoloides  858 Ericrocidini  858 Eucerini  859 Euglossini 851, 858 Exomalopsini  859 Hexepeolini  858

Isepeolini  858 Manueliini  859, 861 Melectini 851, 858 Melipona subnitida 636 Meliponini  66, 658, 858, 861, 862, 880 Neolarrini  858 Nomada 851 Nomadinae  858, 862 Osirini  858 Osiris 284, 858 Parepeolus  858 Peponapis 815 Protepeolini  858 Rhathymini  858 Tapinotaspidini  859 Tetrapedia 852, 858, 859, 861 Tetrapediini  858 Thyreus 851 Townsendiellini  858 Trigona 111 Xolocopinae  859 Xylocopa 71, 852, 861 Xylocopa confusa 108 Xylocopa dejeani 108 Xylocopa nobilis 108 Xylocopinae  859, 861 Apoidea 804, 856 Argidae  67 Aulacidae  66 Bethylidae  66 Braconidae  22, 23, 31, 33, 38, 39, 41, 42, 66, 72, 817 Aphidius ervi 604 Aphidius leclanti 31 Cotesia kazak 817 Diaeretellus svalbardicum  31, 39 Microplitis croceipes 817 Ceraphronidae  22, 41, 66 Chalcididae  22, 66 Hockeria mengenillarum  680, 681 Chalcidoidea  4, 680 Chrysididae  66, 281 Amiseginae 281 Loboscelidiinae 281 Cleptidae  66 Colletidae  66, 683, 688, 855, 857 Callomelittinae  857

Index

Caupolicanini  857 Colletinae  857 Diphaglossinae  857 Diphaglossini  857 Dissoglottini  857 Euryglossinae  857 Hylaeinae  857 Hylaeus 851 Neopasiphaeinae  857 Paracolletes  857 Scrapterinae  857 Xeromelissinae  857 Crabronidae  66, 684, 854 Pemphredoninae 854 Philanthinae 854 Cynipidae  22, 66 Andricus 609 Callirhytis 609 Diapriidae  22, 66 Diprionidae  67 Dorylidae (see Formicidae—Dorylinae) Dryinidae  66, 72 Dicondylus lindbergi 691 Embolemidae  66 Encyrtidae  22, 66 Eucharitidae  66 Eulophidae  22, 33, 66 Eupelmidae  66 Merostenus 681 Eurytomidae  66 Evaniidae  66 Figitidae  22, 66 Formicidae  22, 66, 72, 90, 179, 368, 397, 593, 681, 683, 686, 687, 774, 874, 877, 878, 879, 882, 883, 884, 885 Anoplolepis gracilipes 93 Attinae 372 Basiceros manni 206 Camponotus 732 Camponotus leonardi  732 Crematogaster laeviuscula 688 Dolichoderinae  683, 686, 813 Dorylinae 179 Eciton 814 Ecitoninae  683, 686, 687 Formicinae  683, 686, 687, 688 Iridomyrmex purpureus 813

Labidus 814 Leptogenys 324 Messor  179, 681 Messor barabarus 681 Myrmeciinae  683, 687, 688 Myrmicinae 179, 683, 686 Neivamyrmex 814 Poerinae  683 Ponerinae  179, 686, 687 Prionopelta amabilis 206 Pseudomyrmecinae  683, 686, 687 Pseudomyrmex 368 Pseudomyrmex gracilis 179 Solenopsis 185 Solenopsis invicta  327, 688 Tetramorium bicarinatum 817 Gasteruptiidae  66 Halictidae  66, 683, 688, 855, 856 Augochlorini  857 Caenohalictini  856, 857 Conanthalictini  856 Dialectus 40 Halictinae  856 Halictini 852, 857, 859 Lasioglossum 689 Nomiinae  856 Nomioidinae  856 Penapini  856 Rophitinae  856 Rophitini  856 Sphecodini  856, 857 Thrinchostomini  856, 857 Xeralictini  856 Ibaliidae  66 Ichneumonidae  22, 23, 25, 28, 31, 33, 38, 41, 66, 72 Aclastus borealis 41 Atractodes pusillus 41 Gelis glacialis 41 Stenomacrus groenlandicus  39, 41 Leucospidae  66 Megachilidae  22, 66, 851, 855, 857, 860, 861, 862 Afroheriades  857 Anthidiini  858, 861, 862 Aspidosmiini  857, 861 Dioxyini  857

917

918

Index

Hymenoptera: Megachilidae (contd.) Fidelia 860 Fideliinae 855, 857, 860, 861 Lithurginae  857 Megachile pluto 851 Megachilinae 851, 856, 857, 862 Megachilini  857 Neofidelia 860 Noteriades  857 Ochreriades  858 Osmiini  857 Pararhophitinae  857, 860, 861 Proteriades bullifacies 852 Pseudoheriades  857 Stelis 862 Megalyridae  66 Meliponidae (see Apidae—Meliponini) Melittidae  66, 683, 851, 855, 856, 860, 861 Dasypodainae  856, 860, 861 Dasypodaini  856 Hesperapini  856 Hesperapis leucura 688 Hesperapis rhodocerata 688 Macropidini  856 Macropis nuda 852 Meganomiinae  856, 860 Melittinae 851, 856 Rediviva 851 Sambini  856 Mutillidae  66 Mymaridae  22, 66 Orussidae  67 Pamphiliidae  67 Platygastridae  22, 66, 229 Gryonoides glabriceps  790 Pompilidae  66 Proctotrupidae  22, 66 Pseudomyrmecidae (see Formicidae—Pseudomyrmecinae) Pteromalidae  22, 38, 66 Dibrachys microgasteri 681 Nasonia 606 Rhopalosomatidae  66 Roproniidae  66 Rotoitidae  66 Sapygidae  66 Scelionidae (see Platygastridae)

Scolebythidae  66 Scoliidae  66 Signiphoridae  66 Siricidae  22, 67 Sphecidae  66, 683, 684, 689 Ammophila 690 Chlorion cyaneum 376 Krombeinictus nordenae 855 Stenotritidae 855, 857, 860 Stephanidae  67 Tenthredinidae  22, 23, 25, 26, 28, 67, 72 Nematinae 26 Pristophora 36 Tiphiidae  66 Torymidae Idiomacromerus gregarius 680, 681, 681 Trichogrammatidae  67 Trigonalidae  67 Vespidae  22, 67, 72, 683, 684, 689, 845, 874 Anterhynchium 690 Dolichovespula 40 Masarinae 855 Polistes  689, 690 Polistes carnifex  689, 690 Polistes dominula  680 Polistes gallicus 690 Polistes stabilis 690 Provespa 690 Rhynchium 690 Vespa 690 Vespa analis 690 Vespa crabro 690 Vespa ducalis 690 Vespa dybowskii 690 Vespa mandarinia 690 Vespa simillima 690 Xiphydriidae  67

i

Isopoda 821 Isoptera. See Blattodea —Termitoidae

l

Lepidoptera  5, 17, 18, 21, 23, 25, 27, 28, 31, 33, 36, 41, 61, 67, 71, 72, 73, 228, 367, 405, 457, 596, 597, 651, 715, 726, 752, 756, 758, 759, 762, 769, 792, 804, 805, 810, 811, 817, 883, 890, 892

Index

Adelidae  67 Agonoxenidae  67 Alucitidae  67 Amphiteridae  67 Arctiidae (see Erebidae–Arctiinae) Attevidae  67 Bedelliidae  67 Blastobasidae  67 Bombycidae  67 Bombyx mori 877 Brachodidae  67 Brahmaeidae  67 Callidulidae  67 Carposinidae  67 Choreutidae  67 Coleophoridae  67 Cosmopterigidae  67 Cossidae  67 Crambidae  67 Cnaphalocrocis patnalis 109 Ctenuchidae  67 Cyclotornidae  67 Danaidae (see Nymphalidae) Drepanidae  67 Elachistidae  67 Endromidae  67 Epermeniidae  67 Epicopeiidae  67 Epipyropidae  67 Erebidae  67 Arctiinae  22, 35, 67, 72 Gynaeophora groenlandica  36, 37, 39 Lymantriinae  22, 35, 67 Olene mendosa 596 Pareuchaetes pseudoinsulata 810 Schrankia altivolans  141 Schrankia howarthi  141 Ethmiidae  67 Eupterotidae  67 Gelechiidae  21, 67 Geometridae  22, 25, 35, 67, 72, 810, 821 Biston betularia 810 Eupithecia 91 Operophtera brumata  26, 33 Psychophora 36 Psychophora cinderella 35 Glyphipterigidae  67

Gracillariidae  67, 774 Hepialidae  67 Hesperiidae  21, 67, 72 Himantopteridae  67 Hyblaeidae  67 Immidae  67 Incurvariidae  21 Lasiocampidae  67 Lecithoceridae  67, 72 Limacodidae  67 Lycaenidae  21, 67, 72 Polyommatus 30 Lymantriidae  67 Micropterigidae  67 Neothoridae (see Hepialidae) Nepticulidae  67 Noctuidae  22, 25, 27, 35, 67, 72, 811, 817 Agrotis infusa 811 Apamea zeta 28 Helicoverpa 811 Helicoverpa armigera 817 Helicoverpa zea  108, 108 Xestia aequaeva 35 Nolidae  67 Notodontidae  67 Nymphalidae  21, 28, 36, 67, 72, 757, 762 Boloria  27, 30, 36, 37 Boloria chariclea  26, 42 Danaus plexippus  811, 895 Erebia 27 Nymphalis antiopa 40 Oenis 27 Satyrinae  21, 28 Vanessa cardui 40 Oecophoridae  67, 72 Palaeosetidae  67 Papilionidae  21, 67, 72, 886 Graphium antiphates  71 Papilio 71 Papilio multicaudatus 880 Papilionoidea  872, 874, 878, 880, 883, 884, 885 Pieridae  21, 28, 68, 72 Colias  27, 36, 37 Colias hecla  26, 42 Pieris napi 40

919

920

Index

Lepidoptera (contd.) Plutellidae  21, 807, 811 Plutella polaris 31 Plutella xylostella  40, 807, 811 Psychidae  68 Pterophoridae  21, 68 Pyralidae  22, 68, 72, 457 Pyla fusca 28 Riodinidae  68 Roeslerstammiidae  68 Saturniidae  68 Satyridae (see Nymphalidae–Satyrinae) Scythrididae  68 Sesiidae  68 Sphingidae  22, 68, 72, 85 Manduca 753 Thyrididae  68 Tineidae  68, 813 Pringleophaga marioni 813 Tortricidae  21, 68, 72 Torymidae  66 Uraniidae  68 Urodidae  68 Yponomeutidae  68 Zygaenidae  68

m

Mantodea 61, 68, 73, 219, 359, 389–407 (chapter 15), 650, 673, 683, 686, 687, 871, 878, 885, 886, 891 incertae sedis Haaniinae  392, 404, 405 Ambermantis 406 Haania lobiceps 406 Hoplocoryphinae 405 Acanthopidae  393, 395, 396, 397, 404 Acanthops  395, 396 Acontiothespinae (see Acontistidae) Acontistinae (see Acontistidae) Stenophyllinae (see Stenophyllidae) Acontistidae  393, 395, 396, 397, 404 Acontista 390 Callibia diana 396 Amorphoscelidae  68, 391, 392, 393, 395, 397, 398, 401, 406 Amorphoscelinae 391, 393, 395, 397, 398 Chloroharpax 397

Exparoxypilus africanus 397 Paramorphoscelis 397 Paramorphoscelis gondokorensis 397 Paraoxypilinae 391, 393, 397, 401 Perlamantinae 391, 393, 395, 397 Perlamantis 397 Angelidae  393, 395, 398 Angela 398 Angela guianensis 398 Angelinae  392, 398 Chaeteessidae 391, 393, 395, 395, 398, 402, 406 Chaeteessa  395, 396, 398, 399, 405, 406 Coptopterygidae  393, 395, 399 Brunneria 399 Brunneria borealis 399 Coptopterygini 399 Coptopteryx 399 Empusidae  68, 391, 393, 395, 399, 400, 402 Blepharodinae  393 Empusinae  393 Empusini  393 Gongylus 395 Gongylus gongylodes 399 Idolomantis 391 Idolomantis diabolica 399 Idolomorphini  393 Epaphroditidae  393, 395, 399, 400 Brancsikia  399, 400 Epaphrodita  399, 400, 401 Epaphroditinae 399 Eremiaphilidae 391, 392, 393, 395, 400, 406 Eremiaphila 395 Heteronutarsus  396, 400 Galinthiadidae  393, 395, 396, 400, 401 Congoharpax 400 Galinthias 400 Harpagomantis  397, 400 Pseudoharpax 400 Gryllomantidae  68 Hymenopodidae   68, 391, 393, 395, 396, 399, 400, 401, 402 (see also Galinthiadidae) Acanthopinae (see Acanthopidae) Acromantinae  393 Acromantini  393 Anaxarchini  393 Ceratomantis 401

Index

Chlidonoptera  397, 401 Chrysomantis 401 Creobroter 401 Heliomantis 401 Helvia 401 Hymenopodinae  393, 395, 401 Hymenopodini  393, 400 Hymenopus  395, 400, 401 Hymenopus coronatus  390, 401 Junodia 401 Otomantini  393 Otomantis 401 Oxypilinae  393 Oxypiloidea 401 Oxypilus 401 Pachymantis 401 Parablepharis 399 Phyllocrania 399 Phyllocraniinae  393, 399 Phyllothelyinae  393, 401, 402 Pseudocreobotra  400, 401 Sibyllinae  393, 401, 402 Tithrone 397 Iridopterygidae  68, 392, 393, 395, 401 Ciulfina 401 Cornucollis 401 Enicophlebia hilara  390, 401 Fulcinini  393 Hapalomantinae  393, 401 Hapalomantini  393 Hapalopeza 392, 401 Hyalomantis 401 Ilomantis 401 Iridopteryginae  393, 401 Nanomantinae  393, 401 Nilomantinae  393, 401 Nilomantis 401 Platycalymma 401 Stenomantis 401 Tarachina 392 Tropidomantinae  393, 401 Tropidomantini  393 Jantarimantidae  68 Liturgusidae  68, 391, 392, 393, 395, 401, 402, 406 Corticomantis 402 Fuga 402

Gonatista 400 Hagiomantis 402 Liturgusa  402, 406 Liturgusa cursor  390 Liturgusini 391 Theopompa 406 Velox 402 Mantidae  68, 393, 394, 395, 398, 399, 401, 402, 403, 683, 687 Ameles 395 Amelinae  392, 393, 398, 401 Amelini  392, 393 Amphecostephanus 399 Angela guianensis 398 Antemna 399 Antemninae  392, 393 Apteromantis 395 Archimantinae  393 Callimantis 400 Choeradodinae  393, 402 Choeradodis  395, 402 Chroicoptera 402 Chroicopterinae  392, 393 Compsothespinae  392, 393, 402 Compsothespis 390, 395, 398, 402 Danuriini  392 Deroplatyinae  394, 398 Deroplatys 402 Dystactinae  392, 394 Heterochaetini  392 Hierodula 402 Ligaria 402 Liturgusinae (see Liturgusidae) Mantinae  392, 394 Mantis 402 Mellierinae  394 Miomantinae  394, 404 Mythomantis 398 Orthoderinae  394, 402 Oxyothespinae  392, 394, 402 Photinainae (see Photinaidae) Popa spurca  390 Pseudempusa 395 Pseudoyersinia 395 Rivetina 402 Schizocephala 402 Schizocephalinae 391, 392, 394, 402, 404

921

922

Index

Mantodea: Mantidae (contd.) Sphodromantis 402 Stagmatopterinae  392, 394 Stagmetopterinae  683 Stagmomantinae  392, 394 Tarachodinae (see Tarachodidae) Thespoides bolivari = Angela guianensis 398 Tisma 402 Tisma grandidieri  390 Toxoderinae (see Toxoderidae) Vatinae 391, 392, 394, 402 Mantoididae 391, 394, 395, 395, 398, 402, 403, 406 Mantoida 390, 398, 403, 405, 406 Mantoida maya 402 Paramantoida 403 Metallyticidae  68, 394, 395, 395, 403, 406 Metallyticus  391, 396, 399, 403, 406 Metallyticus splendidus 403 Metallyticus violaceus 403 Photinaidae  394, 395, 399, 403, 404 Cardiopterinae  394, 403 Macromantinae  394, 403 Microphotinini  394 Orthoderellini  394 Photinaini  394 Photiomantinae  394, 403 Photiomantis 404 Stenophyllidae  394, 395, 396, 397, 404 Stenophylla 404 Tarachodidae  392, 394, 395, 400, 402, 404 Caliridinae  392, 394, 404 Leptomantella 392 Oxyelaea elegans  390 Pyrgomantis 404 Iris 404 Oxyelaea 404 Paroxyophthalmus 404 Tarachodes 404 Tarachodula pantherina 404 Thespinae (see Thespidae) Thespidae  68, 394, 395, 404, 405 Aethalochroaini  394 Bantiinae  394 Calamothespis 405 Haaniinae (see Mantodea incertae sedis)

Hoplocoryphinae (see Mantodea incertae sedis) Miobantiinae  394 Musoniellini  394 Pogonogaster 406 Pogonogaster tristani 406 Pseudomiopteryginae  394 Pseudopogonogastrinae  394 Thespinae  392, 394, 402 Thespini  394 Toxoderidae  68, 392, 394, 395, 398, 400, 404, 405 Toxodera 391 Toxodera beieri 405 Toxoderinae  394, 395, 402 Toxoderini 405 Toxoderopsini (see Toxoderini) Mantophasmatodea  73, 199, 210, 246, 343 incertae sedis (extinct) Adicophasma grylloblattoides (extinct)  345, 351 Adicophasma spinosum (extinct)  344, 351 Ensiferophasma velociraptor (extinct)  345, 351 Juramantophasma sinica (extinct)  345, 351 Raptophasma groehni (extinct)  344, 351 Raptophasma kerneggeri (extinct)  344, 351 incertae sedis (Tyrannophasma/ Praedatophasma Clade)  350 Praedatophasma 347 Praedatophasma maraisi  344, 350 Tyrannophasma 347 Tyrannophasma gladiator  344, 350 Austrophasmatidae  340, 344, 347, 350, 351 Austrophasma  344, 350 Austrophasma caledonense  344, 350 Austrophasma gansbaaiense  344, 350 Austrophasma rawsonvillense  344, 350 Hemilobophasma  344, 350 Hemilobophasma montaguense  344, 350 Karoophasma 350 Karoophasma biedouwense  340, 344, 346, 350, 351 Karoophasma botterkloofense  344, 350 Lobatophasma 350 Lobatophasma redelinghuysense  344, 350, 351

Index

Namaquaphasma 350 Namaquaphasma ookiepense  344, 350 Striatophasma naukluftense  344, 350 Viridiphasma 350 Viridiphasma clanwilliamense  340, 344, 346, 350, 351 Mantophasma 349 Mantophasma gamsbergense  344 Mantophasma kudubergense  344 Mantophasma omatakoense  344 Mantophasma zephyrum  344, 349 Pachyphasma  344, 349 Pachyphasma brandbergense  344 Sclerophasma 349 Sclerophasma paresisense  344, 349, 349 Mantophasmatidae  344, 347, 349 Tanzaniophasmatidae  339, 344, 349 Tanzaniophasma subsolanum  344, 349 Mecoptera 61, 68, 73, 77, 705–716 (chapter 23), 756, 756, 762, 876 Aneuretopsychidae (extinct)  756 Jeholopsyche liaoningensis (extinct)  756 Apteropanorpidae 705, 706, 710, 712 Apteropanorpa 712 Austropanorpidae 712 Bittacidae  68, 706, 706, 707, 708, 712 Afrobittacus 709 Anabittacus 709 Anomalobittacus 709 Apterobittacus 709 Austrobittacus 709 Bicaubittacus 709 Bittacus  708, 709 Edriobittacus 709 Eremobittacus 709 Harpobittacus 709 Harpobittacus similis  709 Hylobittacus 709 Issikiella 709 Juracimbrophlebia ginkgofolia (extinct)  763 Kalobittacus 709 Nannobittacus 709 Neobittacus 709 Orobittacus 709 Pazius 709 Symbittacus 709 Terrobittacus 709

Thyridates 709 Tytthobittacus 709 Boreidae  706, 706, 710 Boreinae 710 Boreus 711 Boreus californicus  710 Caurininae 710 Caurinus 710 Hesperoboreus 710 Choristidae  706, 712 Chorista 712 Neochorista 712 Taeniochorista 712 Cimbrophlebiidae (extinct)  707 Juracimbrophlebia ginkgofolia (extinct)  707 Eomeropidae  706, 711, 712 Eomerope (extinct)  711 Notiothauma 711 Meropeidae  68, 705, 706, 711 Austromerope 711 Merope 711 Mesopsychidae (extinct)  756 Lichnomesopsyche gloriae (extinct)  756 Nannochoristidae  706, 706, 707 Nannochorista 707 Nannochorista andica  707, 707 Notioathaumidae (see Eomeropidae) Orthophlebiidae 712 Panorpidae  68, 73, 706, 706, 712, 713 Baltipanorpa 713 Dicerapanorpa 715 Furcatopanorpa 715 Jurassipanorpa (extinct)  713 Leptopanorpa 715 Neopanorpa 715 Panorpa  713, 714, 715 Panorpa coreana  714 Sinopanorpa 715 Panorpodidae  706, 712, 713 Brachypanorpa  712, 713 Panorpodes  712, 713 Panorpodes paradoxa 712, 713 Pseudopolycentropus janeannae (extinct)  756 Pseudopolycentropodidae (extinct)  68, 756

923

924

Index

Megaloptera 24, 61, 68, 73, 77, 642–644 Corydalidae  68, 73, 629, 631, 632, 634, 635, 638, 642, 642, 644 Acanthacorydalis 642 Chauliodinae  638, 642 Chloronia 642 Corydalinae  638, 642 Corydalus 642, 642 Neochauliodes tonkinensis 73 Nigronia 642 Sialidae  631, 634, 638, 642, 642 Sialis lutaria  642 Meganisoptera (extinct) Meganeuridae (extinct)  773 Merostomata—Xyphosura Limulidae 733 Microcoryphia (= Archaeognatha)  61, 73, 168–175, 728, 733 incertae sedis Charimachilis  156, 157, 173, 174 Mesomachilis nearcticus  170 Machilidae  68, 155, 156, 164, 165, 167, 169, 169, 170, 171, 172, 173, 174, 175, 177, 183, 184, 185, 186 Afrochilis  157 Afromachilis  157 Allopsontus  157 Machilanus 170 Bachilis  157, 169, 173 Catamachilis  157 Coreamachilis  157, 170 Corethromachilis (extinct)  157 Cretaceomachilis (extinct)  165, 185 Cretaceomachilis libanensis (extinct)  164 Dilta  157, 167, 169, 170, 171, 172, 186 Dilta bitschi 171 Dilta hybernica 171 Dilta littoralis  171, 172 Dilta machadoi 172 Ditrigoniophthalminae  156, 174, 175 Ditrigoniophthalmus  157, 173, 174 Graphitarsus  157 Haslundichilis  157, 169, 170 Haslundiella  157 Heteropsontus  157 Heteropsontus americanus 175

Himalayachilis  157 Hybographitarsus 167 Janetschekilis  157 Kerkiratrobius  158 Lepismachilis  157, 186 Leptomachilis  158 Machilinae  156, 174, 177 Machilinus  158, 169, 170, 172, 173 Machilis  157, 165, 167, 170, 171, 172, 173 Machilis engiadina 167 Mendeschilis  157 Mesomachilis  156, 157, 168, 171, 173, 174 Mesomachilis nearcticus 168 Metagraphitarsus  157 Metamachilis  157 Meximachilis  158 Neomachilis  158 Neomachilis halophila  172, 173 Paramachilis  157, 186 Parapetrobius  158, 173 Parapetrobius azoricus 173 Pedetontinus  158 Pedetontoides  158 Pedetontus 186 Petridiobius  158, 173 Petrobiellinae  156, 171, 173, 174 Petrobiellus  158 Petrobiinae  156, 171, 173, 174, 183, 186 Petrobius  158, 173 Petrobius brevistylis 173 Praemachilis  157 Praemachiloides  157 Praetrigoniophthalmus  157 Promesomachilis  157, 167, 169, 170, 186 Promesomachilis hispanica  167, 173, 184 Pseudocatamachilis  157 Pseudomachilanus  157 Silvestrichilis  157 Silvestrichiloides  157 Songmachilis  157 Stachilis  158 Trigoniomachilis  158 Trigoniophthalmus  158, 172 Trigoniophthalmus alternatus  171, 185 Turquimachilis  156, 157, 174 Wygodzinskilis  158

Index

Meinertellidae  68, 156, 164, 165, 166, 168, 169, 169, 170, 171, 172, 173, 174, 175, 183, 185 Allomeinertellus  158 Hypermeinertellus  158 Hypomachiloides  158 Machilelloides  158, 172 Machilellus  158 Machilinus rupestris 173 Machilontus 169, 170 Machilontus yoshii 168 Macropsontus  158, 165 Madagaschiloides  158 Meinertellinae 165 Meinertellus  158, 166, 175 Neomachilellus  158, 165, 175, 185 Neomachilellus dominicanus (extinct)  165 Nesomachilis  158 Nesomeinertellus  158 Patagoniochiloides  158 Praemachilellus  158 Pseudomeinertellus  158 Triassomachilidae (see Ephemeroptera–Triassomachilidae) Monura (extinct) Dasyleptidae 159 Dasyleptus (extinct)  159 Dasyleptus brongniarti (extinct)  159 Dasyleptus lucasi (extinct)  159 Lepidodasypus sharovi (extinct)  159

n

Neuroptera 20, 24, 61, 68, 73, 644–656, 756, 756, 759, 762, 763, 796, 883, 891 Ascalaphidae  68, 628, 629, 631, 633, 634, 635, 638, 642, 644, 652 Albardia furcata 644 Albardiinae  638, 644, 645, 652 Ascalaphinae  638, 644 Haplogleniinae  639, 644, 652 Suphalomitus 642 Berothidae  68, 628, 629, 630, 631, 631, 635, 636, 637, 638, 639, 642, 645, 651 Berothimerobiinae  639, 645 Berothinae  639, 645 Cyrenoberothinae  639, 645

Nosybinae  639, 645 Nyrminae  639, 645 Protobiellinae  639, 645 Rhachiberothinae 637, 639, 645, 651 Spermophorella 642 Trichomatinae  639, 645 Chrysopidae  20, 68, 628, 630, 631, 631, 633, 635, 636, 638, 639, 642, 645, 646, 649, 658 Anomalochrysa 646 Apochrysinae  639, 646 Chrysoperla  111, 646, 658, 659 Chrysoperla carnea 111 Chrysopinae  639, 646 Hypochrysa elegans  642 Nothochrysinae  639, 646 Coniopterygidae  68, 628, 631, 633, 634, 638, 639, 647, 649, 658 Aleuropteryginae  639, 647 Brucheiserinae  640, 647 Coniocompsini  639 Coniopteryginae  640, 647 Dilaridae  68, 629, 630, 631, 635, 638, 640, 647, 649 Berothella 647 Dilar 647 Dilarinae  640, 647 Nallachiinae  640, 647 Nallachius 647 Nallachius americanus 647, 649 Neonallachius 647 Hemerobiidae  20, 68, 628, 630, 631, 633, 634, 635, 638, 640, 649, 649, 658 Adelphohemerobiinae  640 Adelphohemerobius enigmaramus  649 Carobiinae  640 Drepanacrinae  640 Drepanepteryginae  640 Drepanepteryx phalaenoides  649 Hemerobiinae  640 Megalominae  640 Microminae  640 Notiobiellinae  640 Psychobiellinae  640 Sympherobiinae  640 Sympherobius 658

925

926

Index

Neuroptera (contd.) Ithonidae  68, 630, 631, 635, 637, 638, 640, 649, 649, 650 Adamsia 650 Fontecilla 650 Ithone 650 Ithone fulva  649 Megalithone 650 Narodona 650 Oliarces 650 Platystoechotes 650 Polystoechotes 650 Rapisma 650 Varnia 650 Kalligrammatidae (extinct)  756, 757, 759, 762, 763, 773 Kalligramma aciedentatus (extinct)  756 Mantispidae  68, 628, 629, 630, 631, 631, 635, 636, 638, 640, 645, 649, 650, 651 Calomantispinae  640, 650, 651 Climaciella brunnea 650 Drepanicinae  640, 650, 651 Mantispinae  640, 650, 651 Plega hagenella 636 Symphrasinae  640, 650, 651 Trichoscelia 650 Zeugomantispa minuta  649 Myrmeleontidae  68, 628, 629, 630, 631, 633, 634, 635, 636, 638, 640, 649, 651, 652, 879 Acanthaclisini  640 Albardia 652 Austrogymnocnemia edwardsi  649 Myrmeleontinae  640, 651 Palparinae  641, 651 Stilbopteryginae  641, 645, 651, 652 Stilbopteryx 652 Synclisis baetica  649 Nemopteridae  629, 630, 631, 634, 635, 638, 641, 649, 652 Crocinae  641, 652, 653 Nemoptera sinuata  649 Nemopterinae  641, 652, 653 Nevrorthidae 629, 631, 633, 634, 638, 641, 653 Austroneurorthus 653 Nevrorthus 653 Nipponeurorthus 653 Sinoneurorthus 653

Nymphidae 630, 631, 631, 632, 635, 638, 641, 653, 654, 655 Myiodactylinae 655 Myiodactylus 655 Nymphes myrmeleonoides 632, 654 Osmylops 655 Osmylidae  68, 628, 631, 634, 635, 638, 641, 654, 655 Eidoporisminae  641, 655 Gumilla 655 Gumillinae  641, 655 Kempyninae  641, 655 Osmylinae  641, 655 Osmylus fulvicephalus 655 Porisminae  641, 655 Porismus strigatus  654 Protosmylinae  641, 655 Spilosmylinae  641, 655 Stenosmylinae  641, 655 Polystoechotidae (see Ithonidae) Psychopsidae  68, 628, 631, 631, 635, 638, 641, 654, 656, 759 Balmes 656 Psychopsinae  641, 656 Psychopsis insolens  654 Zygophlebiinae  641, 656 Rapismatidae (see Ithonidae) Rhachiberothidae (see Berthridae–Rhachiberothinae) Saucrosmylidae (extinct)  762, 763 Bellinympha filicivora (extinct)  763 Sisyridae  68, 631, 632, 634, 638, 641, 653, 654, 656 Climacia 656 Sisyborina 656 Sisyra 656 Sisyra fuscata  654 Sisyra terminalis  654 Sisyrina 656 Notoptera 61, 73, 637 See also Grylloblattodea and Mantophasmatodea

o

Odonata  61, 73, 878, 881, 883, 885, 885, 886, 889 Aeshnidae  19, 68

Index

Agrionidae  68, 73 Amphipterygidae  68 Calopterygidae  68 Chlorocyphidae  68 Chlorogomphidae  68 Chlorolestidae  68 Coenagriidae (see Coenagrionidae) Coenagrionidae  19, 68, 818 Ischnura hastata 92 Megalagrion calliphya 818 Cordulegastridae  68 Corduliidae  19, 68 Disparoneuridae  68 Euphaeidae  68 Gomphidae  68, 73 Lestidae  69 Libellaginidae  69 Libellulidae  69 Megapodagrionidae  69 Platycnemididae  69 Platystictidae  69 Protoneuridae  69 Zygoptera 886 Orthoptera 19, 61, 69, 73, 75, 144, 202, 209, 210, 245–271 (chapter 10), 613, 673, 683, 684, 686, 687, 692, 804, 883 Acrididae  19, 69, 73, 247, 250, 251, 253, 257, 264, 268, 269, 270, 872, 873 Acridinae 268 Bootetix argentatus 251 Calliptaminae  253, 268, 269 Catantopinae  268, 269 Copiocerinae 269 Coptacridinae 268 Cyrtacanthacridinae  253, 268, 269 Egnatiinae 268 Eremogryllinae 268 Euryphyminae 269 Eyprepocnemidinae 269 Gomphocerinae  251, 252, 253, 268, 269 Habrocneminae 269 Hemiacridinae  253, 269 Kosciuscola 251 Leptysminae  253, 269 Marellia remipes 250 Marelliinae 269 Melanoplinae  250, 268, 269

Melanoplus 269 Melanoplus sanguinipes 807 Oedipodinae  252, 253, 268, 269 Ommatolampidinae  251, 269 Oxyinae  250, 251, 269 Pauliniinae 269 Paulininae 250 Proctolabinae  251, 269 Rhytidochrotinae 269 Schistocerca 753 Schistocerca ceratiola 251 Schistocerca gregaria 253 Spathosterninae 269 Teratodinae 269 Tropidopolinae 269 Urnisiella rubropunctata 250 Acridoidea  874, 876, 880, 886 Anostostomatidae  69, 251, 252, 253, 256, 258, 260, 261 Hemideina maori 253 Caelifera 262–266 Charilaidae (see Pamphagodidae) Chorotypidae  69, 256, 265 Chininae 265 Chorotypinae 265 Erianthinae 265 Eruciinae 265 Mnesicleinae 265 Prioacanthinae 265 Coolooidae (see Anostostomatidae) Cylindrachetidae 250, 256, 263 Dericorythidae  69, 257, 268, 269 Conophyminae 269 Dericorys albidula 269 Dericorythinae 269 Iranellinae 269 Ensifera  254–262, 884, 885, 886, 888 Episactidae  256, 264, 265 Episactinae 265 Espagnolinae 265 Miraculinae 265 Pielomastax 265 Teicophryinae 265 Eumastacidae  69, 256, 265, 266 Eumorsea 266 Gomphamastacinae 266 Gomphomastacinae 265

927

928

Index

Orthoptera: Eumastacidae (contd.) Moreseinae 266 Morsea 266 Psychomastax 266 Euschmidtiidae  256, 265, 266 Euschmidtiinae 266 Pseudoschmidtiinae 266 Stenoschmidtiinae 266 Gryllacrididae  69, 251, 256, 258, 260, 261 Gryllacridinae 261 Lezininae 261 Gryllidae  69, 73, 255, 256, 258, 258, 683, 686, 687 Acheta domestica 254 Caconemobius fori  141 Caconemobius sandwichensis  141 Caconemobius varius  141 Eneopterinae 255 Euscyrtinae 255 Gryllinae 255, 258 Gryllomiminae 255 Gryllomorphinae 255 Gryllus bimaculatus 254 Hapithinae 255 Itarinae 255 Landrevinae 255 Luzarinae 255 Nemobiinae 255 Oecanthinae 255 Paragryllinae 255 Pentacentrinae 255 Phalangopsinae 255, 258 Phaloriinae 255 Podoscirtinae 255 Pteroplistinae 255 Sclerogryllinae 255 Trigonidiinae 255 Gryllotalpidae  69, 250, 255, 256, 258, 258, 873, 882 Gryllotalpinae 258 Malgasia 258 Malgasiinae 258 Mogoplistinae 258, 684 Scapteriscinae 258 Lathiceridae 250, 257, 268, 269 Lentulidae 253, 257, 268, 269, 270 Lentulinae 269

Shelforditinae 269 Lithidiidae  257, 268, 269 Mastacideidae  256, 265, 266 Metrodoridae  69 Mogoplistidae  69, 255, 256, 258 Morabidae 253, 256, 265, 266 Biroellinae 266 Morabinae 266 Myrmecophilidae  69, 255, 256, 258 Ommexechidae  257, 268, 270 Aucaridinae 270 Clarazella 270 Illapeliinae 270 Ommexecha 270 Ommexechinae 270 Pamphagidae 252, 257, 264, 268, 270 Akicerinae 270 Echinotropinae 270 Pamphaginae 270 Porthetinae  252, 270 Prionotropis hystrix rhodanica 270 Thrinchinae 270 Pamphagodidae  257, 268, 270 Pamphagodes riffensis 270 Phaneropteridae (see Tettigoniidae—Phaneropterinae) Pneumoridae 252, 257, 264, 267 Bullacris membracioides 252 Prophalangopsidae  256, 258, 260 Aboilomimus 260 Cyphoderrinae  258, 260 Cyphoderris 260 Parachyphoderris 260 Prophalangopsinae 260 Prophalangopsis 260 Tarragoilus 260 Proscopiidae  256, 264, 266 Hybusinae 266 Proscopiinae 266 Xeniinae 266 Pyrgacrididae  257, 268, 270 Acanthophoenix 270 Pyrgacris descampsi 270 Pyrgacris relictus 270 Pyrgomorphidae  69, 251, 253, 257, 264, 267, 268, 270 Orthacridinae 268

Index

Pyrgomorphinae 268 Sphenarium purpurascens purpurascens 254 Rhaphidophoridae  69, 73, 250, 251, 256, 258, 260, 870 Aemodogryllinae 260 Anoplophilinae 260 Ceuthophilinae 260 Ceuthophilus 260 Dolichopodainae 260 Gammarotettiginae 260 Macropathinae 260 Rhaphidophorinae 260 Troglophilinae 260 Trogophilus 870 Tropidischiinae 260 Ripipterygidae 250, 256, 263 Romaleidae  257, 268, 270 Bactrophorinae  251, 271 Romaleinae 271 Titanacris 271 Tropidacris 271 Schizodactylidae  256, 258, 258 Comicinae 258 Comininae  258 Schizodactylinae 258 Schizodactylus 258 Stenopelmatidae  251, 252, 256, 258, 260, 261 Oryctopinae 261 Siinae 261 Stenopelmatinae 261 Tanaoceridae  256, 264, 266, 267 Tetrigidae  69, 250, 251, 256, 263, 264, 265 Batrachideinae  263, 265 Cladonotinae 265 Lophotettiginae 265 Metrodorinae 265 Scelimeninae 250 Tetriginae 265 Tettigoniidae  69, 73, 247, 251, 253, 256, 258, 261, 262, 683, 686, 687, 882, 885 Neduba extincta (extinct)  90 Phaneropterinae 262 Aganacris 253 Austrosaginae 262 Bradyporinae 262 Cedarbergeniana imperfecta 251

Conocephalinae  258, 262 Hexacentrinae 262 Lichenodraculus 253 Listroscelidinae  251, 262 Markia 253 Meconematinae 262 Mecopodinae 262 Microtettigoniinae 262 Phaneropterinae  253, 262 Phasmodinae  251, 262 Phyllophorinae 262 Polyancistrinae 262 Pseudophyllinae  258, 262 Pterochrozinae 253, 258, 262 Pterochrozini 262 Saginae  251, 262 Segestidea novaeguineae 688 Sexava 693 Tettigoniinae 262 Tympanophorinae 262 Typophyllum 253 Zaprochilinae  251, 262 Thericleidae  256, 264, 265, 266 Tridactylidae  69, 250, 256, 263, 264, 675, 684, 692 Laupala 88 Trigonopterygidae  257, 264, 267 Borneacridinae 267 Trigonopteryginae 267 Tristiridae  257, 268, 270 Atacamacridinae 271 Tristirinae 271 Xyronotidae 252, 257, 264, 266, 267

p

Phasmatodea 61, 69, 73, 202, 210, 219, 235, 245, 246, 281–304 (chapter 11), 406 Agathameridae Agathemera  289, 290, 294 Agathemerinae  285 Anisacanthidae  285, 295 Anisacanthini 292 Archipseudophasmatidae 302 Aschiphasmatidae 291 Aschiphasmatinae  285, 290, 291, 294, 300 Aschiphasmatini 290 Dajaca  290, 291, 294

929

930

Index

Phasmatodea: Aschiphasmatidae (contd.) Dajacini 290 Dallaiphasma 290 Bacillidae Antongiliinae  285, 295 Bacillinae  284, 285, 295 Bacillus  295, 303 Bacillus rossius  284 Clonopsis  295, 303 Macynia 295 Macyniinae  285 Phalces 295 Xylica 295 Damasippoididae  294, 295 Damasippoidinae  285 Diapheromeridae  69 Alienobostra brocki  284 Bacteria  293, 294 Bactrododema  294, 297 Bactrododema hippotaurum 294 Clonaria  295, 297 Diapheromera  290, 294, 303 Diapheromera femorata 303 Diapheromerinae  284, 285, 290, 293, 294, 296 Diapheromerini 294 Diesbachia tamyris  284 Gratidia 284, 295 Gratidiini  284, 285, 295, 296, 297 Leptynia  295, 303 Marmessoidea rosea  284 Megaphasma  290, 293 Micrarchus hystriculeus 303 Necrosciinae  284, 285, 295, 297, 299, 300 Neohirasea 300 Ocnophilini 294 Ocnophiloidea 293 Oreophoetini 294 Pachymorpha 297 Pachymorphinae 297 Palophinae  285, 294, 295, 297 Phaenopharos 300 Pijnackeria 295 Pseudosermyle 294 Pterolibethra 294 Sceptrophasma 284, 295, 297 Sceptrophasma hispidulum  284

Sipyloidea  297, 300 Sipyloidea sipylus  284, 300 Trachythorax maculicollis  284 Heteronemiidae Creoxylus spinosus  284 Heteronemiinae  285, 290, 294 Parabacillus 290 Spinonemia 290 Heteropterygidae 292, 293 Dataminae  293 Datamini 292 Epidares 292 Haaniella 292 Heteropteryginae  284, 285, 292, 293, 293 Heteropterygini 292 Heteropteryx  292, 293 Miroceramia 292 Obriminae 292, 293 Obrimini 292 Pterobrimus 292 Sungaya inexpectata  284 Necrosciidae 300 Phasmatidae  69, 289 Acanthoxyla 282, 303 Acanthoxyla inermis  282 Achriopterini  285, 295 Acrophylla 297 Anchiale austrotessulata 303 Apterograeffea 299 Argosarchus 303 Baculofractum 299 Baculofractum insigne  284 Canachus 298 Carausius morosus  282, 299 Carlius 297 Cladomorphinae  285, 294, 296, 297 Clitarchus 303 Clitarchus hookeri  302, 303 Clitumninae  284, 296 Clitumnini  285, 295, 296 Cnipsus 297 Diapherodes gigantea 296 Didymuria violescens  284, 303 Dimorphodes 299 Dryococelus 298, 298 Dryococelus australis  298, 298 Epicharmus 299

Index

Eurycantha 298, 299 Eurycantha calcarata  284 Eurycantha horrida  298 Eurycanthinae 297, 298, 298 Eurycnema 284 Extatosoma 303 Graeffea  299, 303 Graeffea crouanii 303 Hesperophasmatini 297 Leosthenes 299 Leprocaulinus  282, 299 Lobofemora 296 Lonchodinae  282, 284, 285, 297, 298, 298, 299, 300 Macrophasma 282 Malandania 283, 297, 299 Malandania pulchra  283 Medaurini  285, 295, 296 Megacrania 299 Neopromachus 297 Otocrania 293 Parapachymorpha spiniger  284 Pharnaciini  284, 285, 296 Phasmatinae 297 Phobaeticus 282, 284, 296 Phobaeticus chani  282, 296 Phobaeticus kirbyi 296 Phobaeticus serratipes  284 Platycraninae  297, 299 Podacanthus wilkinsoni 303 Ramulus 296 Ramulus nematodes 296 Rhamphophasma spinicorne  284 Rhaphiderus 299 Spathomorpha  285, 295 Staelonchodes harmani  284 Stephanacridini  285, 296, 297 Tainophasma 296 Tropidoderinae  284, 297, 299 Tropidoderus  297, 299 Xeroderinae  297, 299 Xeroderus 299 Phylliidae  69, 289, 291 Chitoniscus 292 Eophyllium messelense (extinct)  291, 300 Nanophyllium 292 Phylliinae  283, 284, 285, 291, 292, 297

Phyllium 283, 291, 292 Phyllium bioculatum  283 Phyllium giganteum  284 Prisopodidae 294 Kalocorinnis 300 Korinninae  289, 291, 294, 300, 301 Korinnis 300 Prisopodinae 300 Prisopus  294, 300 Prispopodidae 294 Pseudophasmatidae  69, 284, 285, 294, 297 Anisomorpha paromalus  284 Melophasma 294 Susumanioidea (extinct)  302 Timematidae 286 Timema  285, 286, 288, 289, 290, 302, 303 Timema cristinae  289, 302 Phasmatoptera. See Phasmatodea Phasmida. See Phasmatodea Phthiraptera 19, 61, 69, 73, 210, 417, 422, 434, 447, 458–465, 637, 886, 891 Boopiidae  458, 459 Echinophthiriidae  19, 458, 463 Antarctophthirus microchir 463 Latagophthirus rauschi 463 Enderleinellidae  69, 458 Microphthirus uncinatus 457 Gyropidae  458, 459 Haematomyzidae  458 Haematopinidae  458 Haematopinus suis  459, 460 Haemodipsus setoni 463 Hamophthiriidae  458 Hoplopleuridae  19, 69, 458 Hybophthiridae  458 Laemobothriidae  458 Laemobothrion vulturis 457 Linognathidae  19, 458 “Mallophaga”  41, 460–463 Menoponidae  19, 69, 458, 477 Actornithophilus 461 Colpocephalum 461 Hohorstiella lata 464 Myrsidea 477 Piagetiella 462 Piagetiella peralis 462, 463

931

932

Index

Phthiraptera: Menoponidae (contd.) Pseudomenopon pilosum 474 Trinoton anserinum 474 Microthoraciidae  458 Neolinognathidae  458 Pecaroecidae  458 Pedicinidae  69, 458 Pediculidae  19, 458 Pediculus 869 Pediculus humanus capitis  457, 473 Pediculus humanus humanus 473 Philopteridae  19, 69, 458, 459, 461 Brueelia  458 Campanulotes compar 464 Coloceras tovornikae 464 Columbicola 461 Columbicola columbae 464 Craspedorrhynchus 461 Haffneria 461 Ornithobius 461 Pectinopygus 461 Pectinopygus farallonii  457 Philopterus  458 Rotundiceps 460 Strigiphilus 461 Polyplacidae  19, 69, 458 Pthiridae  458 Pthirus pubis  459, 473 Ratemiidae  458 Rhynchophthirina 457, 458, 459, 461 Ricinidae  19, 458, 461 Ricinus 464 Trichodectidae  19, 69, 458, 459 Neotrichodectes 461 Thomomydoecus 461 Trichodectes euarctidos  462 Trimenoponidae  458, 459 Phylloptera. See Phasmatodea–Phylliidae Plecoptera 18, 19, 23, 24, 27, 28, 31, 34, 61, 69, 74, 77, 203, 210, 397, 734, 886, 891 Capniidae  19 Chloroperlidae  19 Paraperla frontalis  127, 133 Leuctridae  69 Nemouridae  19, 69 Nemoura arctica 38 Amphinemura palmeni 31

Peltoperlidae  69 Perlidae  19, 69, 74 Perlodidae  19 Pteronarcidae  19 Styloperlidae  69 Protelytroptera (extinct)  324 Psocodea. See Pthiraptera and Psocoptera Psocoptera 23, 25, 61, 69, 74, 199, 210, 417–448 (chapter 16), 484, 637 Amphientomidae  69, 420, 423, 426, 428, 431, 436, 436 Lithoseopsis 437 Lithoseopsis hellmani  436 Marcenendius 437 Nephax 437 Seopsocus 437 Seopsocus rafaeli  420, 423, 426 Stimulopalpus japonicus 437 Amphientomoidea  428, 431, 436 Amphipsocidae  420, 426, 427, 429, 432, 439 Afropsocus 439 Amphipsocus 439 Brachypsocus badonneli 439 Capillopsocus 439 Complaniamphus 439 Ctenopsocus 439 Harpezoneura 439 Kolbia quisquiliarum 439 Pentathyrsus 439 Polypsocus 439 Polypsocus corruptus 439 Polypsocus lineatus  420, 426, 427 Pseudokolbea 439 Schizopechus 439 Siniamphipsocus 439 Taeniostigma 439 Tagalopsocus 439 Xenopsocus 439 Archipsocidae  420, 422, 427, 428, 432, 438 Archipsocopsis 438 Archipsocus 438 Archipsocus indentatus  427 Notarchipsocus 438 Notarchipsocus fasciipennis  420 Pararchipsocus 438 Pseudarchipsocus 438 Asiopsocidae  421, 423, 427, 429, 432, 439

Index

Asiopsocus 439 Asiopsocus sonorensis  423 Notiopsocus 421, 423, 427, 439 Pronotiopsocus 427, 439 Bryopsocidae (see Philotarsidae) Calopsocidae (see Philotarsidae) Caeciliusidae  69, 421, 422, 424, 426, 429, 432, 439, 440, 443 Caecilius  440, 441 Caecilius fuscopterus 440 Caeciliusini 441 Maoripsocini 441 Maoripsocus 441 Valenzuela 440 Valenzuela postica  421, 424, 426 Cladiopsocidae  421, 429, 432, 444 Cladiopsocus 421, 444 Spurostigma 421, 444 Compsocidae  420, 424, 427, 431, 437 Compsocus 437 Electrentomopsis 437 Electrentomopsis variegata  420, 424, 427 Dasydemellidae  425, 427, 429, 432, 436, 439 Dasydemella 439 Matsumuraiella 439 Ptenopsila 439 Teliapsocus conterminus  425, 427, 436, 439 Dolabellopsocidae  425, 429, 432, 444 Auroropsocus 444 Dimidistriata 444 Dolabellopsocus 444 Dolabellopsocus similis  425 Isthmopsocus 444 Ectopsocidae  421, 422, 425, 429, 432, 436, 441 Ectopsocopsis cryptomeriae 441 Ectopsocus 447 Ectopsocus briggsi  421, 425 Ectopsocus meridionalis  436 Ectopsocus pumilis 441 Ectopsocus richardsi 441 Electrentomidae  419, 420, 423, 431, 437 Electrentomum 437 Epitroctes 437 Epitroctes tuxtlarum  419, 420, 423, 424 Manicapsocus 437 Nothoentomum 437

Parelectrentomum 437 Phallopsocus 437 Electrentomoidea  428, 431, 436, 437 Elipsocidae  419, 421, 422, 424, 429, 432, 441, 442 Cuneopalpus 442 Elipsocus 442 Elipsocus guentheri  424 Elipsocus obscurus  421 Hemineura 442 Nepiomorpha peripsocoides  419, 421 Propsocinae 442 Pseudopsocus 442 Reuterella 442 Epipsocidae  69, 419, 421, 423, 429, 433, 444, 445 Bertkauia 445 Bertkauia crosbyana  419, 421 Epipsocopsis 445 Epipsocus 423, 445 Epipsocus foliatus  423 Goja 445 Mesepipsocus 445 Neurostigma 445 Hemipsocidae  421, 424, 430, 433, 445 Cyclohemipsocus 446 Hemipsocus 446 Hemipsocus chloroticus  421, 424, 446 Metahemipsocus 446 Lachesillidae  69, 421, 422, 424, 427, 429, 432, 436, 441, 442 Lachesilla 442 Lachesilla contraforcepeta  436 Lachesilla riegeli  421 Lachesilla tropica  424, 427 Lepidopsocidae  419, 420, 421, 425, 427, 428, 431 Neolepolepis occidentalis  419, 421 Nepticulomima 420, 427 Lesneiidae  429, 442 Lesneia 442 Liposcelididae 417, 419, 428, 431, 434, 435 Belapha 435 Belaphopsocus 435 Belaphotroctes 435 Chaetotroctes 435 Embidopsocinae 435

933

934

Index

Psocoptera: Liposcelididae (contd.) Embidopsocopsis 435 Embidopsocus 423, 435 Embidopsocus bousemani  419 Embidopsocus needhami  419 Liposcelidinae 435 Liposcelis  435, 447 Liposcelis bostrychophila  419 Liposcelis decolor 422 Liposcelis entomophila  419 Lipsoelis 435 Troctulus 435 Troglotroctes 435 Mesopsocidae 422, 424, 426, 427, 429, 432, 441, 442 Mesopsocopsis 443 Mesopsocus 442 Mesopsocus unipunctatus  424, 426, 427 Newipsocus termitiformis 442 Musapsocidae  421, 423, 425, 431, 437, 438 Musapsocoides 438 Musapsocus 438 Musapsocus creole  421 Musapsocus huastecanus  423, 425 Myopsocidae  422, 427, 430, 433, 445, 446 Lichenomima 446 Lichenomima lugens  422, 427 Lophopterygella 446 Myopsocus 446 Pachytroctidae  419, 421, 424, 426, 428, 431, 434, 435 Nanopsocus 435 Nanopsocus oceanicus  421, 447 Pachytroctes 435 Peritroctes 435 Tapinella 435 Tapinella maculata  419, 424, 426 Paracaeciliidae  421, 424, 426, 429, 432, 439, 440 Enderleinella 440 Paracaecilius 440 Xanthocaecilius 440 Xanthocaecilius quillayute 440 Xanthocaecilius sommermanae  421, 424, 426, 440 Peripsocidae  421, 422, 424, 425, 429, 432, 436, 441

Kaestneriella 441 Peripsocus 441 Peripsocus stagnivagus  421, 424, 425 Peripsocus subfasciatus  436 Philotarsidae  69, 421, 422, 424, 426, 427, 429, 432, 443 Aaroniella 443 Abelopsocus 443 Garcialdretia 443 Haplophallus 443 Philotarsopsis 443 Philotarsus 443 Philotarsus arizonicus  421, 424, 426, 427 Tarsophilus 443 Prionoglarididae  420, 422, 424, 428, 431, 434 Speleketor flocki  424 Speleketor irwini  420 Protroctopsocidae  420, 428, 431, 437 Chelyopsocus 437 Philedaphia 437 Protroctopsocus 437 Protroctopsocus enigmaticus  420 Reticulopsocus 437 Pseudocaeciliidae  69, 422, 424, 426, 427, 429, 432, 443, 444 Heterocaecilius 424 Ophiodopelma 447 Pseudocaecilius citricola  426, 427 Scottiella 443 Scytopsocus 443 Trimerocaecilius 443 Psilopsocidae  421, 423, 424, 430, 445 Psilopsocus mimulus 445 Psilopsocus nebulosus  421, 423, 424 Psocidae  69, 419, 422, 424, 425, 427, 430, 433, 436, 445, 446 Amphigerontia 425, 446 Amphigerontia bifasciata  424, 425 Amphigerontiinae 446 Blaste 446 Camelopsocus 447 Camelopsocus bactrianus  419, 425 Cerastipsocini 446 Cerastipsocus 446 Hyalopsocus 436 Indiopsocus 447 Loensia 447

Index

Loensia conspersa  422 Loensia fasciata  422 Metylophorini 447 Neopsocus 447 Psocinae 446 Psocini 447 Psococerastis 447 Psococerastis fasciata  422, 427 Ptycta 447 Ptyctini 447 Steleops 447 Steleops elegans  422 Steleops lichenatus  422 Thyrsophorini 447 Trichadenotecnum 447 Trichadenotecnum quaesitum  422 Psoquillidae  419, 421, 424, 425, 426, 426, 428, 431 Psoquilla marginepunctata  421, 424, 426, 434 Rhyopsocus 434 Rhyopsocus bentonae  419 Rhyopsocus texanus  421 Psyllipsocidae  69, 420, 424, 427, 428, 431, 434 Dorypteryx 434 Pseudorypteryx 434 Psocathropos 434 Psyllipsocus 434 Psyllipsocus huastecanus  420, 427 Psyllipsocus maculatus  424 Psyllipsocus ramburii 434 Ptiloneuridae  420, 423, 433, 444 Loneura 445 Loneura splendida  420, 423 Sabulopsocidae  429, 442 Sphaeropsocidae  419, 424, 426, 427, 428, 431, 434, 435 Badonnelia titei 435 Prosphaeropsocus pallidus  424 Sphaeropsocopsis argentina  426, 427 Sphaeropsocus bicolor 435 Sphaeropsocus kuenowii 435 Troglosphaeropsocus voylesi  419 Spurostigmatidae (see Cladiopsocidae) Stenopsocidae  69, 421, 424, 426, 427, 429, 432, 436, 439, 440

Cubipilis 440 Graphopsocus 440 Graphopsocus cruciatus  436, 440 Malostenopsocus 440 Stenopsocus 424, 440 Stenopsocus nigricellus  421, 426, 427 Trichopsocidae  421, 422, 424, 427, 429, 432, 443, 444 Palaeopsocus 444 Trichopsocus 444 Trichopsocus clarus  421, 424, 427 Troctopsocidae  428, 431, 437 Coleotroctellus 438 Selenopsocus 438 Thaipsocus 438 Troctopsocoides 438 Troctopsocopsis 438 Troctopsoculus 438 Troctopsocus 438 Trogiidae  419, 424, 425, 426, 427, 428, 431 Cerobasis 426 Cerobasis annulata 447 Cerobasis lineata  419, 424, 427 Lepinotus 426 Lepinotus inquilinus 447 Lepinotus patruelis 447 Lepinotus reticulatus  419 Trogium 426 Trogium pulsatorium 447

r

Raphidioptera 61, 69, 74, 77, 657 Inocelliidae  69, 74, 631, 638, 641, 654, 657, 658 Inocelliinae  641 Neghini  641 Parainocellia bicolor  654 Mesoraphidiidae (extinct)  69, 74 Raphidiidae  631, 638, 641, 654, 657, 658 Agulla 654 Agullini  641 Alenini  641 Raphidiinae  641 Raphidiini  641

s

Siphonaptera 21, 23, 61, 69, 74, 465–474, 707, 710, 715, 873, 880, 884, 888, 891 Ancistropsyllidae  69, 465

935

936

Index

Siphonaptera (contd.) Ceratophyllidae  21, 69, 465 Ceratophyllinae  465 Ceratophyllus  464, 469, 472 Ceratophyllus arcuegens 467 Ceratophyllus celsus 472 Ceratophyllus gallinae 475 Ceratophyllus niger 475 Ceratophyllus sciurorum 469 Ceratophyllus scopulorum 472 Ceratophyllus vagabundus 41 Dactylopsyllinae  465 Glaciopsyllus antarcticus 469 Mioctenopylla arctica 41 Opisodasys pseudarctomys  468 Orchopeas caedens 475 Chimaeropsyllidae  465 Chiastopsyllinae  465 Chimaeropsyllinae  465 Epirimiinae  465 Coptopsyllidae  465 Ctenophthalmidae  465 Anomiopsyllinae  465 Anomiopsyllus 464, 467 Brachyctenonotini  465 Ctenophthalminae  465 Dinopsyllinae  465 Doratopsyllinae  465 Listropsyllinae  465 Liuopsyllinae  465 Neopsyllinae  465 Rhadinopsyllinae  465 Stenoponiinae  465 Stenoponia  464, 468 Hectopsyllidae Echidnophaga gallinacea  468, 468, 475 Hystrichopsyllidae  69, 465 Corrodopsylla 472 Corrodopsylla curvata 472 Hystrichopsylla 467, 468, 468 Nearctopsylla hygini 472 Ischnopsyllidae  69, 465 Myodopsylla 464 Thaumapsyllinae  465 Leptopsyllidae  21, 69, 465 Amphipsyllinae  465 Leptopsyllinae  465

Peromyscopsylla catatina 472 Lycopsyllidae  465 Bradiopsyllinae  465 Lycopsyllinae  465 Choristopsyllinae  465 Uropsylla tasmanica 470, 471, 472 Uropsyllinae  465 Macropsyllidae 464, 465 Malacopsyllidae  465 Pulicidae  21, 69, 465 Cediopsylla simplex 473 Ctenocephalides felis  468, 472 Euchoplopsyllus glacialis 469 Pulex 464 Spilopsyllus cuniculi  473, 475 Xenopsylla cheopis  472, 474 Pygiopsyllidae  69, 465 Notiopsylla enciari  471 Rhopalopsyllidae  466 Parapsyllinae  466 Parapsyllus longicornis 472 Rhopalopsyllinae  466 Stephanocircidae 464, 466 Craneopsyllinae  466 Stephanocircinae  466 Stephanocircus dasyuri  466 Stivaliidae  69, 466 Farhangiinae  466 Lentistivalius insolli 74 Stivaliinae  466 Tungidae  466 Tunga penetrans  469, 476 Vermipsyllidae  466 Chaetopsylla tuberculaticeps 473 Vermipsylla alakurt 468 Xiphiopsyllidae  466 Strepsiptera 61, 69, 74, 77, 167, 210, 628, 659, 673–694 (chapter 22), 706, 715 Bahiaxenidae  674, 675, 678, 681, 682 Bahiaxenos 678, 682 Bahiaxenos relictus 678 Bohartillidae  674, 681, 683, 690 Bohartilla  683, 691 Bohartilla megalognatha 690 Callipharixenidae  69, 684, 692 Corioxenidae  69, 674, 675, 676, 681, 682, 685 Australoxenos  682, 685

Index

Blissoxenos  682, 685 Corioxenos  682, 685 Dundoxenos  682, 685 Eocenoxenos (extinct)  682, 685 Floridoxenos  682, 685 Loania  682, 685 Malagasyxenos  682, 685 Malayaxenos  682, 685 Mufagaa  682, 685 Proceroxenos  682, 685 Triozocera  682, 685 Uniclavus  682, 685 Viridipromontoxius  682, 685 Cretostylopidae (extinct)  674, 678, 682 Cretostylops (extinct)  682 Dipterophagidae (see Halictophagidae—Dipterophaginae) Elenchidae  69, 674, 675, 684, 684, 688, 691, 693 Colacina  684, 691 Deinelenchus  684, 691, 693 Deinelenchus australensis 692 Elencholax  684, 691 Elenchus 680, 684, 691 Elenchus japonicus 691 Elenchus tenuicornis 691 Elenchus varleyi  681 Elenchus yasumatsui 691 Protelenchus (extinct)  684, 691 Halictophagidae  69, 674, 675, 684, 684, 688, 691, 692, 693 Blattodeaphagus  684, 692 Callipharixenos  684, 692, 693 Callipharixenos philippines 693 Coriophaginae 692 Coriophagus  684, 692 Dipterophaginae 692 Dipterophagus  684, 692 Dipterophagus daci  692, 693 Halictophaginae 692 Halictophagus  684, 692, 693 Halictophagus australiensis 693 Halictophagus calcaratus 693 Halictophagus fulmeki 693 Halictophagus indicus 693 Halictophagus naulti 693 Halictophagus palmae 693

Halictophagus silwoodensis 692 Halictophagus tryoni 692 Stenocranophilus  684, 692 Tridactylophaginae 692 Tridactylophagus  684, 692 Kinzelbachillidae (extinct)  674, 682 Kinzelbachilla (extinct)  682 Lychnocolacidae  674, 675, 681, 683, 688 Lychnocolax  683, 684, 688 Lychnocolax hispanicus 688 Mengeidae (extinct)  674, 678, 682 Mengea (extinct)  682 Mengenillidae 167, 674, 674, 675, 677, 678, 679, 680, 681, 682 Congoxeninae 678 Congoxenos  678, 679, 682 Eoxenos  678, 679, 680, 681, 682 Eoxenos laboulbenei 167, 679, 680, 681, 681 Iberoxerninae 678 Mengenilla 678, 678, 679, 680, 681, 682 Mengenilla chabauti 167 Mengenillinae 678 Trilineatoxenos  678, 679, 682 Yemengenilla  678, 679, 682 Myrmecolacidae  70, 674, 675, 681, 683, 684, 685, 686, 687, 688, 693 Caenocholax  683, 685, 686, 687, 688, 693 Caenocholax fenyesi  680, 688 Kronomyrmecolax (extinct)  683, 685 Kronomyrmecolax fushunicus (extinct)  687 Myrmecolax  683, 685, 686, 687 Myrmecolax incautus 687 Palaeomyrmecolax (extinct)  683, 685 Stichotrema  683, 685, 686, 687, 688 Stichotrema dallatorreanum  686, 688, 693 Phthanoxenidae (extinct)  674, 682 Phthanoxenos (extinct)  682 Protoxenidae (extinct)  674, 678, 682 Protoxenos (extinct)  682 Stylopidae  70, 674, 675, 683, 684, 688, 689, 693 Crawfordia  683, 688 Eurystylops  683, 688 Halictoxenos  683, 688, 689 Hylecthrus  683, 688 Jantarostylops  683, 688

937

938

Index

Strepsiptera: Stylopidae (contd.) Kinzelbachus  683, 688 Melittostylops  683, 688 Rozenia  683, 688 Stylops 677, 683, 688, 689 Stylops pacificus 689 Xenidae  674, 675, 683, 684, 688, 689, 690 Paragioxenos  683, 689 Paraxenos  683, 689, 690 Paraxenos lugubris  681 Pseudoxenos  683, 689, 690 Pseudoxenos carnifax 689 Pseudoxenos hockeri 689 Pseudoxenos iwatai 689 Xenos  683, 689, 690 Xenos hamiltoni 689 Xenos moutoni 690 Xenos oxyodontes 690 Xenos vesparum  677, 679, 680, 689

t

Thysanoptera 20, 61, 70, 74, 210, 483–496 (chapter 18), 637, 804, 805 Aeolothripidae  70, 484, 485, 486, 490 Aeolothrips 486 Cycadothrips albrechti 489 Fauriellidae  485, 486 Hemithripidae  485 Heterothripidae  485 Aulacothrips 487 Karataothripidae  485 Liassothripidae  485 Melanthripidae 484, 485 Merothripidae  70, 485, 486 Moundthripidae  485 Phlaeothripidae  70, 484, 485, 486, 487, 489, 490, 491 Adrothrips intermedius  489 Crotonothrips polyalthiae 494 Dunatothrips 490 Dunatothrips aneurae 490 Elaphrothrips 488 Eurynothrips magnicollis 491, 492 Grypothrips cambagei 492, 493 Haplothrips  487, 488 Haplothrips leucanthemi  484 Idolothripinae  485, 490

Idolothrips spectrum 490 Karnyothrips flavipes 487 Karnyothrips melaleucus 486 Kladothrips  487, 490 Kladothrips sterni  491 Koptothrips 487 Liothrips  487, 488 Nesothrips lativentris 495 Phlaeothripinae  485 Stenurothripidae  485, 486 Holarthrothrips = Adiheterothrips 486 Thripidae  70, 484, 485, 486, 487, 489, 490, 491 Anaphothrips 488 Arachisothrips 491, 492 Caliothrips fasciatus 493 Chirothrips  487, 495 Craspedothrips 486 Dendrothripinae  485 Echinothrips americanus 495 Frankliniella  488, 491, 495 Frankliniella occidentalis  486, 490, 494 Frankliniella schultzei 486 Limothrips 487 Limothrips cerealium 493 Mycterothrips 487 Panchaetothripinae  485, 491 Parthenothrips 491 Parthenothrips dracaenae 491 Scirtothrips dorsalis 495 Scolothrips 486 Sericothripinae  485 Taeniothrips inconsequens 494 Thripinae  485 Thrips  488, 491 Thrips palmi  490, 495 Thrips tabaci  486, 494 Triassothripidae (extinct)  485 Uzelothripidae  485, 486 Thysanura. See Microcoryphia, Zygentoma and Monura  155–187 (chapter 7) Trichoptera  4, 18, 22, 23, 24, 27, 28, 34, 37, 61, 70, 74, 77, 886 Apataniidae  70 Apatania zonella 37 Brachycentridae  22, 70 Calamoceratidae  70

Index

Dipseudopsidae  70 Ecnomidae  70 Glossosomatidae  22, 70 Goeridae  70 Helicopsychidae  70 Hydrobiosidae  70 Hydropsychidae  70 Hydroptilidae  22, 70 Lepidostomatidae  70 Leptoceridae  22, 70 Limnephilidae  22, 70 Limnocentropodidae  70 Molannidae  70 Odontoceridae  70 Philopotamidae  70 Phryganeidae  22, 70 Phryganopsychidae  70 Polycentropodidae  70 Psychomyiidae  70 Ptilocolepidae  70 Rhyacophilidae  22, 70 Sericostomatidae  70 Stenopsychidae  70 Uenoidae  70 Xiphocentronidae  70

z

Zoraptera 61, 70, 74, 199–212 (chapter 8), 637 Zorotypidae  70, 199 Xenozorotypus 199 Xenozorotypus burmiticus (extinct)  210 Zorotypus 199 Zorotypus absonus  210 Zorotypus acanthothorax  210 Zorotypus amazonensis  210 Zorotypus barberi  202, 203, 206, 207, 209, 209, 210 Zorotypus brasiliensis  199, 202, 210 Zorotypus buxtoni  210 Zorotypus caudelli  202, 203, 206, 207, 210 Zorotypus caxiuana  210 Zorotypus cervicornis  210 Zorotypus ceylonicus  210 Zorotypus congensis  210 Zorotypus cramptoni  210 Zorotypus cretatus  210

Zorotypus delamarei  203, 209, 210 Zorotypus goeleti (extinct)  203, 210 Zorotypus guineensis  210 Zorotypus gurneyi  203, 206, 207, 209, 209, 210 Zorotypus hainanensis  210 Zorotypus hamiltoni  210 Zorotypus hubbardi  199, 200, 202, 203, 204, 206, 210, 210 Zorotypus hudae (extinct)  199, 210 Zorotypus huxleyi  202, 208, 210 Zorotypus impolitus  202, 203, 205, 207, 210 Zorotypus javanicus  210 Zorotypus juninensis  210 Zorotypus lawrencei  210 Zorotypus leleupi  210 Zorotypus longicercatus  210 Zorotypus magnicaudelli  202, 203, 207, 210 Zorotypus manni  210 Zorotypus medoensis 199, 210, 210 Zorotypus mexicanus  210 Zorotypus mnemosyne  210 Zorotypus nascimbenei  210 Zorotypus neotropicus 203, 210 Zorotypus newi  210 Zorotypus novobritannicus  210 Zorotypus palaeus  210 Zorotypus philippinensis  210 Zorotypus sechellensis  210 Zorotypus shannoni  202, 208, 210 Zorotypus silvestrii  210 Zorotypus sinensis 199, 210 Zorotypus snyderi  199, 203, 210 Zorotypus swezeyi  210 Zorotypus vinsoni  210 Zorotypus weidneri  202, 203, 208, 210 Zorotypus weiweii  210 Zorotypus zimmermani  210 Zygentoma  61, 73, 144, 160, 175–183, 675, 766 incertae sedis Onycholepisma arizonae (extinct)  165, 166 Lepidotrichidae (extinct)  159, 165, 175, 180, 181, 186 Lepidotrix pilifera (extinct)  181 Ramsdelepidion (extinct)  159

939

940

Index

Zygentoma (contd.) Lepismatidae  68, 159, 165, 166, 167, 173, 175, 176, 177, 177, 178, 179, 180, 181, 182, 183, 184, 185, 186, 678, 682 Acrotelsa  160, 178 Acrotelsa collaris 184 Acrotelsatinae  159, 182 Acrotelsella  160 Afrolepisma  160, 186 Afrolepisma leleupi 168 Afrolepisma wygodzinskyi 186 Allacrotelsa  160, 178, 186 Allacrostela burmiticus (extinct)  165 Allacrotelsa dubia (extinct)  165, 186 Allacrotelsa kraepelini 186 Allacrotelsa spinulata 186 Anallacrotelsa  160 Anisolepisma  160, 182 Apteryskenoma  160 Archeatelura (extinct)  165, 186 Asiolepisma  160 Ctenolepisma  160, 167, 176, 178, 179, 183, 186, 678 Ctenolepisma ciliata 167 Ctenolepisma diversisquamis 186 Ctenolepisma electrans (extinct)  165, 166 Ctenolepisma kashinicum (extinct)  165 Ctenolepisma lineata  167, 184 Ctenolepisma longicaudata  177, 180, 184, 186 Ctenolepisma nicoleti 167 Ctenolepisma rothschildi  184, 186 Ctenolepisma targioniana 184 Ctenolepisma targionii 167 Ctenolepismatinae  159, 182 Desertinoma  160 Gopsilepisma  160 Grassiellini  165, 186 Hemikulina  160 Hemilepisma  160 Heterolepisma  160, 186 Heterolepisma bisetosa 180 Heterolepismatinae 181 Hyperlepisma  160 Lampropholis burmiticus = Allocrostelsa burmiticus 165 Lepisma  160, 167, 183, 185, 186

Lepisma aurea 681 Lepisma polypoda 155 Lepisma saccharina  155, 167, 176, 184, 185 Lepisma terrestris 155 Lepismatinae  159, 181, 183, 186 Lepismina  160 Lepitrochisma  160 Leucolepisma  161 Mirolepisma  160 Mirolepismatinae 182 Monachina  161 Mormisma  161 “Mormisma” wygodzinskyi  161 Namibmormisma  161 Namunukulina  160 Namunukulina funanbuli 186 Nebkhalepisma  161 Neoasterolepisma  160, 176, 177, 178, 678 Neoasterolepisma palmonii 167 Neoasterolepisma santschi 167 Neoasterolepisma wasmannii 186 Ornatilepisma  161 Panlepisma  161 Paracrotelsa  160 Primacrotelsa  160 Prolepismina  160, 186 Protograssiella 186 Protolepisma tainicum (extinct)  165 Psammolepisma  161 Sabulepisma  161 Sceletolepisma 183 Silvestrella  160 Silvestrellatinae  159, 182 Stylifera  161, 186 Stylifera galapagoensis 186 Stylifera impudica 186 Swalepisma  161 Thermobia  161, 176, 177, 183 Thermobia domestica 184 Tricholepisma  160, 178, 678 Xenolepisma  160 Machilidae Pedetontus  158, 169, 186 Maindroniidae  159, 180, 181, 186 Maindronia  161, 186 Maindronia beieri 186 Maindronia mascatensis 186

Index

Maindronia neotropicalis 186 Meinertellidae Allomachilis  158, 185 Machiloides  158, 170, 185 Nicoletiidae  68, 159, 165, 166, 166, 167, 176, 177, 178, 179, 180, 182, 183, 184, 186 Acanthinonychia  161 Acanthonima  163 Allatelura  161 Allograssiella  161 Allomorphura  161 Allomorphuroides  161, 176 Allonicoletia  163, 176 Allonychella  161 Allotrichotriura  164 Allotrinemurodes  164 Anarithmeus  161 Anelpistina  163, 178 Anelpistina quinterensis 184 Anelpistina specusprofundi 180 Arabiatelura  161 Archeatelura sturmi 165, 166 Assmuthia  161 Atelura  161 Atelura formicaria  179, 180 Atelurina  162 Ateluridae 181 Atelurinae  159, 165, 175, 176, 177, 179, 180, 182, 183, 185, 186 Atelurodes  162 Ateluropsis  162 Atopatelura  162 Attatelura  162 Ausallatelura  162 Australiatelura  162 Bharatatelura  162 Canariletia  163 Cephalocryptina  162 Coletinia  164, 183 Coletiniinae  159, 182 Comphotriura  162 Congoatelura  162 Cryptocephalina  162 Cryptostylea  162 Crypturella  162 Crypturelloides  162 Cubacubaninae  159, 176, 179, 182, 183, 184

Dinatelura  162 Dinatelurini 180 Dionychella  162 Dodecastyla  162, 186 Ecnomatelura  162 Eluratinda  162 Galenatelura  162, 175 Gastrotheellus  162 Gastrotheus  162 Goiasatelura  162 Grassiella  162, 180, 185, 186 Gynatelura  162 Hematelura  164, 180 Hematelura doriae 180 Hemitrinemura  164 Hemitrinemura extincta (extinct)  165 Heterolepidella  162 Heteromorphura  162 Heteronychella  162 Lasiotheus  162 Lasiotheus nanus 186 Lepidina  164 Lepidospora  164, 179 Lepidotriura  162 Linadureta  162 Luratea  162 Machadatelura  162 Malayatelura  162 Mesonychographis  162 Metrinura  164, 186 Metriotelura  162 Natiruleda  163 Neatelura  163 Nicoletia  163, 168 Nicoletia phytophila  184, 186 Nicoletiinae  159, 182, 183 Nipponatelura  163 Nipponatelurina  163 Olarthrocera  163 Olarthroceroides  163 Onychomachilis fischeri  165, 166 Paleograssiella chiapanicum (extinct)  165 Pauronychella  163 Petalonychia  163 Platystylea  163 Principella  163 Proatelura  163

941

942

Index

Zygentoma: Nicoletiidae (contd.) Proatelura pseudolepisma 167 Proatelurina  163 Prosthecina  163, 179 Protonychella  163 Pseudatelura  163 Pseudatelurodes  163 Pseudobrinckina  164 Pseudogastrotheus  163, 186 Rasthegotus  163 Rulenatida  163 Santhomesiella  163, 178 Speleonycta  163, 180 Squamatinia  164 Squamatinia algharbica 180 Squamigera  163 Squamigera latebricola 180 Subnicoletia  164 Subnicoletiinae  159, 182, 186 Subtrinemura  164 Texoreddellia  163, 180, 184

Trichatelura  164, 179 Trichodimeria  163 Trichotriura  164 Trichotriurella  164 Trichotriuroides  164 Trinemura  164 Trinemurodes  164 Trinemurodes antiquus (extinct)  165 Trinemurodes miocenicus (extinct)  165 Troglotheus  163 Wygodzincinus  163 Protrinemuridae  68, 159, 176, 177, 180, 181, 182, 183, 186 Protrinemura  161, 186 Protrinemurella  161 Protrinemuroides  161, 176 Trinemophora  161, 186 Tricholepidiidae  159, 175, 177, 180, 181, 183, 186 Tricholepidion  161, 183 Tricholepidion gertschi  181, 186

943

Index of Arthropod Taxa Arranged Alphabetically. Note: Page numbers in bold indicate table entries, and numbers in italic face indicate entries on figures and in figure captions.

a Aaroniella 443 Abelopsocus 443 Aboilomimus 260 Acanalonia 537 A. conica  531, 537 Acanaloniidae  64, 504, 531, 534, 535, 536, 537, 547, 548 Acanthaclisini  640 Acanthacorydalis 642 Acanthinonychia  161 Acanthonima  163 Acanthophoenix 270 Acanthopidae  393, 395, 396, 397, 404 Acanthops  395, 396 Acanthoxyla 282, 303 A. inermis  282 Acarna 546 Acari  167, 733, 752 Acarnini 546 Acartophthalidae  21 Acheta domestica 254 Achilidae  64, 504, 531, 534, 535, 537, 540 Achilinae  504, 537 Achilixiidae  504, 531, 534, 535, 537 Achilixiinae  504 Achilixius 537 Achriopterini  285, 295 Aclastus borealis 41

Aclerdidae  612 Aconophora 526, 528 Aconophorini 527, 528 Acontista 390 Acontistidae  393, 395, 396, 397, 404 Acothrura 546 Acrididae  19, 69, 73, 247, 250, 251, 253, 257, 264, 268, 269, 270, 872, 873 Acridinae 268 Acridoidea  874, 876, 880, 886 Acroceridae  63 Acromantinae  393 Acromantini  393 Acrometopum 544 A. panoplites 544 Acrophylla 297 Acrotelsa  160, 178 A. collaris 184 Acrotelsatinae  159, 182 Acrotelsella  160 Actornithophilus 461 Acutalini  528 Acyrthosiphon A. pisum  604, 605 A. svalbardicum 31, 36, 39, 42 Adamsia 650 Adelges 598 Adelgidae  599, 602, 608, 612 Adelidae  67

Adelphohemerobiinae  640 Adelphohemerobius enigmaramus 649 Adenissini  535, 538 Adexia erminia  532 Adicophasma A. grylloblattoides (extinct)  345, 351 A. spinosum (extinct)  344, 351 Adrothrips intermedius  489 Aecalus 521 Aedes 3 Aemodogryllinae 260 Aeneolamia 519 Aeolothripidae  70, 484, 485, 486, 490 Aeolothrips 486 Aeshnidae  19, 68 Aetalion 527 A. reticulatum  514 Aetalionidae  502, 514, 516, 521, 522, 525, 526 Aetalioninae  502, 514, 526, 527 Aetalionini 527 Aethalochroaini  394 Afrobittacus 709 Afrochilis  157 Afroheriades  857 Afrolepisma  160, 186 A. leleupi 168 A. wygodzinskyi 186

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

944

Index

Afromachilis  157 Afropsocus 439 Agalliinae 523 Agalmatium bilobum 545 Aganacris 253 Agaonidae  66, 820 Agathemera  289, 290, 294 Agathemerinae  285 Agenia  507 Agonoxenidae  67 Agrionidae  68, 73 Agromyzidae  21, 63, 743, 769 Agrotis infusa 811 Agulla 654 Agullini  641 Akicerinae 270 Albardia 652 A. furcata 644 Albardiinae  638, 644, 645, 652 Alenini  641 Aleurodicinae 594 Aleuropteryginae  639, 647 Aleyrodidae  66, 592, 594, 599, 604, 612, 613 Aleyrodinae 594 Alienobostra brocki  284 Allacrotelsa  160, 178, 186 A. burmiticus (extinct)  165 A. dubia (extinct)  165, 186 A. kraepelini 186 A. spinulata 186 Allatelura  161 Allodahlia scabriuscula  320 Allodapini 852, 859 Allograssiella  161 Allomachilis  158, 185 Allomeinertellus  158 Allomorphura  161 Allomorphuroides  161, 176 Allonicoletia  163, 176 Allonychella  161 Allopsontus  157 Allostethus indicum 325 Allotrichotriura  164 Allotrinemurodes  164 Alloxystidae  22, 66 Alucitidae  67

Aluma 546 Alydidae  65 Amara A. alpina 36 A. quenseli 28 Amastrini  528 Amberana 520 Ambermantis 406 Ambonga 521 Ameles 395 Amelinae  392, 393, 398, 401 Amelini  392, 393 Amiseginae 281 Amitermes laurensis 373 Ammobatini  858 Ammobatoidini  858 Ammophila 690 Amorphoscelidae  68, 391, 392, 393, 395, 397, 398, 401, 406 Amorphoscelinae 391, 393, 395, 397, 398 Amphecostephanus 399 Amphientomidae  69, 420, 423, 426, 428, 431, 436, 436 Amphientomoidea  428, 431, 436 Amphigerontia 425, 446 A. bifasciata  424, 425 Amphigerontiinae 446 Amphinemura palmeni 31 Amphipsocidae  420, 426, 427, 429, 432, 439 Amphipsocus 439 Amphipsyllinae  465 Amphipterygidae  68 Amphiteridae  67 Ampulicidae  66 Amyclinae  505 Anabittacus 709 Anallacrotelsa  160 Anallacta 360 Anaphothrips 488 Anaplecta brachyptera 371 Anaplectidae 362, 363, 364, 371 Anarithmeus  161

Anaxarchini  393 Anchiale austrotessulata 303 Anchistrotus 527, 528 Ancistropsyllidae  69, 465 Ancylini  859 Ancyloscelis  859 Ancyra 531, 542 Andes 538 Andesembia 231 Andesembiidae  220, 231, 233, 236 Andrena 40, 677, 688, 689, 694, 852 A. complexa 688 A. minutula 688 A. stromella 688 A. suavis 688 A. vaga 688 Andrenidae  683, 854, 855, 856, 859, 861 Andricus 609 Anechura harmandi 323 Anelpistina  163, 178 A. quinterensis 184 A. specusprofundi 180 Aneuretopsychidae (extinct)  756 Angela guianensis 398 Angelidae  393, 395, 398 Angelinae  392, 398 Anisacanthidae  285, 295 Anisacanthini 292 Anisembia texana  222, 229, 233 Anisembiidae  220, 222, 232, 233, 236, 238 Anisogamia tamerlana 376 Anisolabidae  320 Anisolabididae  63, 316, 317, 325, 326 Anisolabis A. hawaiiensis  141 A. howarthi  141 A. maritima  141, 317, 323, 326, 327 Anisolepisma  160, 182 Anisomorpha paromalus  284

Index

Anisopodidae  63 Anomalobittacus 709 Anomalochrysa 646 Anomiopsyllinae  465 Anomiopsyllus 464, 467 Anopheles  3, 775 A. gambiae 3 Anophthalmus schmidti 128 Anoplolepis gracilipes 93 Anoplophilinae 260 Anostostomatidae  69, 251, 252, 253, 256, 258, 260, 261 Antarctophthirus microchir 463 Antemna 399 Antemninae  392, 393 Anterhynchium 690 Anthidiini  858, 861, 862 Anthocoridae  19, 65 Anthomyiidae  21, 25, 27, 28, 31, 41, 63 Anthonomus grandis 890 Anthophorini  858 Antianthe 526, 528 Antipaluria 235 A. aequicercata  222 A. urichi  221, 221, 224, 228, 228, 229, 230, 235 Antongiliinae  285, 295 Apachyidae  63, 316, 325, 326 Apamea zeta 28 Apatania zonella 37 Apataniidae  70 Apateson 537 Apatesoninae  504, 537 Aphaeninae  505 Aphalaridae  66, 595 Aphelinidae  66, 686 Aphelocheiridae  65 Aphididae  19, 66, 108, 108, 110, 592, 594, 598, 599, 602, 604, 608, 612, 769, 805 Aphidinae  594, 608, 609 Aphidius A. ervi 604

A. leclanti 31 Aphidoidea  23, 25, 31, 592, 594, 599, 602, 608 Aphis A. glycines 593 A. gossypii 111 A. nerii  71 A. sambuci  592 Aphodius A. pseudolividus 108 A. rufipes 111 Aphrodinae  503 Aphrophora A. cribrata 520 A. maculata 519 Aphrophoridae  64, 502, 510, 512, 518, 519, 520, 521, 675, 684, 692 Aphrosiphon 521 Apia 546 Apidae  22, 23, 28, 66, 72, 109, 636, 684, 764, 814, 815, 851, 855, 858, 861, 862, 870, 873, 874, 884, 884 Apinae  858, 862 Apini  858 Apioceridae 754 Apiomorpha 610 A. macqueeni 610 Apis 40 A. cerana  108, 876 A. dorsata  71 A. mellifera  107, 804, 851, 852, 871, 876, 888 Apochrysinae  639, 646 Apoidea 804, 856 Aposthonia  236, 237, 238 A. ceylonica  228, 237 A. gurneyi 237 A. japonica 219 Aptera fusca  360 Apterobittacus 709 Apterograeffea 299 Apteromantis 395 Apteropanorpa 712 Apteropanorpidae 705, 706, 710, 712

Apteryskenoma  160 Arabiatelura  161 Arachisothrips 491, 492 Arachnocampa luminosa 124 Aradidae  65 Araneae (= Araneida)  651, 733, 878, 885 Archeatelura (extinct)  165, 186 Archeatelura sturmi 165, 166 Archembia  233, 234 Archembiidae  220, 231, 233, 234, 235, 236, 238 Archembiinae 233 Archeognatha. See Microcoryphia Archiblattinae  364, 369, 370 Archimantinae  393 Archipseudophasmatidae 302 Archipsocidae  420, 422, 427, 428, 432, 438 Archipsocopsis 438 Archipsocus 438 A. indentatus  427 Archotermopsidae  67, 363, 372 Arctiinae  22, 35, 67, 72 Arenivaga 365 A. bolliana 376 A. investigata 365 Arexeniidae 457 Argidae  67 Argosarchus 303 Arixenia  323, 325 A. esau  317 Arixeniidae 315, 316, 316, 317, 318, 325 Arytaininae 595 Asantorga 546 Asarcopus palmarum 538 Ascalaphidae  68, 628, 629, 631, 633, 634, 635, 638, 642, 644, 652 Ascalaphinae  638, 644 Aschiphasmatidae 291 Aschiphasmatinae  285, 290, 291, 294, 300 Aschiphasmatini 290

945

946

Index

Asilidae  63 Asiolepisma  160 Asiopsocidae  421, 423, 427, 429, 432, 439 Asiopsocus 439 A. sonorensis  423 Asiracinae  504, 539 Aspidosmiini  857, 861 Assmuthia  161 Asteiidae  63 Asterolecaniidae  612 Atacamacridinae 271 Atelura  161 A. formicaria  179, 180 Ateluridae 181 Atelurina  162 Atelurinae  159, 165, 175, 176, 177, 179, 180, 182, 183, 185, 186 Atelurodes  162 Ateluropsis  162 Athericidae  63 Atheta graminicola  29, 39 Atopatelura  162 Atractodes pusillus 41 Attaphila  364, 366 Attaphilinae. See Blattellinae Attatelura  162 Attevidae  67 Attinae 372 Aucaridinae 270 Augilini 538 Augochlorini  857 Aulacidae  66 Aulacigastridae  63 Aulacothrips 487 Auroropsocus 444 Ausallatelura  162 Australembia  222, 234 Australembiidae  220, 224, 230, 231, 232, 234, 235 Australiatelura  162 Australoxenos  682, 685 Austroagalloidinae  503 Austrobittacus 709 Austrogymnocnemia edwardsi 649

Austromerope 711 Austroneurorthus 653 Austropanorpidae 712 Austrophasma  344, 350 A. caledonense  344, 350 A. gansbaaiense  344, 350 A. rawsonvillense  344, 350 Austrophasmatidae  340, 344, 347, 350, 351 Austrosaginae 262 Aylaella  507

b

Bachilis  157, 169, 173 Bacillinae  284, 285, 295 Bacillus  295, 303 B. rossius  284 Bacteria  293, 294 Bactericera cockerelli 604 Bactocera 693 Bactrododema  294, 297 B. hippotaurum 294 Bactrophorinae  251, 271 Baculofractum 299 B. insigne  284 Badonnelia titei 435 Baetidae  19, 64 Baetis bundyae 37 Bahiaxenidae  674, 675, 678, 681, 682 Bahiaxenos 678, 682 B. relictus 678 Balmes 656 Balta 367 Baltipanorpa 713 Bantiinae  394 Basiceros manni 206 Bathynella 129 Bathysciola 146 Bathysciotes khevenhuelleri 128 Bathysmatophorinae  503 Batrachideinae  263, 265 Beaufortianini 530 Bebaiotes 531, 537 Bebaiotinae  504 Bedelliidae  67

Beesoniella 521 Beesoniidae 595, 612, 613 Behningiidae  64 Belapha 435 Belaphopsocus 435 Belaphotroctes 435 Bellinympha filicivora (extinct)  763 Belostomatidae  65 Bemisia tabaci 591, 592, 604 Bennini 537 Berothella 647 Berothidae  68, 628, 629, 630, 631, 631, 635, 636, 637, 638, 639, 642, 645, 651 Berothimerobiinae  639, 645 Berothinae  639, 645 Bertkauia 445 B. crosbyana  419, 421 Berytidae  65 Bessoniella 146 Bethylidae  66 Bharatatelura  162 Biastini  858 Bibionidae  20, 63 Bicaubittacus 709 Bidessini  142 Binaluana 546 Biolleyana 533, 547 Biroellinae 266 Bisma 546 Biston betularia 810 Bittacidae  68, 706, 706, 707, 708, 712 Bittacus  708, 709 Biturritiinae  502, 514, 527 Blaberidae  63, 360, 361, 362, 363, 364, 366, 368, 373, 374, 376 Blaberinae  364 Blaberoidea 366–369 Blaberus 374 B. craniifer 376 B. discoidalis 374 Bladina molorchus 547

Index

Blaste 446 Blastobasidae  67 Blatta orientalis  362, 369, 370, 374, 375, 376 Blattella  361, 362 B. germanica  362, 366, 373, 374 Blattellidae. See Ectobiidae Blattellinae  363, 364, 366 Blattidae  63, 360, 362, 363, 364, 369, 374, 675, 684, 692 Blattinae  364, 370 Blattodea (= Blattaria, Blattoptera)  61, 63, 70, 130, 144, 199, 209, 219, 359–377 (chapter 14), 389, 602, 673, 684, 705, 805, 886 Blattodeaphagus  684, 692 Blattogryllidae (extinct)  352 Blattoidea 369–373 Blephariceridae  63 Blepharodinae  393 Blissoxenos  682, 685 Boccharini 530 Bohartilla  683, 691 B. megalognatha 690 Bohartillidae  674, 681, 683, 690 Bojophlebia (extinct)  159 Bolitophilidae  24 Boloria  27, 30, 36, 37 B. chariclea  26, 42 Bombini  858, 888, 889 Bombus  40, 107 B. bellicosus 811 Bombycidae  67 Bombyliidae  63 Bombyx mori 877 Boocerini 529 Boopiidae  458, 459 Bootetix argentatus 251 Boreellus atriceps  33 Boreidae  706, 706, 710 Boreinae 710 Boreophila subplana 29

Boreus 710 B. californicus  710 Borneacridinae 267 Borystheninae  504, 538 Bothriocerinae  504, 538 Boundarus 513 Bourgoinrana 520 Brachodidae  67 Brachycentridae  22, 70 Brachyctenonotini  465 Brachynomadini  858 Brachypanorpa  712, 713 Brachypsocus badonneli 439 Braconidae  22, 23, 31, 33, 38, 39, 41, 42, 66, 72, 817 Bradiopsyllinae  465 Bradyporinae 262 Brahmaeidae  67 Brancsikia  399, 400 Braulidae  63 Brixia 538 Brucheiserinae  640, 647 Bruchomorpha jocosa  531 Brueelia  458 Brunneria 399 B. borealis 399 Bryoembia 231 Bryopsocidae. See Philotarsidae Buca  507 Bulbosembia 237 Bullacris membracioides 252 Bumerangum deckerti 547 Bupestridae  20, 871 Burmitembia (extinct)  236 Burmitembiinae (extinct)  239 Buxtoniella 546 Byrrhidae  20, 31

c Caconemobius C. fori  141 C. sandwichensis  141 C. varius  141 Cacopsylla C. groenlandica 26 C. myrtilli 37 Caecilius  440, 441

C. fuscopterus 440 Caeciliusidae  69, 421, 424, 426, 429, 432, 439, 440 Caeciliusini 441 Caelifera 262–271 Caenidae  64 Caenocholax  683, 685, 686, 687, 688, 694 C. fenyesi  680, 688 Caenohalictini  856, 857 Caenoprosopidini  858 Calacanthia trybomi  33 Calamoceratidae  70 Calamoclostes  233, 234 Calamothespis 405 Caliothrips fasciatus 493 Caliridinae  392, 394, 404 Caliscelidae  65, 504, 531, 534, 535, 537, 538, 545 Caliscelinae  504, 538 Caliscelini 538 Callibia diana 396 Callidulidae  67 Callimantis 400 Calliopsini  856 Callipappidae  612 Callipharixenidae  69, 684, 692 Callipharixenos  684, 692 C. philippines 693 Calliphoridae  21, 27, 33, 41, 63, 872 Calliptaminae  253, 268, 269 Callirhytis 609 Callitettix versicolor 519 Callomelittinae  857 Calolampra C. elegans 375 C. solida 375 Calomantispinae  640, 650, 651 Calophyidae  66, 595 Calopsocidae. See Philotarsidae Calopterygidae  68 Camelopsocus 447 C. bactrianus  419, 425 Campanulotes compar 464 Campodeidae 146 Camponotus leonardi  732

947

948

Index

Campylenchia  528 Canaceidae  63 Canachus 298 Canariletia  163 Cantharidae  20 Capillopsocus 439 Capniidae  19 Carabidae  20, 23, 25, 27, 29, 31, 39, 134, 146, 840 Carausius morosus  282, 299 Carbotriplura (extinct)  159 Carcinophoridae  63 Cardiopterinae  394, 403 Cariblatta imitans 367 Carlius 297 Carnidae 457 Carobiinae  640 Carposinidae  67 Carrionia  545, 546 Carrioniini 546 Carsidaridae  66, 595 Cartoblatta pulchra 370 Catamachilis  157 Catantopinae  268, 269 Caupolicanini  857 Caurininae 710 Caurinus 710 Cecidomyiidae 17, 20, 63, 808 Cedarbergeniana imperfecta 251 Cediopsylla simplex 473 Cedusa 541 Cedusinae  505, 540 Celyphidae  63 Centridini  858 Centrocharesini  528, 530 Centronodinae  502, 515, 527, 528, 529 Centrotinae  502, 515, 527, 528, 529, 530 Centrotini 530 Centrotus cornutus 509 Centrotypini 530 Cephalocryptina  162 Cephaloleia 820 Cephisus siccifolia 520 Cerambycidae  20, 70, 883, 886

Ceraphronidae  22, 41, 66 Cerastipsocini 446 Cerastipsocus 446 Cerataphidini  599, 615 Ceratina 852 C. cognata 108 Ceratinini  859 Ceratomantis 401 Ceratophyllidae  21, 69, 465 Ceratophyllus  464, 469, 472 C. arcuegens 467 C. celsus 472 C. gallinae 475 C. niger 475 C. sciurorum 469 C. scopulorum 472 C. vagabundus 41 Ceratopogonidae  17, 18, 20, 24, 42, 63 Ceratopsyllidae  21 Cercopidae  65, 502, 510, 512, 518, 519, 520, 521, 675, 684, 692 Cercopinae  518, 520 Cercopis vulnerata  512 Cercopoidea 518–521 Ceresa 526 Ceresini  528 Cerobasis 426 C. annulata 447 C. lineata  419, 424, 427 Cerococcidae  612 Ceuthophilinae 260 Ceuthophilus 260 Chaeteessa  395, 396, 398, 399, 405, 406 Chaeteessidae 391, 393, 395, 395, 398, 402, 406 Chaetocladius perennis  32 Chaetopsylla tuberculaticeps 473 Chaetospania borneensis 323 Chaetotroctes 435 Chaitophorinae 594 Chalcididae  22, 66 Chalcidoidea  4, 680 Challia 325

C. fletcheri 327 Chamaemyiidae  63 Chamaepsylla hartigii  592 Chaoboridae  20, 24, 35, 63 Charilaidae. See Pamphagodidae Charimachilis  156, 157, 173, 174 Chauliodinae  638, 642 Chelicerca 232 C. galapagensis 233 Chelidurella acanthopygia 327 Chelisoches morio  317, 327 Chelisochidae  63, 316, 317, 318, 325, 326 Chelobasis 820 Chelyoidea  528 Chelyopsocus 437 Chiastopsyllinae  465 Chilopoda 168 Chiloxanthus C. arcticus  33 C. stellatus  33 Chimaeropsyllidae  465 Chimaeropsyllinae  465 Chininae 265 Chironomidae  17, 18, 20, 23, 24, 25, 27, 29, 31, 32, 33, 37, 38, 41, 42, 63 Chirothrips  487, 495 Chitoniscus 292 Chlamydatus Chlamydatus acanthioides  33 C. pullus  35, 36, 41 Chlidonoptera  397, 401 Chlorion cyaneum 376 Chlorocyphidae  68 Chlorogomphidae  68 Chloroharpax 397 Chlorolestidae  68 Chloronia 642 Chloroperlidae  19 Chloropidae  21, 63 Choeradodinae  393, 402 Choeradodis  395, 402 Cholevinae 146 Chondroptera  507

Index

Choreutidae  67 Chorista 712 Choristidae  706, 712 Choristopsyllinae  465 Chorotypidae  69, 256, 265 Chorotypinae 265 Choucentrini 530 Chroicoptera 402 Chroicopterinae  392, 393 Chromatoclothoda 235 Chrysididae  66, 281 Chrysolina 36 C. arctica 31 C. brunnicornis bermani 31 C. subsulcata 39 Chrysomantis 401 Chrysomela septentrionalis  33, 39 Chrysomelidae  20, 25, 26, 27, 32, 33, 808, 820 Chrysoperla  111, 646, 658, 659 C. carnea 111 Chrysopidae  20, 68, 628, 630, 631, 631, 633, 635, 636, 638, 639, 642, 645, 646, 649, 658 Chrysopinae  639, 646 Cicadellidae  2, 4, 19, 65, 503, 508, 509, 513, 514, 516, 521, 522, 523, 524, 525, 675, 684, 692 Cicadellinae  503, 513, 516, 522, 523, 524 Cicadettinae  502 Cicadidae  65, 502, 508, 509, 511, 516, 517, 874, 876, 878, 882, 883, 885, 886 Cicadinae  502 Cicadoidea 516–518 Cicadomorpha 511–530 Cicindela albissima 895 Cimbrophlebiidae (extinct) 707 Cimicidae  65, 457, 873 Cinara 592 C. confinis (= Cinara abieticola) 40

Circulifer tenellus 509 Ciulfina 401 Cixidia 537 C. colorata  531 Cixiidae  65, 504, 508, 516, 530, 531, 534, 535, 536, 537, 538, 539, 545, 546 Cixiinae  504, 538 Cixius 538 Cladiopsocidae  421, 429, 432, 444 Cladiopsocus 421, 444 Cladodipterinae 543 Cladodipterinae. See Cladyphinae Cladomorphinae  285, 294, 296, 297 Cladonota  528 Cladonotinae 265 Cladyphinae  505, 543 Clarazella 270 Clastoptera 512, 520, 521 C. theobromae 521 C. undulata 521 Clastopteridae  502, 510, 512, 516, 518, 519, 520, 521 Clastopterini 521 Cleptidae  66 Climacia 656 Climaciella brunnea 650 Clitarchus hookeri  302, 303 Clitumninae  284, 296 Clitumnini  285, 295, 296 Clonaria  295, 297 Clonaspes 546 Clonopsis  295, 303 Clothoda  229, 235 C. longicauda 235 Clothodidae  220, 221, 222, 224, 228, 230, 234, 235, 239 Clusiidae  63 Cnaphalocrocis patnalis 109 Cnipsus 297 Coccidae  19, 66, 598, 601, 602, 612, 615 Coccinella trasversoguttata 42

Coccinellidae  20 Coccoidea 31, 592, 594, 595, 600 Coccus hesperidum 596 Coelidiidae  65 Coelidiinae  503, 522, 523, 524, 530 Coelioxoides  858 Coelopidae  33, 41 Coelostomidiidae  600, 612 Coenagriidae. See Coenagrionidae Coenagrionidae  19, 68, 818 Cofana spectra 693 Colacina  684, 691 Coleomegilla maculata 110 Coleophoridae  67 Coleoptera  5, 18, 20, 23, 23, 27, 28, 30, 31, 35, 36, 37, 59, 61, 63, 70, 71, 75, 88, 130, 144, 146, 168, 318, 457, 628, 651, 659, 674, 726, 752, 769, 796, 804, 805, 813, 883, 891 Coleoscytidae 530 Coleoscytoidea 530 Coleotroctellus 438 Coletinia  164, 183 Coletiniinae  159, 182 Colias  27, 36, 37 C. hecla  26, 42 Collembola 38, 130, 143, 144, 728, 733 Colletidae  66, 683, 688, 855, 857 Colletinae  857 Colobathristidae  65 Coloceras tovornikae 464 Colophina arma 615 Colpocephalum 461 Colpopterinae  506, 547 Colpopterini 536 Columbicola 461 C. columbae 464 Comicinae 258 Comininae  258 Comphotriura  162

949

950

Index

Complaniamphus 439 Compsocidae  420, 424, 427, 431, 437 Compsocus 437 Compsothespinae  392, 393, 402 Compsothespis 390, 395, 398, 402 Comstockiella  611, 615 Conanthalictini  856 Conchaspidae  612, 613 Conditor 521 Congoatelura  162 Congoharpax 400 Congoxeninae 678 Congoxenos  678, 679, 682 Coniocompsini  639 Coniopterygidae  68, 628, 631, 633, 634, 638, 639, 647, 649, 658 Coniopteryginae  640, 647 Conocephalinae  258, 262 Conophyminae 269 Conopidae  63 Coolooidae. See Anostostomatidae Copicerus irroratus  531 Copiocerinae 269 Coprini 110 Coptacridinae 268 Coptopsyllidae  465 Coptopterygidae  393, 395, 399 Coptopterygini 399 Coptopteryx 399 Coptotermes 373 Cordulegastridae  68 Corduliidae  19, 68 Cordylochernes scorpioides 820 Coreamachilis  157, 170 Coreidae  65, 675, 684, 692 Corethrellidae  63 Corethromachilis (extinct)  157 Corethrura 546 Coriophaginae 692 Coriophagus  684, 692 Corioxenidae  69, 674, 675, 676, 681, 682, 685

Corioxenos  682, 685 Corixidae  19, 35, 65 Cornucollis 401 Corrodopsylla curvata 472 Corticomantis 402 Corydalidae  68, 73, 629, 631, 632, 634, 635, 638, 642, 642, 644 Corydalinae  638, 642 Corydalus 642, 642 Corydiidae  63, 360, 362, 363, 363, 365, 370, 374, 376 Corydiinae 363, 365, 365 Corydioidea 363–366 Cosmopterigidae  67 Cosmoscartinae 520 Cosmozosteria 370 Cossidae  67 Cotesia kazak 817 Crabronidae  66, 684, 854 Crambidae  67 Craneopsyllinae  466 Cranopygia marmoricrura  320, 323 Craspedorrhynchus 461 Craspedothrips 486 Crawfordia  683, 688 Crematogaster laeviuscula 688 Creobroter 401 Creoxylus spinosus  284 Cretaceomachilis (extinct)  165, 185 C. libanensis 164 Cretostylopidae (extinct)  674, 678, 682 Cretostylops (extinct)  682 Crocinae  641, 652, 653 Crotonothrips polyalthiae 494 Crustacea  127, 733 Cryptocephalina  162 Cryptocercidae  63, 359, 360, 362, 363, 364, 371, 372 Cryptocercinae  364 Cryptocercus 360, 361, 362, 371, 373 C. clevelandi 362

Cryptoclothoda 235 C. spinula 235 Cryptostylea  162 Crypturella  162 Crypturelloides  162 Ctenarytaina eucalypti 603 Ctenisolabis 320 Ctenocephalides felis  468, 472 Ctenolepisma  160, 167, 176, 178, 179, 183, 186, 678 C. ciliata 167 C. diversisquamis 186 C. electrans (extinct)  165, 166 C. kashinicum (extinct)  165 C. lineata  167, 184 C. longicaudata  177, 180, 184, 186 C. nicoleti 167 C. rothschildi  184, 186 C. targioniana 184 C. targionii 167 Ctenolepismatinae  159, 182 Ctenophthalmidae  465 Ctenophthalminae  465 Ctenoplectrini  858 Ctenopsocus 439 Ctenostylidae  63 Ctenuchidae  67 Cubacubaninae  159, 176, 179, 182, 183, 184 Cubipilis 440 Cucujidae  20, 38 Cucujus 71 Culex quinquefasciatus 818 Culicidae  20, 27, 41, 42, 63, 775, 873 Cuneopalpus 442 Curculionidae 4, 20, 25, 31, 37, 88, 821 Curtonotidae  63 Cycadothrips albrechti 489 Cyclohemipsocus 446 Cyclotornidae  67 Cydnidae  65, 682, 685 Cylindrachetidae 250, 256, 263 Cymbomorphini 529

Index

Cynipidae  22, 66 Cynomya mortuorum  33 Cyphoderrinae  258, 260 Cyphoderris 260 Cypselosomatidae  63 Cyrenoberothinae  639, 645 Cyrtacanthacridinae 253, 268, 269 Cystococcus 608

d Dachtylembia siamensis  239 Dactylocerca rubra 233 Dactylopiidae 595, 612, 658 Dactylopius  601, 602, 658 Dactylopsyllinae  465 Dajaca  290, 291, 294 Dajacini 290 Dalbulus D. elimatus 693 D. gelbus 693 D. maidis  524, 693 Dallaiphasma 290 Damasippoididae  294, 295 Damasippoidinae  285 Danaidae. See Nymphalidae Danaus plexippus  811, 895 Danuriini  392 Darninae  502, 515, 527, 528, 529 Darthula 526 Darthulini 526 Dasydemella 439 Dasydemellidae  425, 427, 429, 432, 436, 439 Dasyleptidae 159 Dasyleptus (extinct)  159 D. brongniarti 159 D. lucasi 159 Dasypodainae  856, 860, 861 Dasypodaini  856 Dataminae  293 Datamini 292 Deinelenchus  684, 691, 693 D. australensis 692

Delphacidae  19, 65, 504, 509, 531, 534, 535, 538, 539, 540, 674, 675, 684, 691, 692 Delphacinae  504, 539 Delphacini  539, 540 Deltocephalinae  503, 513, 523, 524 Dendrothripinae  485 Deois 519 Derbidae  65, 505, 531, 534, 535, 540, 541, 675, 684, 692, 693 Derbinae  505, 541 Dericorys albidula 269 Dericorythidae  69, 257, 268, 269 Dericorythinae 269 Dermaptera  61, 63, 71, 144, 210, 315–327 (chapter 12), 457, 886, 891 Dermestidae  20 Deroplatyinae  394, 398 Deroplatys 402 Desertinoma  160 Diadocidiidae  24, 63 Diaeretellus svalbardicum  31, 39 Dialectus 40 Diamesia D. aberrata 34 D. arctica 34 D. bertrami 34 D. bohemani 34 Diapherodes gigantea 296 Diapheromera  290, 294 D. femorata 303 Diapheromeridae  69 Diapheromerinae  284, 285, 290, 293, 294, 296 Diapheromerini 294 Diaphorina citri  591, 603, 604, 607 Diapriidae  22, 66 Diaspididae  66, 592, 595, 596, 600, 612, 613

Diastatidae  63 Dibrachys microgasteri 681 Dicerapanorpa 715 Dichopterinae  505, 543 Dicondylus lindbergi 691 Dictyopharidae  65, 505, 532, 534, 535, 541, 543, 675, 684, 691, 692, 693 Dictyopharinae  505, 541, 542 Dictyoptera  210, 246, 389, 398 See also Blattodea and Mantodea Didymuria violescens  284, 303 Dienerella D. elegans 33 D. filum 39 Diesbachia tamyris  284 Dilar 647 Dilaridae  68, 629, 630, 631, 635, 638, 640, 647, 649 Dilta  157, 167, 169, 170, 171, 172, 186 D. bitschi 171 D. hybernica 171 D. littoralis  171, 172 D. machadoi 172 Dimidistriata 444 Dimorphodes 299 Dinatelura  162 Dinatelurini 180 Dinembia 236 Dinidoridae  65 Dinopsyllinae  465 Dionychella  162 Diopsidae  63 Diostrombus mkurangai 541 Dioxyini  857 Diphaglossinae  857 Diphaglossini  857 Diplatyidae  63, 316, 317, 318, 323, 325, 326 Diplatys 323 D. flavicolla 325 Diplopoda 821 Diploptera punctata  361, 362 Diplopterinae  364

951

952

Index

Diplura  130, 144 Diprionidae  67 Dipseudopsidae  70 Diptera 18, 20, 23, 23, 24, 25, 25, 27, 31, 32, 33, 33, 35, 36, 37, 39, 40, 41, 42, 61, 63, 72, 130, 144, 146, 457, 651, 673, 684, 692, 706, 707, 708, 715, 726, 743, 756, 756, 758, 759, 766, 767, 804, 805, 813, 843, 876, 878, 880, 882, 883, 884, 890, 892 Dipterophaginae 692 Dipterophagus  684, 692 D. daci  692, 693 Diradius 239 Disparoneuridae  68 Dissoglottini  857 Ditrigoniophthalminae  156, 174, 175 Ditrigoniophthalmus  157, 173, 174 Diuraphis noxia 593 Dixidae  20, 63 Dodecastyla  162, 186 Dolabellopsocidae  425, 429, 432, 444 Dolabellopsocus 444 D. similis  425 Dolichoderinae  683, 686, 813 Dolichopodainae 260 Dolichopodidae  20, 25, 63 Dolichovespula 40 Doratopsyllinae  465 Doru lineare 324 Dorylinae 179 Dorypteryx 434 Drepanacrinae  640 Drepanepteryginae  640 Drepanepteryx phalaenoides 649 Drepanicinae  640, 650, 651 Drepanidae  67 Drepanosiphinae 594 Drosophila  4, 88, 327, 605, 753, 795, 820, 842

D. melanogaster 5 Drosophilidae  21, 63 Dryinidae  66, 72 Drymopsalta daemeli  511 Dryococelus australis  298, 298 Duchailluia  364 Duchailluiinae  364, 370 Dunatothrips 490 D. aneurae 490 Dundoxenos  682, 685 Duvalius 146 Dynastinae  878, 893 Dysmorphoptilidae (extinct) 516 Dystactinae  392, 394 Dytiscidae  20, 25, 29, 35, 36, 129, 134

e

Ebhuloidesini  528, 530 Echidnophaga gallinacea  468, 468, 475 Echinophthiriidae  19, 458, 463 Echinosoma 317 Echinothrips americanus 495 Echinotropinae 270 Eciton 814 Ecitoninae  683, 686, 687 Ecnomatelura  162 Ecnomidae  70 Ectobiidae (= Blattellidae)  63, 360, 362, 363, 364, 366, 374, 675, 684, 692 Ectobiinae  363, 363, 364, 366, 367 Ectobius  362, 363, 367 E. lapponicus 375 E. pallidus 367 E. panzeri 362 Ectopsocidae  421, 422, 425, 429, 432, 436, 441 Ectopsocopsis cryptomeriae 441 Ectopsocus 447 E. briggsi  421, 425 E. meridionalis  436

E. pumilis 441 E. richardsi 441 Edriobittacus 709 Egnatiinae 268 Egropinae  507, 548 Eicissus 512 Eidoporisminae  641, 655 Elachistidae  67 Elaphrothrips 488 Elasmoscelini 546 Elasmoscelis 546 Elateridae  20, 871 Electrentomidae  419, 420, 423, 431, 437 Electrentomoidea  428, 431, 436, 437 Electrentomopsis 437 E. variegata  420, 424, 427 Electrentomum 437 Elenchidae  69, 674, 675, 684, 684, 688, 691, 694 Elencholax  684, 691 Elenchus 680, 684, 691 E. japonicus 691 E. tenuicornis 691 E. varleyi  681 E. yasumatsui 691 Elicinae  507, 537, 545, 548 Elipsocidae  419, 421, 422, 424, 429, 432, 441, 442 Elipsocus 442 E. guentheri  424 E. obscurus  421 Ellipsidion 367 Eluratinda  162 Embia E. major 236 E. nuragica  224 E. ramburi  229, 236, 238 Embidopsocinae 435 Embidopsocopsis 435 Embidopsocus 423, 435 E. bousemani  419 E. needhami  419 Embiidae  64, 219, 220, 224, 230, 231, 233, 235, 238 Embiidina. See Embiodea

Index

Embiodea  61, 64, 72, 210, 219–240 (chapter 9), 246, 286, 288 Embioptera. See Embiodea Embolemidae  66 Embolyntha E. batesi 219 E. interrupta  222, 236 Embonychidae  64, 220, 220, 222, 231, 236 Emphorini  859 Empididae  20, 25, 38, 41, 63 Empidoidea  24 Empoasca fabae 509 Empusidae  68, 391, 393, 395, 399, 400, 402 Empusinae  393 Empusini  393 Encyrtidae  22, 66 Enderleinella 440 Enderleinellidae  69, 458 Enderleinia 521 Enderleiniinae 521 Endoiastinae  502, 514, 526, 527, 528, 529 Endoiastus 514 Endromidae  67 Eneopterinae 255 Enicophlebia hilara  390, 401 Ensifera  254–262, 884, 885, 886, 888 Ensiferophasma velociraptor (extinct)  345, 351 Eoarthropleura (extinct)  765 Eocenoxenos (extinct)  682, 685 Eocercopidium maculata 518 Eomerope (extinct)  711 Eomeropidae  706, 711, 712 Eophyllium messelense (extinct)  291, 300 Eosembia 237 E. auripecta  223, 226 Eoxenos  678, 679, 680, 681, 682 E. laboulbenei 167, 679, 680, 681, 681 Epaphrodita  399, 400, 401

Epaphroditidae  393, 395, 399, 400 Epeolini  858 Epeoloides  858 Epermeniidae  67 Ephemerellidae  19, 64 Ephemeridae  64 Ephemeroptera 18, 19, 23, 24, 27, 31, 34, 37, 61, 64, 72, 90, 164, 766, 883, 888, 891 Ephydridae  21, 63 Epicharmus 299 Epicopeiidae  67 Epidares 292 Epilampra E. abdomennigrum 369 E. involucris 369 E. involucris 369 E. irmleri 376 E. maya 369 E. rothi 369 Epilamprinae  364, 369 Epipsocidae  69, 419, 421, 423, 429, 433, 444, 445 Epipsocopsis 445 Epipsocus 423, 445 E. foliatus  423 Epipyga 521 Epipygidae  502, 510, 512, 518, 519, 521 Epipyropidae  67 Epirimiinae  465 Episactidae  256, 264, 265 Episactinae 265 Epitroctes 437 E. tuxtlarum  419, 420, 423, 424 Epora subtilis 691 Erebia 27 Erebidae  67 Erechtia  528 Eremiaphila 395 Eremiaphilidae 391, 392, 393, 395, 400, 406 Eremobittacus 709 Eremogryllinae 268

Ergaula capensis 365 Erianthinae 265 Ericrocidini  858 Eriococcidae (sensu lato)  595, 601, 602, 612, 613 Eriophyidae 486 Errhomeninae  503 Errhomenus brachypterus 509 Eruciinae 265 Erythraeidae  69 Erythroneura palimpsesta  513 Erythroneurini  523, 524 Espagnolinae 265 Ethmiidae  67 Euacanthellinae 525 Eublaberus posticus 369 Euborellia 326 E. annulipes 327 Eucerini  859 Eucharitidae  66 Euchoplopsyllus glacialis 469 Euglossini 851, 858 Euherbstiini  856 Eulophidae  22, 33, 66 Eumastacidae  69, 256, 265, 266 Eumetopina flavipes 540 Eumorsea 266 Eupelicinae 523 Eupelix cuspidata 509 Eupelmidae  66 Euphaeidae  68 Euphyllodromia 367 Eupithecia 91 Eupolyphaga sinensis 375 Eupterotidae  67 Eurybrachidae  505, 531, 534, 542, 544, 545, 675, 684, 691, 692, 693 Eurybrachinae  505 Eurybrachys tomentosa 542 Eurycantha 298, 299 E. calcarata  284 E. horrida  298 Eurycanthablatta 370 Eurycanthinae 297, 298, 298 Eurycnema 284

953

954

Index

Euryglossinae  857 Eurymelinae  503, 522, 523 Eurynothrips magnicollis 491, 492 Euryphyminae 269 Eurystylops  683, 688 Eurytomidae  66 Euschmidtiidae  256, 265, 266 Euschmidtiinae 266 Euscyrtinae 255 Euthyrraphinae  365 Euthyrrhaphinae 363 Evacanthinae  503, 513 Evaniidae  66 Evansiola 525 Exomalopsini  859 Exparoxypilus africanus  397 Extatosoma 303 Eyprepocnemidinae 269

f

Fanniidae  63 Farhangiinae  466 Fauriellidae  485, 486 Fidelia 860 Fideliinae 855, 857, 860, 861 Figitidae  22, 66 Flatidae  65, 505, 532, 534, 537, 542, 543, 675, 684, 691, 692, 693 Flatinae  505, 542 Flatoidinae  505, 542 Flexamia pict  513 Flexocentrus  528 Floridoxenos  682, 685 Florinemestrius pulcherrimus (extinct)  756 Fontecilla 650 Forficula F. auricularia 315, 317, 319, 322, 323, 324, 326, 327 F. senegalensis  317, 324 Forficulidae  63, 71, 316, 318, 320, 321, 325, 326 Forficulina  316, 318

Formicidae  22, 66, 72, 90, 179, 368, 397, 593, 681, 683, 686, 687, 774, 874, 877, 878, 879, 882, 883, 884, 885 Formicinae  683, 686, 688 Formininae  683, 687 Frankliniella  488, 491, 495 F. occidentalis  486, 490, 494 F. schultzei 486 Fuga 402 Fulcinini  393 Fulgoridae  65, 505, 508, 533, 534, 541, 542, 543, 691 Fulgorinae  505 Fulgoroidea  84, 144, 146, 530–548, 693, 730 Furcatopanorpa 715

g Gaetuliina 535 Gaetuliini 548 Galenatelura  162, 175 Galerucella interrupta 31 Galinthiadidae  393, 395, 396, 400, 401 Galinthias 400 Galloisiana 340, 341, 343 G. chujoi  342 G. kiyosawai  342 G. kosuensis  342 G. nipponensis  342, 343 G. notabilis  342 G. odaesanensis  342 G. olgae  342 G. sinensis  342 G. sofiae  342 G. ussuriensis  342 G. yezoensis 341, 342 G. yuasai  340, 342 Gammarotettiginae 260 Garcialdretia 443 Gargara 526 Gargarini  529, 530 Gastererion  507 Gasterophilidae. See Oestridae Gasteruptiidae  66

Gastriniinae  506, 547 Gastrotheellus  162 Gastrotheus  162 Gelastocoridae  65 Gelechiidae  21, 67 Gelis glacialis 41 Gengidae  506, 533, 534, 542, 544 Gengis. See Acrometopum Geometridae  22, 25, 35, 67, 72, 810, 821 Geoscapheinae  364, 368 Geoscapheini  364 Geracinae 318 Gerridae  65, 885 Gibocercus napoe  235, 239 Giustina 524 Glaciopsyllus antarcticus 469 Glaphyropyga dryas 206 Glosselytrodea (extinct)  627 Glossina 774 Glossosomatidae  22, 70 Glyphembia 233 Glyphipterigidae  67 Goeridae  70 Goiasatelura  162 Goja 445 Gomphomastacinae 266 Gomphidae  68, 73 Gomphocerinae  251, 252, 253, 268, 269 Gomphomastacinae 265 Gonatista 400 Gongylus 395 G. gongylodes 399 Gonolabis electa 324 Gopsilepisma  160 Gracillariidae  67, 774 Graeffea  299, 303 G. crouanii 303 Grallipeza G. cliffi 848 G. placidoides 848 G. spinuliger 848 Graphitarsus  157 Graphium antiphates  71 Graphopsocus 440

Index

G. cruciatus  436, 440 Grassiella  162, 180, 185, 186 Grassiellini  165, 186 Gratidia 284, 295 Gratidiini  284, 285, 295, 296, 297 Grellaphia 521 Gromphadorhina portentosa 376 Gryllacrididae  69, 251, 256, 258, 260, 261 Gryllacridinae 261 Gryllidae  69, 73, 255, 256, 258, 258, 683, 686, 687 Gryllinae 255, 258 Grylloblatta  338, 341 G. barberi  342 G. bifratrilecta  342 G. campodeiformis  340, 342 G. chandleri  342 G. chintimini  342 G. chirurgica  342 G. gurneyi  342 G. marmoreus  342 G. newberryensis  342 G. oregonensis  342 G. rothi  342 G. scudderi  342 G. sculleni  342 G. siskiyouensis  343 G. washoa  343 Grylloblattella 340, 343 G. cheni  343 G. pravdini  343 G. sayanensis  343 Grylloblattidae 336, 337, 338, 338, 339, 341, 342, 351 Grylloblattina 343 G. djakonovi  343 Grylloblattodea  73, 199, 210, 246, 336–343 Gryllomantidae  68 Gryllomiminae 255 Gryllomorphinae 255 Grylloptera. See Ensifera Gryllotalpidae  69, 250, 255, 256, 258, 258, 873, 882

Gryllotalpinae 258 Gryllus bimaculatus 254 Gryonoides glabriceps  790 Grypothrips cambagei 492, 493 Guayaquila 526, 528 Gumilla 655 Gumillinae  641, 655 Gyna henrardi 368 Gynaeophora groenlandica  36, 37, 39 Gynatelura  162 Gyninae  364 Gyrinus opacus 35 Gyropidae  458, 459

h Haania lobiceps 406 Haaniella 292 Haaniinae  392, 404, 405 Habrocneminae 269 Habronattus pugillis  82, 89 Haematomyzidae  458 Haematopinidae  458 Haematopinus suis  459, 460 Haemodipsus setoni 463 Haffneria 461 Hagiomantis 402 Halictidae  66, 683, 688, 855, 856 Halictinae  856 Halictini 852, 857, 859 Halictophagidae  69, 674, 675, 684, 684, 688, 691, 692, 693 Halictophaginae 692 Halictophagus  684, 692, 693 H. australiensis 693 H. calcaratus 693 H. fulmeki 693 H. indicus 693 H. naulti 693 H. palmae 693 H. silwoodensis 692 H. tryoni 692 Halictoxenos  683, 688, 689 Halipliidae  20 Hamophthiriidae  458

Hamza ciliaris 517 Hapalomantinae  393, 401 Hapalomantini  393 Hapalopeza 392, 401 Hapithinae 255 Haplaxius crudus  536, 539 Haploembia 237 H. solieri  223, 227, 237 H. tarsalis  223, 224, 227, 228, 237 Haplogleniinae  639, 644, 652 Haplophallus 443 Haplothrips  487, 488 H. leucanthemi  484 Harmonides  528 Harpagomantis  397, 400 Harpezoneura 439 Harpobittacus similis  709 Haslundichilis  157, 169, 170 Haslundiella  157 Hebridae  65 Heleomyza 133 H. borealis  33, 38 Heleomyzidae  21, 33, 38, 63 Helicopsychidae  70 Helicoverpa 811 H. armigera 817 H. zea  108, 108 Heliomantis 401 Helotrephidae  65 Helvia 401 Hematelura  164, 180 H. doriae 180 Hemerobiidae  20, 68, 628, 630, 631, 633, 634, 635, 638, 640, 649, 649, 658 Hemerobiinae  640 Hemiacridinae  253, 269 Hemiberlesia lataniae 596 Hemideina maori 253 Hemikulina  160 Hemikyptha 515 Hemikypthini  515, 527, 528, 529 Hemilepisma  160 Hemilobophasma montaguense  344, 350

955

956

Index

Hemimeridae 315, 316, 316, 318, 325, 457 Hemimerus  315, 318, 323, 325 Hemineura 442 Hemipsocidae  421, 424, 430, 433, 445 Hemipsocus chloroticus  421, 424, 446 Hemiptera  19, 23, 27, 30, 31, 32, 33, 33, 35, 36, 37, 61, 64, 65, 66, 71, 72, 75, 77, 130, 144, 210, 457, 484, 487, 501–551 (chapter 19), 591–616 (chapter 20), 658, 673, 682, 684, 691, 692, 693, 715, 730, 752, 804, 805, 883 Hemisphaeriini  506, 544 Hemithripidae  485 Hemithyrsocera histrio 366 Hemitrinemura  164 H. extincta (extinct)  165 Hemizygon 521 Henicocephalidae  65 Hepialidae  67 Heptageniidae  19, 64 Heranice miltoglypta  514 Hespenedra 513 Hesperapini  856 Hesperapis H. leucura 688 H. rhodocerata 688 Hesperiidae  21, 67, 72 Hesperoboreus 710 Hesperophasmatini 297 Hesticus  507 Heterocaecilius 424 Heterochaetini  392 Heterogamisca H. chorpardi 365 H. marmorata 365 Heterolepidella  162 Heterolepisma  160, 186 H. bisetosa 180 Heterolepismatinae 181 Heteromorphura  162

Heteromyzidae  63 Heteronemiinae  285, 290, 294 Heteronotinae  503, 514, 527, 528, 529 Heteronotus 514 Heteronutarsus  396, 400 Heteronychella  162 Heteropsontus  157 H. americanus 175 Heteroptera  24, 65, 72, 144, 168, 501, 508, 510, 682, 685, 691, 692, 886 Heteropterygidae 292, 293 Heteropteryginae  284, 285, 292, 293, 293 Heteropterygini 292 Heteropteryx  292, 293 Heterothripidae  485 Hexacentrinae 262 Hexepeolini  858 Hierodula 402 Hilda patruelis  533, 548 Hildinae  507, 548 Himalayachilis  157 Himantopteridae  67 Hindoloides 521 Hindoloidini 521 Hippoboscidae  63, 457 Hiracia  507 Hockeria mengenillarum  680, 681 Hodotermitidae  67, 180, 183, 363, 372 Hohorstiella lata 464 Holarthrothrips = Adiheterothrips 486 Holdgateilla chepuensis  515 Holocompsinae 363, 365 Homotomidae  66, 595 Hoplocoryphinae 405 Hoplophorionini  515, 527, 528 Hoplopleuridae  19, 69, 458 Hormaphidinae 608 Hyalesthes obsoletus  536, 539 Hyalomantis 401 Hyalopsocus 436 Hyblaeidae  67

Hybographitarsus 167 Hybophthiridae  458 Hybotidae  63 Hybusinae 266 Hydrobaenus 42 Hydrobiosidae  70 Hydrometridae  65 Hydrophilidae  20 Hydroporini  142 Hydroporus H. morio 28 H. polaris 28 Hydropsychidae  70 Hydroptilidae  22, 70 Hylaeinae  857 Hylaeus 851 Hylecthrus  683, 688 Hylicellidae 516 Hylicelloidea 516 Hylicidae. See Cicadellidae Hylicinae  503, 522 Hylobittacus 709 Hymenopodidae  68, 391, 393, 395, 396, 399, 400, 401, 402 See also Galinthiadidae Hymenopodinae  393, 395, 401 Hymenopodini  393, 400 Hymenoptera 18, 22, 23, 25, 26, 28, 30, 31, 33, 36, 37, 39, 41, 42, 61, 66, 67, 71, 72, 77, 130, 144, 229, 596, 611, 636, 651, 673, 675, 677, 683, 686, 688, 715, 726, 752, 758, 759, 767, 768, 789, 791, 792, 792, 793, 796, 804, 805, 815, 851–864 (chapter 28), 883, 885, 886, 890, 892 Hymenopus  395, 400, 401 H. coronatus  390, 401 Hymetta balteata  513 Hyperlepisma  160 Hypermeinertellus  158 Hyphinoini 529 Hypochrysa elegans  642 Hypochthonella caeca  532, 544

Index

Hypochthonellidae  506, 532, 534, 535, 544 Hypoderma  42, 870 Hypomachiloides  158 Hypothenemus hampei 821 Hypsaucheniini  515, 528, 530 Hypsipterygidae  65 Hypsoprora  528 Hypsoprorini  528 Hystrichopsylla 467, 468, 468 Hystrichopsyllidae  69, 465

i

Iassinae  503, 513, 523, 524 Iba  518, 520, 521 Ibaliidae  66 Iberoxerninae 678 Icaniidae  65 Icerya purchasi  592 Iceryini  610, 611 Ichneumonidae  22, 23, 25, 28, 31, 33, 38, 41, 66, 72 Idiocerinae  503, 513, 523 Idiocerus 513 I. atkinsoni 693 I. clypealis 693 I. niveosparsus 693 Idiomacromerus gregarius 680, 681, 681 Idioscopus clypealis 693 Idolomantis 391 I. diabolica 399 Idolomorphini  393 Idolothripinae  485, 490 Idolothrips spectrum 490 Illapeliinae 270 Ilomantis 401 Ilvia 537 Ilybus angustior 36 Imblattella impar 367 Immidae  67 Incurvariidae  21 Indiopsocus 447 Inocelliidae  69, 74, 631, 638, 641, 654, 657, 658 Inocelliinae  641 Iranellinae 269

Iridomyrmex purpureus 813 Iridopterygidae  68, 392, 393, 395, 401 Iridopteryginae  393, 401 Iris 404 Ischnocentrus  528 Ischnopsyllidae  69, 465 Ischnorhininae 520 Ischnura hastata 92 Isepeolini  858 Isochnus flagellum 39 Isonychiidae  64 Isopoda 821 Isoptera. See Termitoidae Issidae  65, 506, 532, 534, 535, 536, 537, 538, 544, 545, 547, 548 Issikiella 709 Issinae 544 Issini  506, 544 Isthmopsocus 444 Itarinae 255 Ithone 650 I. fulva  649 Ithonidae  68, 630, 631, 635, 637, 638, 640, 649, 649, 650

j

Janetschekilis  157 Jantarimantidae  68 Jantarostylops  683, 688 Javesella dubia 691 Jeholopsyche liaoningensis (extinct)  756 Jingkara hyalipunctata  515 Jugoda 546 Junodia 401 Juracimbrophlebia ginkgofolia (extinct) 707, 763 Juramantophasma sinica (extinct)  345, 351 Jurassipanorpa (extinct)  713

k Kaestneriella 441 Kalligramma aciedentatus (extinct)  756

Kalligrammatidae (extinct)  756, 757, 759, 762, 763, 773 Kallitaxila granulata 548 Kalobittacus 709 Kalocorinnis 300 Kalotermitidae  67, 363, 372 Karajassidae 516 Karataothripidae  485 Karna  507 Karnyothrips K. flavipes 487 K. melaleucus 486 Karoophasma 350 K. biedouwense  340, 344, 346, 350, 351 K. botterkloofense  344, 350 Karschiellidae  316, 318, 323, 324, 325 Kasserota 546 Kelisiinae  504, 539 Kempyninae  641, 655 Kerkiratrobius  158 Kermesidae  612 Kermesiinae  506, 547 Keroplatidae  63, 124 Kerriidae  66, 601, 612 Kinnara 545 Kinnaridae  65, 506, 532, 534, 535, 545, 546 Kinnarinae  506 Kinzelbachilla (extinct)  682 Kinzelbachillidae (extinct)  674, 682 Kinzelbachus  683, 688 Kladothrips  487, 490 K. sterni  491 Kolbia quisquiliarum 439 Koptothrips 487 Korinninae  289, 291, 294, 300, 301 Korinnis 300 Kosciuscola 251 Krombeinictus nordenae 855 Kronomyrmecolax (extinct)  683, 685 K. fushunicus 687

957

958

Index

Kruegeria 547 Kuschelochilis. See Allomachilis Kuwaniidae  612

l Labia minor  319, 323, 327 Labidura herculeana 92, 320, 323 Labidura L. herculeana 327 L. riparia  317, 320, 322, 323, 324, 326, 327 L. truncata 327 Labiduridae  63, 316, 320, 325, 326 Labidus 814 Labiidae  63 Labramachaerota 521 Lachesilla 442 L. contraforcepeta  436 L. riegeli  421 L. tropica  424, 427 Lachesillidae  69, 421, 422, 424, 427, 429, 432, 436, 441, 442 Lachninae 594 Lachnodius  611, 615 Laemobothriidae  458 Laemobothrion vulturis 457 Lamproblatta albipalpus 370 Lamproblattidae 362, 363, 364, 370, 371 Lamproglandifera 370 Lampyridae  882, 885 Landrevinae 255 Languriidae 457 Laodelphax striatella 540 Laodelphax striatellus  536, 691 Lapithasa 546 Largidae  65 Lasiocampidae  67 Lasioglossum 689 Lasiotheus  162 Lasiotheus nanus 186 Latagophthirus rauschi 463 Lathiceridae 250, 257, 268, 269 Lathridiidae. See Latridiidae

Latiblattella lucifrons 367 Latindiinae 363, 365 Latridiidae  20, 25, 32, 33 Laupala 88 Lauraesilpha  364, 371 Lauraesilpha mearetoi 371 Lauxaniidae  63 Lavora 549 Lecanodiaspididae  601, 612 Lecithoceridae  67, 72 Ledra aurita 509 Ledrinae  65, 503, 513, 522, 523 Leiodidae  138, 139, 146, 457 Lentistivalius insolli 74 Lentulidae 253, 257, 268, 269, 270 Lentulinae 269 Leosthenes 299 Lepidina  164 Lepidodasypus sharovi (extinct) 159 Lepidophorus lineaticollis 31 Lepidopsocidae  419, 420, 421, 425, 427, 428, 431 Lepidoptera  5, 17, 18, 21, 23, 25, 27, 28, 31, 33, 36, 41, 61, 67, 71, 72, 73, 228, 367, 405, 457, 596, 597, 651, 715, 726, 752, 756, 758, 759, 762, 769, 792, 804, 805, 810, 811, 817, 883, 890, 892 Lepidospora  164, 179 Lepidostomatidae  70 Lepidotrichidae (extinct)  159, 165, 175, 180, 181, 186 Lepidotriura  162 Lepidotrix pilifera (extinct)  181 Lepinotus 426 L. inquilinus 447 L. patruelis 447 L. reticulatus  419 Lepisma  160, 167, 183, 185, 186 L. aurea 681 L. polypoda 155

L. saccharina  155, 167, 176, 184, 185 L. terrestris 155 Lepismachilis  157, 186 Lepismatidae  68, 159, 165, 166, 167, 173, 175, 176, 177, 177, 178, 179, 180, 181, 182, 183, 184, 185, 186, 678, 682 Lepismatinae  159, 181, 183, 186 Lepismina  160 Lepitrochisma  160 Leprocaulinus  282, 299 Leptinidae 457 Leptinotarsa decemlineata 109 Leptobelini 530 Leptocentrini 530 Leptocentrus 526 Leptoceridae  22, 70 Leptodirus hochenwartii 123, 124, 124, 125, 133, 134, 135 Leptogenys 324 Leptomachilis  158 Leptomantella 392 Leptopanorpa 715 Leptophlebiidae  19, 64 Leptopodidae  65 Leptopsyllidae  21, 69, 465 Leptopsyllinae  465 Leptynia  295, 303 Leptysminae  253, 269 Lesneia 442 Lesneiidae  429, 442 Lestidae  69 Leucolepisma  161 Leucospidae  66 Leuctridae  69 Lezininae 261 Liassothripidae  485 Libellaginidae  69 Libellulidae  69 Lichenodraculus 253 Lichenomima 446 L. lugens  422, 427 Lichnomesopsyche gloriae (extinct)  756

Index

Ligaria 402 Limacodidae  67 Limnephilidae  22, 70 Limnocentropodidae  70 Limnophyes 36 L. brachytomus  32 L. eltoni  32 L. pumilio  32 Limonia lindrothi 39 Limoniidae 37, 63 Limothrips 487 L. cerealium 493 Limulidae 733 Linadureta  162 Linognathidae  19, 458 Liothrips  487, 488 Liphistiidae  63 Liposcelididae 417, 419, 428, 431, 434, 435 Liposcelidinae 435 Liposcelis  435, 447 L. bostrychophila  419 L. decolor 422 L. entomophila  419 Lipsoelis 435 Liriomyza 810 L. trifolii 810 Listropsyllinae  465 Listroscelidinae  251, 262 Lithidiidae  257, 268, 269 Lithoseopsis 437 L. hellmani  436 Lithurginae  857 Littorophiloscia hawaiiensis  141 Liturgusa  402, 406 L. cursor  390 Liturgusidae  68, 391, 392, 393, 395, 401, 402, 406 Liturgusini 391 Liuopsyllinae  465 Liviidae  66 Loania  682, 685 Lobatophasma 350 L. redelinghuysense  344, 350, 351 Lobocentrini 530 Lobofemora 296

Loboscelidiinae 281 Lobosembia 237 Loensia 447 L. conspersa  422 L. fasciata  422 Lonchodinae  282, 284, 285, 297, 298, 298, 299, 300 Lonchopteridae  63 Loneura 445 L. splendida  420, 423 Lophopidae  65, 506, 533, 534, 542, 544, 545, 546, 547 Lophopinae  506, 546 Lophopini 546 Lophops 533, 546 L. saccharicida  533 Lophopterygella 446 Lophotettiginae 265 Lucanidae  318, 884, 886 Luratea  162 Luridiblatta trivittata 367 Lutzomyia 775 Luzarinae 255 Lycaenidae  21, 67, 72 Lychnocolacidae  674, 675, 681, 683, 688 Lychnocolax  683, 684, 688 L. hispanicus 688 Lycopsyllidae  465 Lycopsyllinae  465 Lycorma delicatula 543 Lycosa L. hawaiiensis  141 L. howarthi  141 Lygaeidae  19, 27, 65, 682, 685 Lygus 3 Lymantriinae  22, 35, 67 Lyncidinae  505, 543 Lyoriella 29 Lystrinae  505

m Maana 546 Maarbarini 530 Machadatelura  162 Machaerota 521 M. coomani  512

Machaerotidae  65, 502, 510, 512, 518, 519, 521 Machaerotini 521 Machilanus 170 Machilelloides  158, 172 Machilellus  158 Machilidae  68, 155, 156, 164, 165, 167, 169, 169, 170, 171, 172, 173, 174, 175, 177, 183, 184, 185, 186 Machilinae  156, 174, 177 Machilinus  158, 169, 170, 172, 173 M. rupestris 173 Machilis  157, 165, 167, 170, 171, 172, 173 M. engiadina 167 Machiloides  158, 170, 185 Machilontus 169, 170 M. yoshii 168 Macrembia 224, 230 Macrocercinae  364, 370 Macromantinae  394, 403 Macropanesthia rhinoceros  362, 368 Macropathinae 260 Macrophasma 282 Macropidini  856 Macropis nuda 852 Macropsinae  503, 523 Macropsontus  158, 165 Macropsyllidae 464, 465 Macrosiphum euphorbiae  604 Macrosteles fascifrons 509 Macrotermitinae 372 Macynia 295 Macyniinae  285 Madagaschiloides  158 Magia 546 M. subocellata 545 Magicicada 517 Mahanarva fimbriolata 519 Maindronia  161 M. beieri 186 M. mascatensis 186 M. neotropicalis 186

959

960

Index

Maindroniidae  159, 180, 181, 186 Makota 546 Malacopsyllidae  465 Malagasyxenos  682, 685 Malandania 283, 297, 299 M. pulchra  283 Malayatelura  162 Malayaxenos  682, 685 Malcidae  65 Malgasia 258 Malgasiinae 258 “Mallophaga”  41, 460–463 Malostenopsocus 440 Manduca 753 Manicapsocus 437 Mantidae  68, 393, 394, 395, 398, 399, 401, 402, 403, 683, 687 Mantis 402 Mantispidae  68, 628, 629, 630, 631, 631, 635, 636, 638, 640, 645, 649, 650, 651 Mantispinae 651 Mantodea  61, 68, 73, 219, 359, 389–407 (chapter 15), 650, 673, 683, 686, 687, 871, 878, 885, 886, 891 Mantoida  390, 398, 403, 405, 406 M. maya  402 Mantoididae 391, 394, 395, 395, 398, 402, 403, 406 Mantophasma 349 M. gamsbergense  344 M. kudubergense  344 M. omatakoense  344 M. zephyrum  344, 349 Mantophasmatidae  344, 347, 349 Mantophasmatodea  73, 199, 210, 246, 343–351 Manueliini  859, 861 Maoripsocini 441 Maoripsocus 441 Marava arachidis 323 Marcenendius 437

Marchalinidae  612 Marellia remipes 250 Marelliinae 269 Margarodidae  66, 607, 612 Margattea nimbata 367 Margatteoidea amoena (extinct) 376 Markia 253 Marmessoidea rosea  284 Masarinae 855 Mastacideidae  256, 265, 266 Mastotermes 373 M. darwiniensis 372, 373, 389 Mastotermitidae  363, 372 Matsucoccidae 607, 612 Matsumuraiella 439 Maxudeini 521 Meconematinae 262 Mecopodinae 262 Mecoptera  61, 68, 73, 77, 705–716 (chapter 23), 756, 756, 762, 876 Medaurini  285, 295, 296 Meenoplidae  65, 66, 506, 532, 534, 535, 545, 546 Meenoplinae  506, 547 Megacarna 546 Megachile pluto 851 Megachilidae  22, 66, 851, 855, 857, 860, 861, 862 Megachilinae 851, 856, 857, 862 Megachilini  857 Megacopta M. cribaria 605 M. punctatissima 605 Megacrania 299 Megalagrion calliphya 818 Megalithone 650 Megaloblatta 367 Megalominae  640 Megaloptera  24, 61, 68, 73, 77, 642–644 Megalyridae  66 Megamerinidae  63 Meganeuridae (extinct)  773

Meganomiinae  856, 860 Megaphasma  290, 293 Megapodagrionidae  69 Megapomponia imperatoria 511 Megophthalminae  503, 525 Meinertellidae  68, 156, 164, 165, 166, 168, 169, 169, 170, 171, 172, 173, 174, 175, 183, 185 Meinertellinae 165 Meinertellus  158, 166, 175 Melanaspis obscura  592 Melangyna 40 Melanoliarus 538 Melanoplinae  250, 268, 269 Melanoplus 269 M. sanguinipes 807 Melanozosteria 369 Melanthripidae 484, 485 Melectini 851, 858 Melipona subnitida 636 Meliponini  66, 658, 858, 861, 862, 880 Melittidae  66, 683, 851, 855, 856, 860, 861 Melittinae 851, 856 Melittini  856 Melittostylops  683, 688 Melitturga clavicornis 688 Melitturgini  856 Melizoderes 514 Melizoderidae  502, 514, 521, 522, 525, 526 Mellierinae  394 Meloidae 886 Melophasma 294 Melyroidea magnifica 359 Membracidae  65, 265, 502, 508, 514, 515, 521, 522, 525, 526, 527, 528, 675, 684, 692 Membracinae  503, 515, 527, 528, 529 Membracini  528 Membracis 526 Membracoidea 521–530

Index

Mendeschilis  157 Mengea (extinct)  682 Mengeidae (extinct)  674, 678, 682 Mengenilla 678, 678, 679, 680, 681, 682 M. chabauti 167 Mengenillidae 167, 674, 674, 675, 677, 678, 679, 680, 681, 682 Menneus unifasciatus 376 Menoponidae  19, 69, 458, 477 Menosca 546 Menoscinae  506, 546 Merope 711 Meropeidae  68, 705, 706, 711 Merostenus 681 Merothripidae  70, 485, 486 Mesepipsocus 445 Mesomachilis  156, 157, 168, 171, 173, 174 M. nearcticus 168, 170 Mesonychographis  162 Mesopsocidae 422, 424, 426, 427, 429, 432, 441, 442 Mesopsocopsis 443 Mesopsocus 442 M. unipunctatus  424, 426, 427 Mesopsychidae (extinct)  756 Mesoraphidiidae (extinct)  69, 74 Mesoveliidae  65 Messor  179, 681 M. barabarus 681 Metagraphitarsus  157 Metahemipsocus 446 Metallyticidae  68, 394, 395, 395, 403, 406 Metallyticus  391, 396, 399, 403, 406 M. splendidus 403 M. violaceus 403 Metamachilis  157 Metcalfa pruinosa 542 Metcalfiella 526 M. vicina  515

Metoligotoma  229, 231, 234 M. brevispina  230 M. incompta 222, 224, 226 M. rileyi  232 Metretopodidae  19 Metrinura  164, 186 Metriocnemus 36 M. ursinus  32 Metriotelura  162 Metrodoridae  69 Metrodorinae 265 Metylophorini 447 Meximachilis  158 Micralymma brevilingua  23, 33, 39 Micrarchus hystriculeus 303 Microchorista 707 Microcoryphia (= Archaeognatha)  61, 73, 168–175, 728, 733 Microembia 238 Microeurybrachys 544 M. vitrifrons  533 Microminae  640 Micronectidae  65 Micropezidae  21, 63, 840, 841, 842, 845, 846, 847, 848 Microphotinini  394 Microphthirus uncinatus 457 Microplitis croceipes 817 Micropterigidae  67 Microtettigoniinae 262 Microthoraciidae  458 Micrutalini  528 Micrutalis 526 Mijas  507 Mileewinae  503 Milichiidae  21, 63 Miobantiinae  394 Miocalles 88 Mioctenopylla arctica 41 Miomantinae  394, 404 Miraculinae 265 Miridae 4, 19, 65 Miroceramia 292 Mirolepisma  160 Mirolepismatinae 182

Mnesicleinae 265 Mogoplistidae  69, 255, 256, 258 Mogoplistinae 258, 684 Molannidae  70 Molopoterus theae 693 Monachina  161 Monastria biguttata 369 Monobelini 529 Monophlebidae  592, 595, 600, 610, 612 Morabidae 253, 256, 265, 266 Morabinae 266 Moreseinae 266 Morica hybrida 813 Mormisma  161 “Mormisma” wygodzinskyi  161 Morsea 266 Morychus viridis 31 Moundthripidae  485 Mufagaa  682, 685 Musapsocidae  421, 423, 425, 431, 437, 438 Musapsocoides 438 Musapsocus 438 M. creole  421 M. huastecanus  423, 425 Muscidae 18, 21, 25, 28, 41, 64 Musoniellini  394 Mutillidae  66 Mycetophilidae  17, 18, 20, 24, 25, 31, 33, 41, 64 Myconinae  504, 537 Mycterothrips 487 Mydidae  64 Myerslopella 525 Myerslopia rakiuraensis  512 Myerslopiidae  503, 508, 512, 516, 521, 522, 524, 525 Myiodactylinae 655 Myiodactylus 655 Mymaridae  22, 66 Myodopsylla 464 Myopsocidae  422, 427, 430, 433, 445, 446 Myopsocus 446

961

962

Index

Myrmeciinae  683, 687, 688 Myrmecolacidae  70, 674, 675, 681, 683, 684, 685, 686, 687, 688, 694 Myrmecolax  683, 685, 686, 687 M. incautus 687 Myrmecophilidae  69, 255, 256, 258 Myrmeleontidae  68, 628, 629, 630, 631, 633, 634, 635, 636, 638, 640, 649, 651, 652, 879 Myrmeleontinae  640, 651 Myrmicinae 179, 683, 686 Myrsidea 477 Mysidia 531, 541 Mystacinobiidae 457 Mythicomyiidae  64 Mythomantis 398 Myzus persicae 604

n

Nabidae  65, 168 Nala tenuicornis  320 Nallachiinae  640, 647 Nallachius 647 N. americanus 647, 649 Namaquaphasma ookiepense  344, 350 Namibmormisma  161 Namkungia  341, 343 N. biryongensis  343 N. magnus  343, 343 Namunukulina  160 N. funanbuli 186 Nannobittacus 709 Nannochorista 707 N. andica  707 Nannochoristidae  706, 706, 707 Nanomantinae  393, 401 Nanophyllium 292 Nanopsocus 435 N. oceanicus  421, 447 Narodona 650 Nasonia 606

Natiruleda  163 Naucoridae  65 Nauphoeta cinerea  374, 376 Nearctopsylla hygini 472 Neatelura  163 Nebkhalepisma  161 Necrosciidae 300 Necrosciinae  284, 285, 295, 297, 299, 300 Neduba extincta (extinct)  90 Neffapini  856 Neghini  641 Neivamyrmex 814 Nematinae 26 Nemestrinidae  64, 754, 756 Nemobiinae 255 Nemoptera sinuata  649 Nemopteridae  629, 630, 631, 634, 635, 638, 641, 649, 652 Nemopterinae  641, 652, 653 Nemoura arctica 38 Nemouridae  19, 69 Neoaliturus N. haematoceps 524 N. tenellus 524 Neoasterolepisma  160, 176, 177, 178, 678 N. palmonii 167 N. santschi 167 N. wasmannii 186 Neobalinae  503 Neobittacus 709 Neochauliodes tonkinensis 73 Neochorista 712 Neococcoidea 611 Neocoelidiinae  503 Neoephemeridae  64 Neofidelia 860 Neohirasea 300 Neolarrini  858 Neolepolepis occidentalis 419, 421 Neoleria prominens  33 Neolinognathidae  458 Neomachilellus  158, 165, 175, 185

N. dominicanus (extinct) 165 Neomachilis  158 N. halophila  172, 173 Neonallachius 647 Neopanorpa 715 Neopasiphaeinae  857 Neophilaenus lineatus  512 Neopromachus 297 Neopsinae  503 Neopsocus 447 Neopsyllinae  465 Neorhagadochir moreliensis 238 Neostylopyga rhombifolia  374, 375 Neothoridae. See Hepialidae Neotrichodectes 461 Nephax 437 Nephotettix 693 Nepidae  65, 886 Nepiomorpha peripsocoides 419, 421 Nepticulidae  67 Nepticulomima 420, 427 Neriidae  64 Nerthridae  65 Nesidiolestes N. ana  141 N. selium  141 Nesomachilis  158 Nesomeinertellus  158 Nesothrips lativentris 495 Nessorhinini 529 Neurochaetidae  64 Neuromachaerota 521 Neuroptera  20, 24, 61, 68, 73, 644–656, 756, 756, 759, 762, 763, 796, 883, 891 Neurostigma 445 Nevrorthidae 629, 631, 633, 634, 638, 641, 653 Nevrorthus 653 Newipsocus termitiformis 442 Nezara viridula 818 Nicoletia  163, 168 N. phytophila  184, 186

Index

Nicoletiidae  68, 159, 165, 166, 166, 167, 176, 177, 178, 179, 180, 182, 183, 184, 186 Nicoletiinae  159, 182, 183 Nicomiinae  503, 515, 527, 528, 529 Nigronia 642 Nilaparvata lugens  509, 535, 536, 540, 691 Nilomantinae  393, 401 Nilomantis 401 Nioniinae  503, 523 Niphargus 137 Nipponaphis monzeni 609 Nipponatelura  163 Nipponatelurina  163 Nipponeurorthus 653 Nirvaninae 523 Nisia 532 N. carolinensis 547 N. nervosa 547 Nocticola australiensis 366 Nocticolidae 362, 363, 365, 365 Noctuidae  22, 25, 27, 35, 67, 72, 811, 817 Nogodina 547 Nogodinidae  506, 533, 534, 535, 536, 547 Nogodininae  506, 547 Nolanomelissini  856 Nolidae  67 Nomada 851 Nomadinae  858, 862 Nomadini  858 Nomiinae  856 Nomioidinae  856 Nosybinae  639, 645 Notarchipsocus 438 N. fasciipennis  420 Noteriades  857 Nothochrysinae  639, 646 Nothoentomum 437 Nothybidae  64 Notioathaumidae. See Eomeropidae

Notiobiellinae  640 Notiophilus aquaticus 37 Notiopsocus 421, 423, 427, 439 Notiopsylla enciari  471 Notiothauma 711 Notodontidae  67 Notoligotoma 236 N. hardyi  224, 226, 229, 234, 236 N. nitens 236 Notoligotomidae  220, 224, 231, 233, 234, 236, 238, 239 Notonectidae  65, 886 Notoptera  61, 73, 637 See also Grylloblattodea and Mantophasmatodea Notozulia 519 Numicia viridis 548 Nycteribiidae 457 Nyctibora N. acaciana 368 N. brunnea 368 N. sericea 368 Nyctiboriinae  363 Nyctiborinae  364, 366, 367 Nymphalidae  21, 28, 36, 67, 72, 757, 762 Nymphalis antiopa 40 Nymphes myrmeleonoides  632, 654 Nymphidae 630, 631, 631, 632, 635, 638, 641, 653, 654, 655 Nyrminae  639, 645 Nysius groenlandicus  33, 35, 36, 37, 40, 41

o Obriminae 292, 293 Obrimini 292 Ochlerotatus nigripes  34, 37 Ochreriades  858 Ochteridae  65 Ocnophilini 294 Ocnophiloidea 293 Ocytata pallipes 324

Odiniidae  64 Odonata  61, 73, 878, 881, 883, 885, 885, 886, 889 Odontoceridae  70 Oecanthinae 255 Oeclidius fraternus  532 Oecophoridae  67, 72 Oedipodinae  252, 253, 268, 269 Oenis 27 Oestridae  21, 42, 64 Olarthrocera  163 Olarthroceroides  163 Olene mendosa 596 Oliarces 650 Oliarus  141, 538 O. koanoa  141 O. lorettae  141 O. makaiki  141 O. polyphemus  141 Oligembia 239 Oligotoma  237, 238 O. nigra 221, 222, 223, 229, 237 O. saundersii 237 Oligotomidae  64, 219, 220, 222, 223, 224, 236, 239 Oliveridia 42 Omalium caesum 28 Ommatidiotinae  504, 538 Ommatissus 549 O. lybicus 548 Ommatolampidinae  251, 269 Ommexecha 270 Ommexechidae  257, 268, 270, 271 Ommexechinae 270 Omolicna joi 541 Oniticellini 110 Onitini 110 Onthophagini 110 Onthophagus O. hecate 108 O. pennsylvanicus 108 O. taurus 108 Onycholepisma arizonae (extinct)  165, 166 Onychomachilis fischeri  165, 166

963

964

Index

Onycta 546 Operophtera brumata  26, 33 Ophiodopelma 447 Opisodasys pseudarctomys  468 Opisthoplatia orientalis 375 Orchopeas caedens 475 Oreophoetini 294 Orgeriinae  505, 541 Orius 495 Ornatilepisma  161 Ornithobius 461 Orobatida 737 Orobittacus 709 Orthacridinae 268 Ortheziidae  19, 600, 611, 612 Orthocladiinae 34 Orthoderellini  394 Orthoderinae  394, 402 Orthophlebiidae 712 Orthoptera  19, 61, 69, 73, 75, 144, 202, 209, 210, 245–271 (chapter 10), 613, 673, 683, 684, 686, 687, 692, 804, 883 Orussidae  67 Oryctopinae 261 Osirini  858 Osiris 284, 858 Osmiini  857 Osmylidae  68, 628, 631, 634, 635, 638, 641, 654, 655 Osmylinae  641, 655 Osmylops 655 Osmylus fulvicephalus 655 Otinotus 526 Otiocerinae  505, 541 Otiorhynchus sulcatus 812 Otocrania 293 Otomantini  393 Otomantis 401 Oulema melanopus 808 Oxaeinae  856 Oxyelaea 404 O. elegans  390 Oxyhaloinae  364 Oxyinae  250, 251, 269 Oxyothespinae  392, 394, 402

Oxypilinae  393 Oxypiloidea 401 Oxypilus 401 Oxyrhachini  528, 530 Oxyrhachis 526

p Pachymantis 401 Pachymorpha 297 Pachymorphinae 297 Pachyphasma 349 P. brandbergense  344 Pachypsylla venusta 603 Pachytroctes 435 Pachytroctidae  419, 421, 424, 426, 428, 431, 434, 435 Paedembiidae  220, 220, 231, 238 Painella simmondsi 546 Palaeomyrmecolax (extinct)  683, 685 Palaeontinidae (extinct)  516 Palaeopsocus 444 Palaeosetidae  67 Paleograssiella chiapanicum (extinct) 165 Palingeniidae  64 Palophinae  285, 294, 295, 297 Palparinae  641, 651 Pamphagidae 252, 257, 264, 268, 270 Pamphaginae 270 Pamphagodes riffensis 270 Pamphagodidae  257, 268, 270 Pamphiliidae  67 Panchaetothripinae  485, 491 Panchlora 368 Panchlorinae  364 Panesthia  361, 369 P. cribata 362 Panesthiidae  63 Panesthiinae  364 Panlepisma  161 Panorpa  713, 714, 715 P. coreana  714 Panorpidae  68, 73, 706, 706, 712, 713

Panorpodes  712, 713 P. paradoxa 712, 713 Panorpodidae  706, 712, 713 Panurginae  856, 859 Panurgini  856 Papilio 71 P. multicaudatus 880 Papilionidae  21, 67, 72, 886 Papilionoidea  872, 874, 878, 880, 883, 884, 885 Parabacillus 290 Parablepharis 399 Paracaeciliidae  421, 424, 426, 429, 432, 439, 440 Paracaecilius 440 Paracentronodus 515 Parachyphoderris 260 Paracolletes  857 Paracorethrura 546 Paracrotelsa  160 Paragioxenos  683, 689 Paragryllinae 255 Parahindoloides  518, 520, 521 P. lumuana 520 Parahiraciini  506, 544 Parainocellia bicolor  654 Paralabis 324 Paralappida 532 Paramachilis  157, 186 Paramantoida 403 Paramorphoscelis gondokorensis 397 Paramuzoa alsopi  361, 367 Paranauphoeta  364 P. formosana 359, 360 Paranauphoetinae  364 Parancyra bivulnerata 542 Paraoxypilinae 391, 393, 397, 401 Parapachymorpha spiniger  284 Paraperla frontalis  127, 133 Parapetrobius  158, 173 P. azoricus 173 Paraphaenocladius impensus  32 Parapsyllinae  466 Parapsyllus longicornis 472

Index

Pararchipsocus 438 Pararhophitinae  857, 860, 861 Parasphaeria boleiriana  361, 369 Paratropes bilunata  368, 376 Paraxenos  683, 689, 690 P. lugubris  681 Parcoblatta  361, 362, 366, 376 Parelectrentomum 437 Parepeolus  858 Pareuchaetes pseudoinsulata 810 Paroligembia 239 Paroxyophthalmus 404 Parthenembia reclusa 236 Parthenolecanium 611 Parthenothrips 491 Parthenothrips dracaenae 491 Patagoniochiloides  158 Patrobus assimilis 37 Paulianiana 525 Pauliniinae 269 Paulininae 250 Pauronychella  163 Pawiloma 513 Pazius 709 Pecaroecidae  458 Pectinariophyes 521 Pectinopygus 461 P. farallonii  457 Pedetontinus  158 Pedetontoides  158 Pedetontus  158, 169, 186 Pedicia hannai  36, 39 Pediciidae  64 Pedicinidae  69, 458 Pediculidae  19, 458 Pediculus 869 P. humanus capitis  457, 473 P. humanus humanus 473 Pelitropis rotulata  533 Pelmatosilpha lenti 370 Peloridiidae 525 Peltonotellini 538 Peltoperlidae  69 Pemphiginae 608 Pemphigus obesinymphae 609

Pemphredoninae 854 Penapini  856 Pentacentrinae 255 Pentathyrsus 439 Pentatomidae  65, 675, 682, 684, 685, 692, 818, 878 Peponapis 815 Perditini  856, 859 Peregrinus maidis  536, 540 Periplaneta 362 P. aboriginea 370 P. americana  369, 370, 373, 374, 375, 376 P. australasiae  374, 375 P. brunnea  374, 375 P. fuliginosa  361, 370, 374, 375 Peripsocidae  421, 422, 424, 425, 429, 432, 436, 441 Peripsocus 441 P. stagnivagus  421, 424, 425 P. subfasciatus  436 Periscelididae  64 Perisphaeria 368 Perisphaeriinae  364 Peritroctes 435 Perkinsiella saccharicida  536, 540 Perlamantinae 391, 393, 395, 397 Perlamantis 397 Perlidae  19, 69, 74 Perlodidae  19 Peromyscopsylla catatina 472 Petalonychia  163 Petridiobius  158, 173 Petrobiellinae  156, 171, 173, 174 Petrobiellus  158 Petrobiinae  156, 171, 173, 174, 183, 186 Petrobius  158, 173 P. brevistylis 173 Phacopteronidae  66, 595 Phaedon amoraciae 31 Phaenopharos 300 Phalangopsinae 255, 258

Phalces 295 Phalixinae  507, 548 Phallopsocus 437 Phaloriinae 255 Phaneropterinae  253, 262 Pharnaciini  284, 285, 296 Pharsalinae  507, 547 Pharsalus 547 Phasmatidae  69, 289 Phasmatinae 297 Phasmatodea  61, 69, 73, 202, 210, 219, 235, 245, 246, 281–304 (chapter 11), 406 Phasmatoptera. See Phasmatodea Phasmida. See Phasmatodea Phasmodinae  251, 262 Phenacinae  505 Phenacococcinae  600 Phenacoleachiidae  612 Phereurhininae  503 Phidippus rimator 109 Philaenus spumarius 509, 519, 520 Philagra cf. parva  512 Philanthinae 854 Philatis 536 Philedaphia 437 Philopotamidae  70 Philopteridae  19, 69, 458, 459, 461 Philopterus  458 Philotarsidae  69, 421, 422, 424, 426, 427, 429, 432, 443 Philotarsopsis 443 Philotarsus 443 P. arizonicus  421, 424, 426, 427 Philya  528 Phlaeothripidae  70, 484, 485, 486, 487, 489, 490, 491 Phlaeothripinae  485 Phobaeticus 282, 284, 296 P. chani  282, 296 P. kirbyi 296 P. serratipes  284

965

966

Index

Phoenicococcidae 595, 612, 613 Phoridae  21, 64 Photinaidae  394, 395, 399, 403, 404 Photinainae. See Photinaidae Photinaini  394 Photiomantinae  394, 403 Photiomantis 404 Phrictus quinquepartitus  533 Phryganeidae  22, 70 Phryganopsychidae  70 Phthanoxenidae (extinct)  674, 682 Phthanoxenos (extinct)  682 Phthiraptera  19, 61, 69, 73, 210, 417, 422, 434, 447, 458–465, 637, 886, 891 Phylliidae  69, 289, 291 Phylliinae  283, 284, 285, 291, 292, 297 Phyllium 283, 291, 292 P. bioculatum  283 P. giganteum  284 Phyllocrania 399 Phyllocraniinae  393, 399 Phyllodromica  363, 367 P. marginata 367 P. megerlei 367 Phyllophorinae 262 Phylloptera. See Phylliidae Phylloscelis rubra 541 Phyllothelyinae  393, 401, 402 Phylloxeridae  599, 602, 608 Piagetiella peralis 462, 463 Pielomastax 265 Pieltainellini 529 Pieridae  21, 28, 68, 72 Pieris napi 40 Piesmatidae  65 Pijnackeria 295 Pineus 598 Pinotini 110 Pintaliini 540 Piophilidae  21, 41, 64 Pipunculidae  21, 64 Pisaurina mira 109

Pissonotus 540 Pityococcidae  595, 612, 613 Planococcus citri 603 Plataspidae  65 Platybrachinae  505 Platybrachus 692 Platycalymma 401 Platycentrini 529 Platycentrus  528 Platycnemididae  69 Platycraninae  297, 299 Platygastridae  22, 66, 229 Platypezidae  21, 64 Platypleura hirtipennis  511 Platypsyllinae 457 Platystictidae  69 Platystoechotes 650 Platystomatidae  64 Platystylea  163 Platyzosteria 369 Plecoptera 18, 19, 23, 24, 27, 28, 31, 34, 61, 69, 74, 77, 203, 210, 397, 734, 886, 891 Plectopterinae. See Pseudophyllodromiinae Plega hagenella 636 Pleidae  65 Plesiodelphacinae  505, 539 Plokiophilidae  65 Plutella P. polaris 31 P. xylostella  40, 807, 811 Plutellidae  21, 807, 811 Pneumoridae 252, 257, 264, 267 Podacanthus wilkinsoni 303 Podoschtroumpfa 546 Podoscirtinae 255 Pogonogaster 406 Pogonogaster tristani 406 Polistes P. carnifex  689, 690 P. dominula  680 P. gallicus 690 P. stabilis 690

Polyancistrinae 262 Polycentropodidae  70 Polyctenidae  65, 457 Polyglyptini  514, 528 Polymitarcyidae  64 Polyommatus 30 Polyphaga saussurei 374 Polyphagidae. See Corydiidae Polyphagoides 365 Polyplacidae  19, 69, 458 Polypsocus 439 P. corruptus 439 P. lineatus  420, 426, 427 Polystoechotes 650 Polystoechotidae. See Ithonidae Polyzosteria P. flavomaculosa 370 P. mitchelli 359 P. obscuroviridis 370 P. pubescens 370 Polyzosteriinae  364, 370 Pompilidae  66 Ponerinae 179, 683, 686, 687 Popa spurca  390 Porisminae  641, 655 Porismus strigatus  654 Porthetinae  252, 270 Potamanthidae  64 Potnia  528 Praedatophasma 347 P. maraisi  344, 350 Praemachilellus  158 Praemachilis  157 Praemachiloides  157 Praetrigoniophthalmus  157 Primacrotelsa  160 Principella  163 Pringleophaga marioni 813 Prioacanthinae 265 Prionoglarididae  420, 422, 424, 428, 431, 434 Prionopelta amabilis 206 Prionotropis hystrix rhodanica 270 Prisopodidae 294 Prisopodinae 300 Prisopus  294, 300

Index

Prispopodidae 294 Pristophora 36 Proatelura  163 P. pseudolepisma 167 Proatelurina  163 Procercopidae  516, 519 Proceroxenos  682, 685 Proconiini 522 Proctolabinae  251, 269 Proctotrupidae  22, 66 Prolepismina  160, 186 Promesomachilis  157, 167, 169, 170, 186 P. hispanica  167, 173, 184 Pronotiopsocus 427, 439 Prophalangopsidae  256, 258, 260 Prophalangopsinae 260 Prophalangopsis 260 Propsocinae 442 Prosapia 519 Prosboloidea (extinct)  516 Proscopiidae  256, 264, 266 Proscopiinae 266 Prosopistomatidae  64 Prosoplecta 367 Prosotropinae  506, 545 Prosphaeropsocus pallidus  424 Prosthecina  163, 179 Protandrenini  856, 859 Protelenchus (extinct)  684, 691 Protelytroptera (extinct)  324 Protepeolini  858 Proteriades bullifacies 852 Protobiellinae  639, 645 Protoculicoides (extinct)  775 Protograssiella 186 Protolepisma tainicum (extinct) 165 Protomeliturgini  856 Protonemestrius jurassicus (extinct)  756 Protoneuridae  69 Protonychella  163 Protophormia terraenovae  33, 37 Protosmylinae  641, 655

Protoxenidae (extinct)  674, 678, 682 Protoxenos (extinct)  682 Protrinemura  161, 186 Protrinemurella  161 Protrinemuridae  68, 159, 176, 177, 180, 181, 182, 183, 186 Protrinemuriodes  161 Protrinemuroides 176 Protroctopsocidae  420, 428, 431, 437 Protroctopsocus 437 P. enigmaticus  420 Proutista moesta  541, 693 Provespa 690 Psammolepisma  161 Pselaphidae 146 Pseudanophthalmus 146 Pseudarchipsocus 438 Pseudatelura  163 Pseudatelurodes  163 Pseudempusa 395 Pseudoanaplectinia yumotoi 366 Pseudobrinckina  164 Pseudocaeciliidae  69, 422, 424, 426, 427, 429, 432, 443, 444 Pseudocaecilius citricola 426, 427 Pseudocatamachilis  157 Pseudoclastoptera 521 Pseudococcidae  19, 66, 595, 600, 612 Pseudococcinae  600 Pseudococcus obscurus 613 Pseudocorethrura 546 Pseudocreobotra  400, 401 Pseudoderopeltis albilatera 359, 360 Pseudodiamesa 42 Pseudogastrotheus  163, 186 Pseudoharpax 400 Pseudoheriades  857 Pseudokolbea 439 Pseudomachaerota 521

Pseudomachilanus  157 Pseudomarava prominens 323 Pseudomeinertellus  158 Pseudomenopon pilosum 474 Pseudomiopteryginae  394 Pseudomyrmecinae  683, 686, 687 Pseudomyrmex 368 P. gracilis 179 Pseudophasmatidae  69, 284, 285, 294, 297 Pseudophyllinae  258, 262 Pseudophyllodromiinae  363, 364, 366 Pseudopogonogastrinae  394 Pseudopolycentropodidae (extinct)  68, 756 Pseudopolycentropus janeannae (extinct)  756 Pseudopsocus 442 Pseudoregma bambucicola 609 Pseudorypteryx 434 Pseudoschmidtiinae 266 Pseudosermyle 294 Pseudosinella 135, 144 P. violenta 134 Pseudosmittia  32 Pseudotyxis 546 Pseudoxenos  683, 689, 690 P. carnifax 689 P. hockeri 689 P. iwatai 689 Pseudoyersinia 395 Psilidae  64 Psilopsocidae  421, 423, 424, 430, 445 Psilopsocus P. mimulus 445 P. nebulosus  421, 423, 424 Psocathropos 434 Psocidae  69, 419, 422, 424, 425, 427, 430, 433, 436, 445, 446 Psocinae 446 Psocini 447 Psococerastis 447 P. fasciata  422, 427

967

968

Index

Psocodea. See Pthiraptera and Psocoptera Psocoptera  23, 25, 61, 69, 74, 199, 210, 417–448 (chapter 16), 484, 637 Psoquilla marginepunctata  421, 424, 426, 434 Psoquillidae  419, 421, 424, 425, 426, 426, 428, 431 Psychidae  68 Psychobiellinae  640 Psychodidae  24, 64, 775 Psychomastax 266 Psychomyiidae  70 Psychophora 36 P. cinderella 35 Psychopsidae  68, 628, 631, 631, 635, 638, 641, 654, 656, 759 Psychopsinae  641, 656 Psychopsis insolens  654 Psyllidae  19, 66, 592, 595, 675, 684, 692 Psyllipsocidae  69, 420, 424, 427, 428, 431, 434 Psyllipsocus 434 P. huastecanus  420, 427 P. maculatus  424 P. ramburii 434 Psylloidea  31, 36, 592, 594, 595, 599, 612, 613 Ptenopsila 439 Pterobrimus 292 Pterochrozinae 253, 258, 262 Pterochrozini 262 Pterolibethra 294 Pteromalidae  22, 38, 66 Pteronarcidae  19 Pterophoridae  21, 68 Pteroplistinae 255 Pterostichus 39 P. brevicornis 92 P. empetricola 92 P. melanarius 111 Pthiridae  458 Pthirus pubis  459, 473 Ptilocerembia  231, 236

P. catherinae 238 P. rossi 238 Ptilocerembiidae  220, 238 Ptilocolepidae  70 Ptiloneuridae  420, 423, 433, 444 Ptomaphagus 137, 138, 139 P. brevior  138, 139 P. cavernicola  138, 139 P. hirtus 137, 138, 138, 139 P. loedingi 137, 138, 139 P. longicornis 137, 138, 139 P. valentinei  138, 138, 139 Ptychopteridae  64 Ptycta 447 Ptyctini 447 Ptyelus goudoti 520 Pulex 464 Pulicidae  21, 69, 465 Pulvinaria urbicola 817 Putoidae  600, 612 Pycnoscelinae  364 Pycnoscelus surinamensis  374, 375 Pygidicranidae  63, 316, 317, 320, 323, 325, 326 Pygiopsyllidae  69, 465 Pyla fusca 28 Pyragropsis 325 Pyralidae  22, 68, 72, 457 Pyrgacrididae  257, 268, 270 Pyrgacris P. descampsi 270 P. relictus 270 Pyrgomantis 404 Pyrgomorphidae  69, 251, 253, 257, 264, 267, 268, 270 Pyrgomorphinae 268 Pyrgotidae  64 Pyrilla perpusilla 546 Pyrrhocoridae  65

q

Quadrinareini 527, 528, 529 Quesada gigas 517

r Ramphomyia 41

Ramsdelepidion (extinct)  159 Ramulus nematodes 296 Raphidiidae  631, 638, 641, 654, 657, 658 Raphidiinae  641 Raphidiini  641 Raphidiomimidae  63 Raphidioptera  61, 69, 74, 77, 657 Rapisma 650 Rapismatidae. See Ithonidae Raptophasma (extinct) R. groehni  344, 351 R. kerneggeri  344, 351 Rasthegotus  163 Ratemiidae  458 Rediviva 851 Reduviidae  65, 168 Reticulopsocus 437 Reuterella 442 Rhachiberothinae 637, 639, 645, 651 Rhadinopsyllinae  465 Rhagadochir virgo  229, 238 Rhagionidae  20, 64 Rhamphixius cf. championi 531 Rhamphophasma spinicorne 284 Rhaphiderus 299 Rhaphidophoridae  69, 73, 250, 251, 256, 258, 260, 870 Rhaphidophorinae 260 Rhathymini  858 Rhiniidae  64 Rhinotermitidae  67, 363 Rhizoecidae  600 Rhodnius 327 Rhopalidae  65 Rhopalopsyllidae  466 Rhopalopsyllinae  466 Rhopalosiphum padi 614 Rhopalosomatidae  66 Rhotanini 541 Rhyacophilidae  22, 70 Rhynchium 690 Rhynchomitra 541

Index

Rhynchophthirina 457, 458, 459, 461 Rhyncophorus ferrugineus 326 Rhyopsocus 434 R. bentonae  419 R. texanus  421 Rhyparobia maderae 374 Rhyparochromidae  65 Rhytidochrotinae 269 Ricania speculum 548 Ricaniidae  65, 507, 533, 534, 535, 542, 544, 547, 548 Ricaniinae  507 Ricinidae  19, 458, 461 Ricinus 464 Riodinidae  68 Ripipterygidae 250, 256, 262 Rivetina 402 Roeslerstammiidae  68 Romaleidae  257, 268, 270 Romaleinae 271 Rophitinae  856 Rophitini  856 Roproniidae  66 Rotoitidae  66 Rotundiceps 460 Rozenia  683, 688 Rugosana querci  513 Rulenatida  163

s

Sabulepisma  161 Sabulopsocidae  429, 442 Saccharosydne saccharivora 540 Saccharosydnini 539 Saginae  251, 262 Sagmation 525 Salda littoralis  33 Saldidae  19, 27, 65 Salticidae 168 Saltoblattella montistabularis 366 Sambini  856 Santhomesiella  163, 178 Sapadrama 846 Sapygidae  66

Sarcophagidae  21, 64 Sarebasa 546 Sarima nigroclypeata 545 Saturniidae  68 Satyrinae  21, 28 Saucrosmylidae (extinct) 762, 763 Saussurembia S. calypso 229 S. davisi 233 Scabina antipoda 370 Scaphytopius nitrides 524 Scapteriscinae 258 Scarabaeidae  109, 318, 457, 804, 882, 886 Scarabaeus sacer 872 Scathophaga S. furcata  33 S. litorea  33 Scathophagidae  21, 25, 33, 64 Scatopsidae  20, 33, 64 Scelembia. See Rhagadochir Scelembiidae  220, 235, 238 Sceletolepisma 183 Scelimeninae 250 Scelionidae. See Platygastridae Scenergates viridis 523 Scenopinidae  64 Sceptrophasma 284, 295, 297 Sceptrophasma hispidulum 284 Schistocerca 753 S. ceratiola 251 S. gregaria 253 Schizocephala 402 Schizocephalinae 391, 392, 394, 402, 404 Schizodactylidae  256, 258, 258 Schizodactylinae 258 Schizodactylus 258 Schizopechus 439 Schizoproreus volcanus  317 Schizopteridae  65 Schrankia S. altivolans  141 S. howarthi  141 Sciaridae  20, 33, 41, 64

Sciomyzidae  21, 64 Scirtothrips dorsalis 495 Scitodidae 168 Sclerodepsa granulosa  533 Sclerogryllinae 255 Sclerophasma 349 S. paresisense  344, 349, 349 Scolebythidae  66 Scoliidae  66 Scolothrips 486 Scolypopa australis  533, 548 Scorpiones 733 Scottiella 443 Scrapterinae  857 Scutelleridae  65, 675, 684, 692, 693 Scythrididae  68 Scytopsocus 443 Segestidea novaeguineae 688 Selenopsocus 438 Selizini 542 Seopsocus 437 S. rafaeli  420, 423, 426 Sepsidae  64 Sepullia 521 Sepulliini 521 Sericostomatidae  70 Sericothripinae  485 Serritermitidae  363 Sesiidae  68 Sevia 537 Sexava 694 Shelfordella lateralis 374 Shelfordina orchidae 367 Shelforditinae 269 Sialidae  631, 634, 638, 642, 642 Sialis lutaria  642 Sibyllinae  393, 401, 402 Signiphoridae  66 Signoretiinae  504 Siinae 261 Silphidae  20 Silvanana 547 Silvestrella  160 Silvestrellatinae  159, 182 Silvestrichilis  157

969

970

Index

Silvestrichiloides  157 Simplocaria metallica 39 Simuliidae  2, 5, 20, 25, 42, 64 Sinella 135 Siniamphipsocus 439 Sinonele (extinct)  351 Sinoneurorthus 653 Sinopanorpa 715 Siphonaptera  21, 23, 61, 69, 74, 465–474, 707, 710, 715, 873, 880, 884, 888, 891 Sipyloidea  297, 300 S. sipylus  284, 300 Siricidae  22, 67 Sisyborina 656 Sisyra 656 S. fuscata  654 S. terminalis  654 Sisyridae  68, 631, 632, 634, 638, 641, 653, 654, 656 Sisyrina 656 Sitobion calvulus  31, 36, 39 Sitodiplosis mosellana 808 Smiliinae  503, 514, 527, 528, 529 Smittia  29, 36, 37, 41 S. brevipennis  32 S. extrema  32, 37 Sogatella furcifera  536, 540, 680, 691 Solenopsis 185 S. invicta  327, 688 Songmachilis  157 Sorellembiidae (extinct)  239 Spathomorpha  285, 295 Spathosterninae 269 Speleketor S. flocki  424 S. irwini  420 Speleonycta  163, 180 Speonomus  128, 146 Spermophorella 642 Sphaeroceridae  21, 33, 64 Sphaeropsocidae  419, 424, 426, 427, 428, 431, 434, 435 Sphaeropsocopsis argentina 426, 427

Sphaeropsocus S. bicolor 435 S. kuenowii 435 Sphecidae  66, 683, 684, 689 Sphecodini  856, 857 Sphenarium purpurascens purpurascens 254 Sphingidae  22, 68, 72, 85 Sphodromantis 402 Spilogona  41, 42 Spilopsyllus cuniculi 473, 475 Spilosmylinae  641, 655 Spinonemia 290 Spissistilus 526 Spongiphoridae  63, 316, 318, 325, 326 Spurostigma 421, 444 Spurostigmatidae. See Cladiopsocidae Squamatinia  164 S. algharbica 180 Squamigera  163 S. latebricola 180 Srokalarva berthei (extinct)  758 Stachilis  158 Staelonchodes harmani  284 Stagmatopterinae  392, 394, 683 Stagmomantinae  392, 394 Stagobius troglodytes 123 Staphylinidae 4, 20, 25, 26, 29, 31, 32, 33, 457 Stegaspidinae  503, 515, 527, 528, 529 Stegaspis  528 Steingeliidae  612 Steleops 447 S. elegans  422 S. lichenatus  422 Stelis 862 Stenocraninae  505, 539 Stenocranophilus  684, 692 Stenomacrus groenlandicus  39, 41

Stenomantis 401 Stenopelmatidae  251, 252, 256, 258, 260, 261 Stenopelmatinae 261 Stenophylla 404 Stenophyllidae  394, 395, 396, 397, 404 Stenophyllinae  396, 404 Stenoponia  464, 468 Stenoponiinae  465 Stenopsocidae  69, 421, 424, 426, 427, 429, 432, 436, 439, 440 Stenopsocus 424, 440 S. nigricellus  421, 426, 427 Stenopsychidae  70 Stenoschmidtiinae 266 Stenosmylinae  641, 655 Stenotritidae 855, 857, 860 Stenurothripidae  485, 486 Stephanacridini  285, 296, 297 Stephanidae  67 Stephanocircidae 464, 466 Stephanocircinae  466 Stephanocircus dasyuri  466 Stephanocleonus 31 Steraspis squamosa 871 Stichotrema  683, 685, 686, 687, 688 S. dallatorreanum 686, 688, 694 Stictocephala 526 Stictococcidae 595, 612, 613 Stictococcus  611, 615 Stigmacoccidae  612 Stigmacoccus garmilleri 593 Stilbopteryginae  641, 645, 651, 652 Stilbopteryx 652 Stimulopalpus japonicus 437 Stivaliidae  69, 466 Stivaliinae  466 Stobaera 540 Stolotermitidae  363, 372 Stratiomyidae  64, 804 Streblidae  64, 457

Index

Strepsiptera  61, 69, 74, 77, 167, 210, 628, 659, 673–694 (chapter 22), 706, 715 Striatophasma naukluftense  344, 350 Strigiphilus 461 Strongylodematinae  505, 543 Stylifera  161, 186 S. galapagoensis 186 S. impudica 186 Styloperlidae  69 Stylopidae  70, 674, 675, 683, 684, 688, 689, 694 Stylops 677, 683, 688, 689 S. pacificus 689 Stylotermitidae  363 Subnicoletia  164 Subnicoletiinae  159, 182, 186 Subtrinemura  164 Sungaya inexpectata  284 Supella longipalpa  362, 366, 367, 374 Suphalomitus 642 Surijokocixiidae 530 Susumanioidea (extinct)  302 Swalepisma  161 Symbittacus 709 Sympherobiinae  640 Sympherobius 658 Symphrasinae  640, 650, 651 Symplecta scotica 37 Synclisis baetica  649 Syrphidae  21, 25, 41, 64, 804 Syrphus 40

t

Tabanidae  20, 64, 884 Tachinidae  21, 41, 64, 324, 840 Taeniapterinae 840 Taeniochorista 712 Taeniostigma 439 Taeniothrips inconsequens 494 Tagalina  318, 323 T. papua 325 Tagalopsocus 439 Tainophasma 296 Talipedini 527, 528, 529

Tambinia 549 Tanaoceridae  256, 264, 266, 267 Tanypezidae  64 Tanzaniophasma subsolanum  344, 349 Tanzaniophasmatidae  339, 344, 349 Taphrotylus 521 Tapinacaena 521 Tapinella 435 T. maculata  419, 424, 426 Tapinotaspidini  859 Tarachina 392 Tarachodes 404 Tarachodidae  392, 394, 395, 400, 402, 404 Tarachodula pantherina 404 Tarragoilus 260 Tarsophilus 443 Tartessinae  504, 524 Teicophryinae 265 Telamonini  528 Teliapsocus conterminus  425, 427, 436, 439 Teloganodidae  64 Temnopteryx phalerata 366 Tenebrionidae  146, 813, 871 Tenthredinidae  22, 23, 25, 26, 28, 67, 72 Tephritidae  64, 675, 684, 692, 693, 846, 847 Teratembia 239 Teratembiidae  64, 220, 239 Teratodinae 269 Terentiini  528, 530 Termitidae  67, 363, 371, 372, 373 Termitoidae  61, 67, 70, 72, 362, 363, 364, 371, 372, 805, 891 Termopsidae. See Archotermopsidae and Stolotermitidae Terrobittacus 709 Tessaratomidae  65 Tetragnatha 88

Tetramorium bicarinatum 817 Tetrapedia 852, 858, 859, 861 Tetrapediini  858 Tetrigidae  69, 250, 251, 256, 263, 264, 265 Tetriginae 265 Tettigarcta 517 T. crinita  511 Tettigarctidae  502, 508, 511, 516, 517 Tettigometridae  507, 533, 534, 535, 548, 675, 684, 692 Tettigometrinae  507, 548 Tettigoniidae  69, 73, 247, 251, 253, 256, 258, 261, 262, 683, 686, 687, 882, 885 Tettigoniinae 262 Texoreddellia  163, 180, 184 Thabena brunnifrons 545 Thaipsocus 438 Thaumaleidae  64 Thaumapsyllinae  465 Theopompa 406 Therea T. petiverana 376 T. petiveriana  361, 362, 365 Therevidae  64 Thericleidae  256, 264, 265, 266 Therioaphis trifolii (f. maculata) 614 Thermobia  161, 176, 177, 183 T. domestica 184 Thespidae  68, 394, 395, 404, 405 Thespinae  392, 394, 402 Thespini  394 Thionia 532 T. simplex 545 Thomomydoecus 461 Thorax porcellana  360, 368 Thrinchinae 270 Thrinchostomini  856, 857 Thripidae  70, 484, 485, 486, 487, 489, 490, 491 Thripinae  485

971

972

Index

Thrips  488, 491 T. palmi  490, 495 T. tabaci  486, 494 Thuridini 527, 528, 529 Thyreus 851 Thyridates 709 Thyrididae  68 Thyrsophorini 447 Thysanoptera  20, 61, 70, 74, 210, 483–496 (chapter 18), 637, 804, 805 Tibicininae  502 Timema  285, 286, 288, 289, 290, 302, 303 T. cristinae  289, 302 Timematodea 286 Timomenus lugens  321 Tineidae  68, 813 Tingidae  65 Tiphiidae  66 Tipula carinifrons  35, 36 Tipulidae  20, 28, 39, 42, 64 Tipuloidea  25, 31 Tisma 402 T. grandidieri  390 Titanacris 271 Tithrone 397 Tivia  365 Tiviinae 363, 365 Tonginae  506, 535, 547 Tortricidae  21, 68, 72 Torymidae  66 Townsendiellini  858 Toxodera 391 T. beieri 405 Toxoderidae  68, 392, 394, 395, 398, 400, 402, 404, 405 Toxoderini 405 Toxoderopsini. See Toxoderini Trachythorax maculicollis  284 Tragopini  528, 529 Trechiama 146 Trechini 142 Trechus pertyi 30 Tremapterus 521 Triarthria setipennis 324

Triassomachilidae (extinct) 164 Triassomachilis uralensis (extinct) 159 Triassothripidae (extinct)  485 Tribolium  753, 795 T. castaneum  758 Tricentrus 526 Trichadenotecnum 447 T. quaesitum  422 Trichatelura  164, 179 Trichocera  33 T. borealis 38 Trichoceridae  20, 27, 33 Trichodectes euarctidos  462 Trichodectidae  19, 69, 458, 459 Trichodimeria  163 Trichogrammatidae  67 Tricholepidiidae  159, 175, 177, 180, 181, 183, 186 Tricholepidion  161, 183 T. gertschi  181, 186 Tricholepisma  160, 178, 678 Trichomatinae  639, 645 Trichopsocidae  421, 422, 424, 427, 429, 432, 443, 444 Trichopsocus 444 T. clarus  421, 424, 427 Trichoptera  4, 18, 22, 23, 24, 27, 28, 34, 37, 61, 70, 74, 77, 886 Trichoscelia 650 Trichotriura  164 Trichotriurella  164 Trichotriuroides  164 Tricorythidae  64 Tridactylidae  69, 250, 256, 263, 264, 675, 684, 692 Tridactylophaginae 692 Tridactylophagus  684, 692 Trienopinae 535 Trienopini 548 Trigona 111 Trigonalidae  67 Trigonidiinae 255 Trigoniomachilis  158

Trigoniophthalmus  158, 172 T. alternatus  171, 185 Trigonopterygidae  257, 264, 267 Trigonopteryginae 267 Trigonoscuta 90 Trilineatoxenos  678, 679, 682 Trimenoponidae  458, 459 Trimerocaecilius 443 Trinemophora  161, 186 Trinemura  164 Trinemurodes  164 T. antiquus (extinct)  165 T. miocenicus (extinct)  165 Trinoton anserinum 474 Triozidae  66, 595 Triozocera  682, 685 Tristiridae  257, 268, 270, 271 Tristirinae 271 Troctopsocidae  428, 431, 437, 438 Troctopsocoides 438 Troctopsocopsis 438 Troctopsoculus 438 Troctopsocus 438 Troctulus 435 Trogiidae  419, 424, 425, 426, 427, 428, 431 Trogium 426 T. pulsatorium 447 Troglophilinae 260 Troglosphaeropsocus voylesi  419 Troglotheus  163 Troglotroctes 435 Trogophilus 870 Tropidacris 271 Tropidaspis 514 Tropidischiinae 260 Tropidocephalini 539 Tropidoderinae  284, 297, 299 Tropidoderus  297, 299 Tropidomantinae  393, 401 Tropidomantini  393 Tropidopolinae 269 Tropiduchidae  65, 507, 533, 534, 535, 537, 545, 548, 549, 675, 684, 692

Index

Tropiduchinae  507, 548 Tropiphlepsia 537 Tryonicidae 362, 363, 364, 371 Tryonicinae  364 Tryonicus  364, 371 Trypetinae  846 Tuberaphis styraci 609 Tunga penetrans  469, 476 Tungidae  466 Turquimachilis  156, 157, 174 Tympanophorinae 262 Typhlocybinae  504, 513, 523, 524 Typophyllum 253 Tyrannophasma 347 T. gladiator  344, 350 Tytthobittacus 709

u Udamoselinae 594 Uenoidae  70 Ugyops 539 Ulidiidae  64 Ulopa U. carneae 509 U. reticulata  509, 692 Ulopinae  504, 522, 523 Ultracoelostoma 593 Umbelligerus woldai  515 Uniclavus  682, 685 Uniptera 537 Uraniidae  68 Urnisiella rubropunctata 250 Urodidae  68 Uropsylla tasmanica 470, 471, 472 Uropsyllinae  465 Urostylididae  65 Uzelothripidae  485, 486

v Valenzuela 440 V. postica  421, 424, 426 Vanduzea 526 Vanessa cardui 40 Varnia 650 Vatinae 391, 392, 394, 402

Veliidae  65 Velocipedidae  65 Velox 402 Venisiella 546 Vermileonidae  64 Vermipsylla alakurt 468 Vermipsyllidae  466 Vespa V. analis 690 V. crabro 690 V. ducalis 690 V. dybowskii 690 V. mandarinia 690 V. simillima 690 Vespidae  22, 67, 72, 683, 684, 689, 845, 874 Vietnamellidae  64 Virgilia 546 V. luzonensis 546 Virgiliini 546 Viridiphasma 350 V. clanwilliamense  340, 344, 346, 350, 351 Viridipromontoxius  682, 685 Vitavitus thulius 31 Vizcayinae  505, 539 Vostox brunneipenis 324 Wygodzincinus  163 Wygodzinskilis  158

X. vesparum  677, 679, 680, 689 Xenozorotypus 199 X. burmiticus (extinct)  210 Xeralictini  856 Xeroderinae  297, 299 Xeroderus 299 Xeromelissinae  857 Xestia aequaeva 35 Xestoblatta X. cantralli 366 X. hamata 366 Xestocephalinae 523 Xiphiopsyllidae  466 Xiphocentronidae  70 Xiphopoeini 530 Xiphydriidae  67 Xolocopinae  859 Xosopharinae  505 Xylica 295 Xylococcidae  612 Xylocopa 71, 852, 861 X. confusa 108 X. dejeani 108 X. nobilis 108 Xylocopinae  859, 861 Xylocopini  859 Xylomyidae  64 Xylophagidae  64 Xyronotidae 252, 257, 264, 266, 267

x

y

w

Xanthocaecilius 440 X. quillayute 440 X. sommermanae  421, 424, 426, 440 Xenidae  674, 675, 683, 684, 688, 689, 690 Xeniinae 266 Xenolepisma  160 Xenopsocus 439 Xenopsylla cheopis  472, 474 Xenos  683, 689, 690 X. hamiltoni 689 X. moutoni 690 X. oxyodontes 690

Yemengenilla  678, 679, 682 Yponomeutidae  68

z Zanna 543 Zanninae  506, 543 Zaprochilinae  251, 262 Zeleja 546 Zelus 111 Zelus renardii 111 Zetoborinae  364 Zeugomantispa minuta  649 Ziartissus  507

973

974

Index

Zophiuma Z. butawengi 546 Z. lobulata 546 Zoraptera  61, 70, 74, 199–212 (chapter 8), 637 Zorotypidae  70, 199 Zorotypus 199 Z. absonus  210 Z. acanthothorax  210 Z. amazonensis  210 Z. barberi  202, 203, 206, 207, 209, 209, 210 Z. brasiliensis  199, 202, 210 Z. buxtoni  210 Z. caudelli  202, 203, 206, 207, 210 Z. caxiuana  210 Z. cervicornis  210 Z. ceylonicus  210 Z. congensis  210 Z. cramptoni  210 Z. cretatus  210 Z. delamarei  203, 209, 210 Z. goeleti (extinct)  203, 210

Z. guineensis  210 Z. gurneyi  203, 206, 207, 209, 209, 210 Z. hainanensis  210 Z. hamiltoni  210 Z. hubbardi  199, 200, 202, 203, 204, 206, 210, 210 Z. hudae (extinct)  199, 210 Z. huxleyi  202, 208, 210 Z. impolitus  202, 203, 205, 207, 210 Z. javanicus  210 Z. juninensis  210 Z. lawrencei  210 Z. leleupi  210 Z. longicercatus  210 Z. magnicaudelli  202, 203, 207, 210 Z. manni  210 Z. medoensis 199, 210, 210 Z. mexicanus  210 Z. mnemosyne  210 Z. nascimbenei  210 Z. neotropicus 203, 210

Z. newi  210 Z. novobritannicus  210 Z. palaeus  210 Z. philippinensis  210 Z. sechellensis  210 Z. shannoni  202, 208, 210 Z. silvestrii  210 Z. sinensis 199, 210 Z. snyderi  199, 203, 210 Z. swezeyi  210 Z. vinsoni  210 Z. weidneri  202, 203, 208, 210 Z. weiweii  210 Z. zimmermani  210 Zulia 519 Zuluia 360 Zygaenidae  68 Zygentoma  61, 73, 144, 160, 175–183, 675, 766 Zygon 521 Zygonini 521 Zygophlebiinae  641, 656 Zygoptera 886

975

Index of non‐Arthropod Taxa Arranged Alphabetically. Note: Page numbers in bold indicate table entries, and numbers in italic face indicate entries on figures and in figure captions.

a Abies balsamea 2 Acacia  368, 487, 490 A. cambagei 492 Acanthocheilonema reconditum 475 Acer  128, 172 A. plantanoides 172 A. saccharum (sugar maple) 494 Ailanthus altissima 543 algae  226, 265 Alnus 26 Annandia 602 A. adelgestsuga  599 A. pinicola  599 Annonaceae 376 Anolis 376 apple (Malus) 483 Aquifoliales  752 Arachis hypogaea  483, 544 Araucariaceae 91 Ardipithicus 869 Arecaceae  545, 546 Arecales  752 Arsenophonus  599, 603 Artemisia 238 Arxula  601 Asclepias 811 Aspalathus linearis 693 aspen. See Populus

Asteraceae  172, 487, 540, 811, 853, 859, 863 Asterales  752 Aureobasidium  601 Australopithecus afarensis 775

Buchnera 602 B. aphidicola 598, 599, 604 Bufo melanostictus 168 Bullera  601 Bunyaviridae 494

b

c

Bacteroidaceae  600 banana (Musa)  435, 483, 494 Bartonella 475 B. henselae 475 B. quintana 473 bat  316, 464 Beamys 315 bean 822 navy (Phaseolus vulgaris) 375 Betaproteobacteria  598, 602 Betula  26, 172 B. nana 34 B. verrucosa 172 Bison 42 Blattabacterium 602 Borrelia recurrentis 473 Bovicola tibialis 474 Brassica 111 B. oleracea 109 Brassicaceae  813, 863 bromeliads 367 Brownia rhizoecola  600 Bryophyta  231, 239, 250, 265, 741

cabbage. See Brassica Capnodium 591 Capparaceae 863 capsicum  483, 495 Cardinium  600, 603 Carex ursina  32 caribou/reindeer (Rangifer tarandus) 42 Caryophyllales  752 Carsonella ruddii  599, 603 Cassiope tetragona 42 Casuarina  236, 489, 521 cat (Felis) 474 cattle (Bos taurus) 463 Cedrelospermum nervosum (extinct)  750 Ceratiola ericoides 251 Cervicola 474 Cetraria delisei  32 cherry (Prunus) 494 chimpanzee (Pan) 869 Chloranthiales  752 Chromolaena odorata 810 Chrysanthemum 487 Cicadophyta  739

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

976

Index

Cinchona pubescens 375 citrus  483, 493, 548, 591 Cladonia 236 Clethrionomys rutilis 41 coconut (Cocos nucifera)  495, 536 coffee (Coffea)  483, 548, 809 coot (Fulica) 474 Cordaites (extinct)  733, 739 Coriaria arborea 548 Cornales  752 Corydalis solida 26 Corylus 172 C. avelana 172 cotton (Gossypium) 375, 483, 548 Coxiella burnettii 475 Cricetomys 315

d Dactylopiibacterium carminicum  601 Damalinia 474 Dasyuridae 470 deer, black‐tailed (Odocoileus hemionus) 474 Deschampsia alpina  32 Dipsicales  752 Dipylidium caninum 475 dog (Canis)  474, 475 Dothieraceae  601 Dryas 42 D. integrifolia 40 D. octopetala 31, 32, 40 dunlin (Calidris alpina) 42

e

Ecksteinia adelgidicola  599 elephant (Elephantidae)  459 Eliomys quercinus 168 Empetrum nigrum 34 Enterobacteriaceae 598, 599, 600, 601, 602, 604 Entomophaga grylli 807 Equisetales  733, 766 Equisetopsida (= Sphenopsida)  739 Ericales  752

Erwinia  600 E. aphidicola  599 Eucalyptus  236, 297, 373, 441, 490, 520, 656 Eulimdana 474

guillemot, Brunnich’s (Uria lomvia) 42 guinea pig (Cavia porcellus) 474

f

Halomonadaceae  599 Hamiltonella defensa  599, 604 hantavirus 477 hare arctic hare (Lepus arcticus) 469 snowshoe hare (Lepus americanus) 42 Hartigia pinicola  599 Heliconia  435, 518, 539 Hirundo pyrrhonota 472 Hoataupuhia coelostomidicola  600 hog, red river (Potamochoerus porcus 459 Holozamites (extinct)  763 Homo erectus 775 horse (Equus ferus) 463 Hymenolepis H. citelli 475 H. diminuta 475 H. nana 475 Hypermastigia 371 Hypocreales  732

Fabaceae  595, 819, 853, 859 Fabales  752 Fagaceae 537 Fagales  752 Fagus 128, 130 ferns  251, 405, 739 ferret, black‐footed (Mustela nigripes) 475 Ficus 820 fig (Ficus carica) 487 filarial nematodes  474 flagellates 371 Flavobacteriia 602 flying squirrel (Glaucomys) 457, 468 fox, arctic (Vulpes lagopus) 42 Francisella tularensis 475 fulmar (Fulmarus) 469 Fusarium 613 F. solani 613

g Gammaproteobacteria 602 garlic (Allium) 494 Garryales  752 geminivirus 591 Gentianales  752 Geraniaceae 754 Gigantopterida  739, 766 Gillettellia cooleyia  599 Ginkgo 707, 741, 745 G. biloba  745 Ginkgoites 762, 763 goose (Anserinae)  474 lesser snow goose (Anser caerulescens) 35 Gorilla 869 grape. See Vitis grasses. See Poaceae Gregarina 324 Gregarinidae 167

h

i Inga 819 Iridaceae 754 Ishikawaella 605

j Jamestown Canyon virus  42 Juncales  752

k kiwifruit (Actinidia) 548 knot (Calidris canutus) 42

l Laboulbeniaceae 168 Lamiales  752 lantana 526 Larrea tridentata 251

Index

Laurales  752 Ledum (section of Rhododendron) 26 Leishmania 775 lettuce (Lactuca) 483 Leucosporidiaceae  601 Leucosporidium  601 Liaoningocladus (extinct)  738 Liberibacter  599, 604 L. psyllaurous 604 lichen  224, 226, 229, 231, 233, 236, 237, 238, 239 Liliaceae  375, 859 Littoraria scabra 821 Lysimachia 852

m Macrotyloma geocarpum 544 Macrozamia 489 Magnoliales  752 maize (Zea mays)  375, 540, 544, 817, 822 Malpighiales  752 Malvales  752 mango (Mangifera) 483 maple. See Acer Marattiales (tree ferns)  733 Medullosales (seed ferns)  733, 739, 766 Megaderma lyra 376 Mermithidae 167 Metzgeriothallus sharonae  739 mink (Mustela vison) 475 Moranella endobia  600, 602, 603 Moricandia moricandoides 813 moss. See Bryophyta Mustelidae 472 Myodes gapperi 473 Myrtales  752

n nectarine (Prunus persica)  483, 494 Nilssonia (extinct)  763 Nyctaginaceae 817 Nymphaeales  752

o oak. See Quercus oil palm (Elaeis) 548 onion (Allium cepa)  483, 494 Ophiocordyceps unilateralis 732 Ophiocordycipitaceae 767 Opuntia 658 Orchidaceae  367, 754 Oryctolagus cuniculus 473 Oryza sativa 109 otter, North American river (Lontra canadensis) 463 oxeye daisy (Leucanthemum vulgare)  484 Oxymonadida 371

p Paleohaematopus 775 Pallenis 172 palm  299, 375, 538, 546 Pandanus 299 Pantoea agglomerans  599 Parnassius mnemosyne 26 Parvileguminophyllum coloradoensis (extinct)  750 peanut. See Arachis Pedicularis 42 Pelecitis fulicaeatrae 474 pelican (Pelicanus)  463 Peltaspermales (extinct)  739, 766 penguin, white‐flippered (Eudyptula minor albosignata) 472 Penicillium chrysogenum 179 Peromyscus maniculatus 472 Phalaropus 42 Phoenix 486 P. dactylifera 548 phytoplasmas  524, 536, 539 pig (Sus) 463 Pinaceae 537 Pinophyta  739, 766 Pisonia grandis 817 Planchonella 491

Platanaceae  743 Platanus raynoldsi  743 Pleurococcus 172 Poaceae  179, 251, 487, 523, 541, 545 Poales  752 poinsettia (Euphorbia pulcherrima) 375 Polyalthia longifolia 494 Polyporales 537 Populus  172, 375, 750 P. tremula 172 P. wilmattae  750 Portiera aleyrodidarum  599, 603, 604 potato. See Solanum tuberosum Profftella 603 P. armatura  599, 603, 604 P. tarda  599 P. tarda  599 progymnosperms  745 Proteales  752 Proteus anguinus 123 Prototaxites (extinct)  765 Psammodromus hispanica 168 Psaronius (extinct)  737, 738, 766 P. chasei  737, 738, 766 Pseudomonadaceae  599 Pseudomonas 603 P. adelgestsugas  599 Pseudotsuga menziesii 186 Pteridophyta  251, 405, 739

q Quercus 593 Q. alba 809

r rabbit (Leporidae)  473, 474 See also hare eastern cottontail (Sylvilagus flordianus) 463 Ranunculales  752 Ranunculus acris 40 rat (Rattus) 474 Regiella insecticola  599, 604

977

978

Index

Rhizobiaceae  599 Rhodocyclaceae  601 Rhodotorula  601 rice  483, 519, 540, 547 Ricketsiella 167 Rickettsia  600, 603 R. felis 475 R. prowazeckii 473 R. typhi 475 Rickettsiaceae  600, 601 rodents 464 Rosales  752 rose (Rosa) 375 Rutaceae  743

s

Saccharomycetaceae  601 Salicaceae  750 Salix  26, 35, 172 S. caprea 172 S. polaris 31 sandpiper, purple (Calidris maritima)  41, 42 Santalum album  542, 545 sap sucker (Sphyrapicus) 813 Sapindales  752 Sarconema eurycerca 474 Saxifraga oppositifolia  32, 40 Saxifragales  752 Scinax ardeous 376 seal (Pinnipedia)  463 seed ferns (Spermatophyta)  733, 739, 766 Serratia 603 S. symbiotica  599, 604 sheep (Ovis) Dall sheep (O. dalli) 42 domestic sheep (O. aries) 463 shrew (Soricidae)  464, 472 sloth 457 snow bunting (Plectrophenax nivalis) 42 Sodalis 603 Solanum tuberosum  109, 483 Soldalis  600 Solenodontidae 775

Sorex araneus 41 sorghum (Sorghum bicolor) 375 southern beech (Nothofagus)  593, 711 soybean (Glycine max) 375 Speleomantes italicus 168 Sphenopsida. See Equisetopsida Spiroplasma S. citri 524 S. kunkelii 524 Sporobolomyces  601 Steffania adelgidicola  599 strawberry (Fragaria)  483, 487, 494, 520 sugarcane (Saccharum) 483, 494, 519, 540, 548 Sulcia 509 S. muelleri 602 sunflower (Helianthus) 110, 375 swine (Sus) 463

t tea (Camellia sinensis) 483 Termitomyces 372 Thoropa miliaris 376 tobacco (Nicotiana tabacum)  270, 375, 544 Toddalia schaarschmidti  743 tomato (Solanum lycopersicum)  483, 817 tree ferns  733 Tremblaya T. phenacola  600 T. princeps  600, 602, 603 Tremellaceae  601 Triceratops (extinct)  488 Trypanosoma 475 T. lewisi 475 turnstone (Arenaria) 42

u Ulmaceae  750 Ursus U. americanus (black bear)  462

U. arctos 473 Uvaria elmeri 376 Uzinura diaspidicola  600

v Vaccinium  26, 37, 107, 483, 541 V. vitis‐idaea 40 Vallotia V. cooleyi  599 V. tarda  599 V. virida  599 Vanilla 483 Vidania 509 Vigna subterranean 544 Violaceae 36 viruses 477 Vitales  752 Vitis  536, 543, 545

w Walczuchella monophlebidarum  600 warthog, desert (Phacochoerus aethiopicus) 459 watermelon (Citrullus lanatus) 110 wheat (Triticum) 375 Wolbachia  600, 601, 602, 603, 615 woodpecker, red‐cockaded (Leuconoptopicus borealis) 376

x Xylella fastidiosa 519

y Yersinia pestis 474 Yimaia capituliformis (extinct) 707, 763 Yucca elata 367

z Zingiberales  752 Zosterophyllopsida (extinct)  744

979

Subject Index Note: Page numbers in bold indicate table entries, and numbers in italic face indicate entries on figures and in figure captions.

a acoustic communication  235, 251, 255, 258, 259, 260, 261, 267, 270, 292, 346, 349, 350, 516, 525, 545, 659 adaptation  35, 88, 134, 810 adventive species  93, 327, 367, 425, 441, 446, 447, 520, 521, 538, 542, 543, 545, 548, 593, 805, 818 Afghanistan 238 Africa  180, 185, 229, 235, 236, 238, 239, 248, 251, 253, 254, 259, 260, 265, 266, 267, 268, 269, 285, 294, 295, 297, 315, 325, 343, 345, 363, 365, 368, 369, 370, 371, 373, 397, 398, 399, 400, 401, 406, 434, 435, 441, 442, 443, 446, 488, 495, 517, 521, 535, 538, 541, 542, 544, 546, 548, 642, 645, 646, 647, 649, 652, 653, 656, 657, 658, 659, 685, 687, 688, 693, 707, 802, 822, 855 Afrotropical Region  186, 335, 397, 401, 405, 430, 434, 439, 443, 445, 446, 447, 520, 523, 537, 542, 543, 544, 545, 547, 548, 549, 636, 644, 645, 651 aggressive behavior  368 agricultural ecosystems  105, 801 Alaska [United States of America]  15, 28, 29, 30, 34, 35, 40 Alberta [Canada]  341 Aleutian Islands [Alaska, United States of America] 92 Alexander, Charles P.  3 allergens  185, 375, 447 alpine/montane habitats  251

Alps (Europe)  173 Altai‐Sayan mountains  336 alternation of generations  608, 609 amateur entomologists  283, 303, 447, 848 Amazon basin  488 amber and copal  300, 324, 406, 435, 437, 519, 674, 685, 687, 691, 712, 724, 729, 733, 773, 774 Baltic  165, 166, 186, 300, 302, 344, 351, 519, 674, 682, 683, 685, 688, 712, 713, 728, 729, 735 Burmese  164, 165, 239, 301, 682, 774, 775 Dominican 165, 166, 239, 300, 676, 683, 691, 735, 775 East Prussian  444 Fushun  683, 685, 687 Jordanian 199 Lebonese 164 Mexican  165, 444 New Jersey  164 American Insects [web site]  843 American Philosophical Society 6 Anatolia 645 Anatomy Ontology (HAO)  791 Andes Mountains  173, 234 Angola  186, 239, 439, 444 ant attendance  523, 527, 608 Antilles  437, 438, 439, 775 aposematic coloration  251, 268, 271, 294, 370, 519, 520 aquatic insects  34, 38, 518, 629, 634, 642, 644, 647, 653, 655, 656

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

980

Index

Arctic region  15, 18, 688, 735 Argentina  158, 161, 250, 290, 433, 517, 526, 647, 688, 693, 707, 861 arrhenotoky 611 art, literature and religion, insects in  253, 326, 659, 869–896 (chapter 29) Ascension Island [United Kingdom, overseas territory] 435 asexual reproduction  36, 92, 171, 229, 236, 238, 282, 302, 303, 509, 610, 613, 614, 686, 693 Asia  237, 248, 254, 259, 369, 371, 398, 399, 425, 434, 439, 440, 441, 443, 445, 446, 447, 475, 645, 646, 653, 687, 688, 707, 708 Asia, central  269, 371 Assam [India]  322 Australasian Region  297, 430, 520, 521, 542, 543, 549 Australia  142, 146, 185, 186, 234, 236, 237, 250, 260, 261, 262, 266, 297, 299, 303, 319, 365, 369, 370, 371, 375, 376, 397, 398, 440, 441, 443, 445, 446, 488, 491, 495, 511, 517, 521, 523, 525, 535, 537, 538, 543, 544, 546, 637, 642, 645, 646, 650, 651, 653, 655, 656, 679, 685, 687, 688, 689, 691, 692, 706, 707, 709, 711, 855, 860 Austria  173, 179, 180, 512 Azores [Portugal]  83, 92, 173

b bacteria, as pathogens  524 alfalfa dwarf  519 almond leaf scorch  519 bacterial leaf scorch  526

bois noir grapevine yellows 539 citrus stubborn disease  524 corn stunt  524 curcubit bacteria wilt  774 lethal yellowing pathogen  536, 539 olive quick decline syndrome 519 spike disease (of sandalwood)  542, 545 bacteria, endosymbiotic  598 Gammaproteobacteria  602 Flavobacteriia  600, 601, 602, 613 bacteriocytes  596, 598, 602, 603, 604, 605, 615 Baja California [Mexico]  186 Balbiani, Édouard‐Gérard  4 Ball’s Pyramid [Australia]  298 banana (Musa)  483, 494 Banks, Joseph  81 Barberini, Maffeo (Pope Urban VIII)  884, 884 Barcode of Life Data Systems 5 barcodes, DNA  5, 24, 819 Bates, Henry  6 beneficial insects  105 Bering Land Bridge  30 Bering Strait  15, 27, 186 Beringia  25, 30, 31, 173 Bermuda 529 Bernhauer, Max  4 Bhutan 715 biodiversity hotspot  76, 93, 351 biological control  109, 110, 111, 646, 647, 649, 658, 659, 688, 693, 809, 817 blood feeding  41, 457, 775 Bolivia 709 Borneo  180, 283, 296, 300, 375, 376, 546 Bosnia and Hercegovina  142, 143, 144 Botswana 183

brachypterous insects  39, 440, 441, 523 Brazil  126, 134, 199, 202, 208, 319, 369, 375, 376, 434, 439, 440, 488, 533, 535, 637, 644, 678, 709, 822 brochosomes 523 brood parasitism. See kleptoparasitism Brooks Range [Alaska, United States of America]  28 Brueghel the Elder, Pieter  883 BugGuide [web site]  447, 843 Burma  322, 546 Bylot Island [Canada]  42

c calcrete aquifers  127, 128, 142 California [United States of America]  173, 237, 336, 341, 495, 545, 709, 710, 712 Cambodia 59, 531, 692 Cameron, Malcolm  4 Cameroon 521 camouflage  235, 253, 261, 281, 291, 292, 297, 395, 396, 401, 404, 405, 527, 707, 762, 763 Canada  15, 34, 175, 302, 336, 439, 440, 650, 658, 714, 735, 745 Canadian Journal of Arthropod Identification 845 Canary Islands [Spain]  93, 128, 131, 141, 143, 146, 186, 322, 535, 595 Cape Floristic Province  754 Carabidae of the World [website] 840 Carboniferous Period  159, 351, 753 Caribbean region  235, 294, 296, 529, 537, 545, 594 Caribou/reindeer  42, 43, 870 Carvalho, Jos&eaucte; C. M.  4

Index

caves and cave dwelling arthropods  89, 123, 124, 172, 175, 179, 180, 185, 250, 259, 322, 338, 365, 368, 369, 417, 535, 538, 545, 547 Cenozoic era  628, 723, 735, 753, 863 Central America  248, 261, 265, 269, 271, 294, 296, 367, 370, 371, 399, 437, 438, 439, 441, 444, 519, 521, 537, 545, 594, 650, 708, 709 Cercopoidea Organised On Line (COOL) 510 Chad 774 Chile  290, 437, 444, 514, 515, 525, 645, 647, 650, 707, 709, 711, 861 China  59, 62, 128, 156, 157, 168, 186, 210, 236, 258, 260, 269, 283, 290, 296, 301, 342, 343, 343, 345, 351, 375, 376, 437, 438, 439, 440, 441, 442, 443, 444, 446, 510, 515, 517, 519, 521, 530, 538, 540, 543, 544, 545, 546, 548, 549, 637, 642, 649, 650, 653, 674, 685, 692, 706, 708, 709, 712, 714, 715, 724, 734, 735, 738, 743, 745, 756, 762, 763, 802, 819, 822, 863, 871, 876 Christmas Island [Australia]  93 Chukotka Peninsula [Russia] 28 citizen‐science 848 cleptoparasitism. See kleptoparasitism climate change  15, 42, 93, 185, 546, 765, 775, 801, 863 climatic optima  723, 862 Climatic Relict Hypothesis  141 CLIMEX [modeling software] 806

Colombia  231, 234, 283, 515, 709, 740 colonization. See migration and dispersal colony collapse disorder  895 Colorado [United States of America]  239, 676, 687 commensalism  179, 457, 473, 681 community structure  112 Congo, Democratic Republic of  376 conservation  327, 376 convergent evolution  89, 406, 756 Cook, James  81 Cooks Islands  92 corn, sweet. See maize Cornwallis Island [Canada]  26 Costa Rica  207, 233, 376, 512, 518, 532, 533, 649, 709, 820, 821, 841 courtship. See reproductive behavior Cretaceous Period  239, 258, 268, 301, 302, 324, 351, 508, 516, 519, 523, 628, 674, 705, 706, 711, 712, 723, 729, 734, 735, 738, 753, 759, 767, 769, 773, 774, 855, 862, 863 Cretaceous–Tertiary extinction. See extinction events (K–Pg) Critical Ecosystem Partnership Fund (CEPF)  76 Croatia  124, 142, 146 crypsis. See camouflage cryptic species  820 Cuba  126, 322 Cyprus 437

d Darwin, Charles  6, 81 database  396, 510, 511, 523, 597, 840 da Vinci, Leonardo  883

decomposers  109, 199, 820 defensive behavior  324, 603 DeLong, Dwight  4 deserts  23, 32, 40, 82, 89, 179, 234, 250, 267, 322, 359, 363, 365, 391, 400 detritivore  226, 338, 483 Devonian period  159, 726, 728, 733, 746, 760, 765 Devon Island [Canada]  28 digital insect collections  789, 839, 840, 841, 847 diseases of animals avian malaria  818 hantavirus 477 Jamestown Canyon disease 42 leishmaniasis 775 malaria 774 myxomomatosis 475 nagana 774 Northway Virus  42 diseases of plants alfalfa dwarf  519 almond leaf scorch  519 bacterial leaf scorch  526 barley yellow dwarf  769, 774 black wood  536 bois noir grapevine yellows 539 citrus greening  591 citrus stubborn  524 corn stunt  524 curcubit bacteria wilt  774 Dutch elm disease  774 Finschhafen disorder of oil palms 546 lethal yellowing  536, 539 olive quick decline syndrome 519 pseudo‐curly top of tomato 526 red ring Disease  774 spike disease (of sandalwood)  542, 545 tospovirus diseases  483, 486, 494

981

982

Index

dispersal. See migration and dispersal distribution, geographic  160, 806, 815 DNA sequencing  5, 15, 17, 210, 335, 347, 350, 594, 595 See also barcodes, DNA DNA, complementary (cDNA) 226 Dominican Republic  690 Dryad Digital Repository 796 Dürer, Albrecht  884

e Early Cenozoic Thermal Maximum (ECTM)  750 Early Eocene Climatic Optimum  723, 862 ecosystem services  107 ectoparasites  41, 457–479 (chapter 17) ectosymbionts  315, 457 Ecuador  168, 231, 234, 239, 514, 515 Egypt  325, 494, 861, 871 Ellesmere Island [Canada]  28, 40, 42 Encyclopaedia Biospeologica  144 endangered species  270, 376 Endeavour, HM Bark 81 endemism  31, 91, 351, 547, 708, 709, 710, 712, 714, 815, 821, 852 endosymbiosis  372, 457, 461, 509, 510, 598 entomophagy (by humans)  74, 254, 375, 658, 870, 882 Eocene epoch  239, 300, 302, 508, 518, 674, 687, 723, 730, 735, 747, 761, 762, 862 epikarst. See karst landscapes Equilibrium Theory of Island Biogeography 82 Ethiopian Region. See Afrotropical Region

Europe  125, 260, 326, 399, 425, 434, 439, 440, 441, 443, 446, 447, 495, 642, 646, 685, 692, 707 evolution  89, 535, 597 extinct and fossil species  29, 159, 166, 181, 239, 291, 300, 323, 324, 347, 351, 406, 485, 517, 518, 530, 604, 655, 674, 678, 685, 687, 691, 706, 707, 715, 723, 775 extinction events Cretaceous–Paleogene crisis (K–Pg)  726, 735, 747, 770, 771, 775, 861 Permian–Triasic (P–Tr) crisis  724, 726, 729, 734, 735, 747, 749, 770, 771, 775

f Fiji  92, 292, 297, 816 filarial nematodes  474 Finnmark [Norway]  18, 31 Flickr photograph sharing web site 843 flightlessness  91, 370, 524 Florida [United States of America]  84, 179, 237, 250, 402, 439, 441, 495, 521, 541 forest canopy  251 fossiliferous deposits Cerrejón Formation  740 Colwell Creek Pond Formation  742 Dakota Formation  753 Florissant Formation  239, 729 Green River Formation  676, 683, 687 Jiulongshan Formation  773 Karabastau formation  734 Messel deposit  732, 737 Patagonian Eocene deposits  742

Rhynie chert deposit  733 Yixian Formation  734, 773 fossil insects. See extinct and fossil species France  132, 146, 159, 270, 397, 870 Franz Josef Land [Russia]  28 French Polynesia  93 Fulgoromorpha Lists on The Web (FLOW)  511 fungal parasites of insects  167, 759 fungivores  509, 540

g Gabon 173 Galápagos Islands [Ecuador]  81, 83, 93, 141, 146, 186, 233, 322, 535, 537 gall induction  487, 523, 596, 608, 609 Gaspé [Québec, Canada]  159 generalist  35, 813 generation time  362 gene transfer  288 Germany  319, 676, 685, 743, 746 Ghana 397 Girault, Alexandre A.  4 glaciation  15, 92, 523, 772 glossary 792 Glow Worm Cave [New Zealand] 146 Gondwana  85, 185, 186, 233, 260, 263, 525, 707 grassland  249, 250, 538, 540, 747, 769, 770, 852, 853 Greece  437, 871 Greenland  15, 28, 30, 31, 34, 35, 41, 84, 488 grooming 206 Guadeloupe 848 Guam 811 Grundzüge einer Theorie der phylogenetischen Systematik 6

Index

h habitat disturbance  35, 93, 146, 802 habitat restoration  112 hantavirus 477 Harris, Thaddeus  2 Hawa’ii [United States of America]  85, 87, 91, 92, 93, 126, 132, 141, 146, 185, 322, 535, 538, 540, 816, 817 Hennig, Willi  6 herbivory 813 hermaphroditism 610 Herschel Island [Canada]  42 High Arctic  15, 735 Holocene Epoch  15, 724, 726 Honduras 690 honeydew  591, 593, 596 Hooke, Robert  1 host specificity  302, 549, 596, 597 HOSTS (database of the world’s Lepidopteran hostplants) 597 Hox gene  757 Huautla cave [Mexico]  180 Hudson Bay [Canada]  15, 27, 42 Hustache, Adolphe  4 hybridization 88

i Iceland  15, 86, 319 Idaho [United States of America] 341 identification keys  6, 174, 175, 181, 362, 524, 645, 646, 649, 650, 844 Illinois [United States of America] 733 immature insects  629, 632, 634, 635, 636, 708 immature stages  633, 673, 677, 677, 680, 681, 685, 686, 687, 689, 690, 691, 692, 693

iNaturalist citizen‐science network 848 India  185, 239, 258, 259, 260, 266, 296, 299, 373, 376, 400, 406, 440, 488, 494, 521, 541, 546, 692, 715, 811 Indian Ocean islands  425 indicator species  549 Indo‐Burma Hotspot  76 Indochina  59–78 (chapter 3), 708, 709, 715 Indomalayan Region. See Oriental Region Indonesia  267, 316, 440, 488, 542, 712, 715 industrialization 802 Inner Mongolia [China]  239, 351 inquline species  258 insects as food. See entomophagy integrated pest management 817 Intergovernmental Panel on Climate Change (IPCC)  801, 816 International Code of Zoological Nomenclature (ICZN) 7 International Cooperation Unit on Biodiversity and Environment Conservation (ICUBEC) 77 International Union for Conservation of Nature (IUCN)  270, 376 interstitial habitats  126 Ireland 816 islands  81, 82, 810, 815 Israel 495 Italy  126, 437, 548, 743

j Jamaica  439, 535, 541, 848 Jamestown Canyon Virus  42

Japan  92, 128, 296, 336, 343, 375, 439, 521, 692, 708, 709, 712, 817 Java [Indonesia]  495, 546, 692 Juan Fernández Islands [Chile]  92, 525 Jurassic period  239, 266, 336, 351, 516, 519, 523, 628, 705, 706, 707, 711, 726, 729, 734, 759, 761, 762, 767, 769, 773

k karst landscapes  123, 126, 128 Karst Research Institute of Slovenia 125 Kazakhstan  706, 861 Kentucky [United States of America]  143 Kenya 322 Kerguelen Islands [France, overseas territory]  488 kleptoparasite  487, 852 kleptoparasitism 861 Knight, Harry H.  3 Komsomolets Island [Russia] 23 Korean peninsula  296, 319, 336, 343, 709, 714 Korea, South  343, 543 Krakatoa Island [Indonesia]  86 K–T event. See Cretaceous– Paleogene event Kyrgyzstan 735

l Laos 59 Lapland Fininsh  28, 34 Swedish 34 Laurasia 186 lava tubes  141, 146 le Clerc, Guillaume  879 Liaoning [China]  738 Linné, Carl von (Linnaeus)  627 literature. See art, literature and religion

983

984

Index

liverwort (Hepatophyta)  746 lizard (Sauria)  376 longevity 362 Lord Howe Island [Australia] 298 lycopods (Lycopodophyta)  733, 740, 766

m Mackenzie River [Canada]  27 macroptery 252 Madagascar  82, 85, 91, 93, 185, 258, 265, 266, 289, 292, 294, 295, 300, 365, 488, 520, 525, 538, 543, 544, 547 Madeira [Portugal]  186, 817 Magnetic Island [Australia] 226 maize  373, 375, 519, 544, 693, 804 Majorca. See Mallorca Malaise, René  4 Malawi  345, 349, 521 Malaysia  85, 124, 260, 292, 297, 316, 445, 511, 520, 521, 533, 546, 685, 847 Maldives 816 Malicky, Hans  4 Mallorca [Spain]  146 Malpighian tubules  518, 522, 632 Mammoth Cave [Kentucky, United States of America] 143 mangrove 86 Marion Islands (Crozet Islands) [France, overseas territory] 813 Marquesas [France, overseas territory] 83 Martinique [France, overseas territory] 848 Mascarene Islands  83, 270, 297, 299, 300 maternal care. See parental care Mauritius  270, 299

medicine, use of insects in  375 Medieval Warm Period  29 Mediterranean region  186, 237, 397, 437, 494, 642, 653, 678, 772 meiosis 610 Mekong River  59 melanism 37 Merian, Maria Sibylla  883 Mesoamerica. See Central America Mesozoic Era  185, 186, 628, 729, 753, 757, 761, 767 metamorphosis 629 Mexico  175, 180, 184, 233, 238, 266, 267, 322, 434, 437, 438, 439, 440, 441, 443, 444, 510, 521, 529, 535, 537, 538, 650, 658, 685, 689, 708, 709, 714, 807, 811, 822 microptery 435 Middle Eocene Climatic Optimum 862 migration and dispersal  39, 84, 86, 91, 138, 142, 253, 434, 509, 807, 811 Milieu souterrain superficiel (MSS) 128 mimesis. See camouflage mimicry  281, 367, 762 Miocene Epoch  30, 239, 335, 546, 769 Mississipian Subperiod  726, 728, 733 mitochondrial genes/ genome  183, 210, 219, 347, 350, 594, 595, 820 COI  183, 347, 350 modeling  374, 806, 807, 811 Moffen, Spitzebergen [Norway] 29 molecular data  246, 347, 628 Moluccas [Indonesia]  81, 290 Monarch Butterfly Biosphere Reserve 811

Mongolia 706 Montana [United States of America] 341 Montenegro 142 Morocco 861 Muséum Nationale d’Historie Naturelle 6 Museum of Comparative Zoology Entomology Type Collection Entomology Type Collection 840 Myanmar  59, 239, 267, 674 mythology. See art, literature and religion myxoma virus  475

n   179, 180, 239, 250, 269 Namibia  179, 186, 269, 343, 349, 350, 434, 544, 861 National Institute of Biological Resources (NIBR)  77 Neartic Region  186, 325, 335, 399, 521, 527, 529, 537, 541, 543, 545, 548, 549, 637, 685 Neogene Period  726, 729 neoteny  372, 441, 685, 693 Neotropical Region  186, 359, 398, 399, 402, 405, 425, 435, 444, 445, 446, 447, 491, 521, 523, 527, 529, 537, 542, 543, 545, 547, 549, 644, 647, 688, 689, 851 Nepal  173, 179, 715 Networked Organisms and Habitats (NOAH)  848 New Caledonia  82, 90, 91, 92, 289, 297, 299, 371, 525, 535, 816 New Guinea  285, 297, 298, 299, 397, 443, 445, 488, 542, 543, 545, 649, 655, 691 See also ­Papua‐ New Guinea

Index

New York [United States of America]  531 New Zealand  90, 92, 93, 146, 186, 260, 261, 285, 289, 297, 302, 303, 443, 444, 446, 472, 495, 525, 533, 544, 593, 645, 707, 816 Nicaragua 319, 512, 532 Nigeria 373 North America  185, 260, 261, 271, 289, 303, 326, 338, 425, 435, 440, 441, 442, 443, 444, 446, 447, 472, 473, 495, 517, 642, 646, 650, 658, 709, 807, 811 North Carolina [United States of America] 107, 210, 533, 685 North Dakota [United States of America] 746 Northway Virus  42 Norway  26, 31, 41 nuclear genes 18S rRNA  183, 210 28S rRNA  183, 210 nunataks 30 Nunavut [Canada]  34

o Oceania  425, 446, 520, 544, 549, 637, 688 Ogalalla Aquifer  127 Oligocene Epoch  516, 711, 712, 769 Oman 186 On the Origin of Species 6 ontogenetic development  757, 791 Oregon [United States of America] 300, 341, 710 Oriental Region  186, 294, 297, 430, 520, 521, 527, 537, 539, 542, 543, 544, 545, 547, 548, 549, 636, 644, 647, 655, 656, 861 ovoviparity  361, 368

p paedomorphism. See neoteny Pakistan 400 Palaearctic Region  186, 235, 325, 401, 430, 530, 539, 541, 544, 545, 548, 549, 636, 647, 689, 861 Paleocene Epoch  516, 518, 687, 863 Paleocene–Eocene Thermal Maximum (PETM)  723, 747, 749, 770, 772, 775 Paleodictyoptera  729, 734, 773 Paleogene Period  437, 444, 726, 729, 757, 769, 774 Paleozoic Era  726, 728, 749, 753, 765 Panama  173, 207, 239, 488, 521, 650, 690 Papua New Guinea  86, 263, 266, 322, 521, 546, 686, 688 parasites and parasitoids  41, 281, 324, 673, 690, 692, 759, 767, 852 parasitoid‐protection services 604 parenchyma 596 parental care  224, 229, 235, 238, 258, 315, 323, 324, 327, 359, 362, 368, 371, 490, 523, 525, 526, 527, 762 parthenogenisis. See asexual reproduction Paternal Genome Elimination (PGE)  611, 615 Pbase photograph sharing web site 843 Pennsylvania (United States of America) 543 Pennsylvanian Subperiod  723, 726, 728, 729, 733, 737, 738, 753, 757, 766 Pergande, Theodore  2 permafrost 24

Permian Period  159, 260, 263, 324, 361, 501, 516, 530, 628, 705, 726, 728, 729, 738, 742, 746, 749, 753, 757, 760, 766, 771 Permian–Triasic (P–Tr) extinction 724 Peru  173, 186, 438 pesticides 112 pest insects  113, 253, 269, 326, 366, 370, 373, 374, 447, 493, 494, 517, 519, 520, 536, 539, 541, 543, 547, 548, 593, 688, 805, 811 pets, insects as  376 PHASMA (society)  303 Phasmid Study Group 303 phenotype 789 Philippines  267, 285, 290, 300, 316, 520, 521, 546, 692 phloem feeding  593, 596 phoresy  608, 687, 690 photograph sharing internet sites  840, 843 photography  447, 839 PhyloCode 7 phylogenetic analysis  325, 347, 389, 628, 691, 853, 854 Phylogenetic Systematics 6 phylogeny 246 phytoplasmas  509, 524, 539, 542 bois noir grapevine yellows 539 lethal yellowing  536 spike disease (of sandalwood)  542, 545 Piedmont physiographic province (United States of America)  376 planidium (larval type)  673, 677, 677, 680, 681, 685, 686, 687, 689, 690, 691, 692, 693 Planina Cave [Slovenia] 128, 143

985

986

Index

Pleistocene Epoch  251, 335, 724, 726, 770, 772 pollination  40, 42, 107, 109, 110, 327, 368, 376, 483, 705, 754, 763, 764, 809, 814, 820, 822 Portugal  173, 397 Postojnska cave [Slovenia]  123, 146 Primorsky Region [Russia]  343 pteridisperms  739 Puerto Rico  184, 376, 818 poultry pests  475 Pyrenees (Europe)  142

q Quaternary Period  15 Qur’an 879

r rabies 477 Rafinesque, Constantine Samuel 2 rearing insects  303 Réaumur, Réné  6 Red List 376 red ring disease (nematode) 774 refugium, glacial  30, 92 reindeer. See caribou/reindeer religion. See art, literature and religion reproductive behavior  171, 207, 231, 259, 261, 323, 339, 346, 361, 461, 490, 525, 530, 613, 646, 657, 659, 708, 715 resilin  319, 467, 710 Respiratory Quotient  38–39 Réunion [France, oversees territory]  270, 445 Riley, Charles V.  4, 6 Romania  143, 692 root feeding  812 Ross, Ronald  3 Russia  15, 29, 31, 40, 336, 343, 649, 652, 711

Russian Far East  343, 708, 714 Ryukyu Islands [Japan]  186

s Saint Helena [United Kingdom, overseas territory]  92, 186, 323, 417, 435, 447 saline habitats  33, 89, 173 Samoa  92, 146, 535, 816, 817 São Tomé e Príncipe  173, 180 Sarawak [Indonesia]  533 Say, Thomas  6 ScaleNet 597 Scandinavia 15 semelparity 323 Senegal 810 Serbia 142 Severnaya Zemlya [Russia]  35, 39, 40 sex determination  610, 611, 615 sexual dimorphism  177, 206, 252, 281, 291, 486, 609, 677, 685, 706 Seychelles  295, 297, 299, 376, 443, 544, 817 Siberechts, Jan  869 Siberia [Russia]  25, 31, 159, 338, 343 Sicily [Italy]  816 Signor–Lipps effect  729 silk 223, 224, 226, 228, 229, 231, 234, 235, 236, 237, 238, 239, 261, 490, 631, 632, 633, 636, 646, 651 Silurian Period  733, 757, 765 sky islands  89, 251 Slovenia  124, 126, 128, 131, 142, 143 snowshoe hare meningoencephalitis 42 snowshoe hare virus  42 social behavior  206, 371, 372, 490, 608, 609, 762 in insects  83, 229, 235, 239, 371, 372, 490, 608, 615 social parasite  852

soldier caste  372, 490, 608, 615 Solomon Islands  487, 546, 692 sound production. See acoustic communication South Africa  167, 175, 183, 186, 267, 269, 270, 345, 350, 511, 521, 533, 547, 693, 709, 734, 745, 754, 861 South America  185, 186, 233, 235, 248, 250, 254, 259, 262, 268, 270, 271, 290, 294, 367, 370, 371, 399, 435, 437, 438, 439, 442, 443, 444, 445, 447, 488, 519, 521, 537, 594, 642, 646, 650, 651, 653, 655, 678, 688, 692, 707, 708, 709, 747 Southampton Island [Nunavut, Canada] 29 South Carolina [United States of America]  210, 376 Southeast Asia  238, 251, 265, 290, 296, 299, 401, 434, 435, 494, 519, 649, 656, 679, 707 Spain  185, 397, 437, 688, 870 specialist  35, 87, 141, 302, 604, 605, 606, 659 speciation 606 species concepts  17 species diversity  18, 85, 725, 727, 728, 730, 804 Species File software and online catalogs 248, 249, 396 species richness  25, 27, 86, 142, 248, 725, 768, 809, 860 Spence, William  6 Spice Islands. See Moluccas spiroplasmas 509 Spitsbergen [Norway]  28 Sri Lanka  82, 437, 446, 542, 546, 811 subsocial behavior. See parental care

Index

Sudan  186, 324, 679 Sulawesi [Indonesia]  521, 546 Sunda islands  290 supernumary chromosomes  613 superparasitism  690, 692 Suriname 883 Svalbard [Norway]  15, 28, 29, 36, 39, 41, 42 swan (Cygnus) 474 Sweden 39 Switzerland 175 Systema Naturae  5, 155, 627

t Taimyr [Russia]  15, 39 Taiwan  439, 709, 714, 715 Tanzania  322, 345, 349, 397 Tasmania [Australia]  435, 443, 444, 544, 707, 712 taxonomic impediment  93 technology inspired by insects  228, 374 temperature tolerance  38, 338 Tertiary Period (= Paleogene + Neogene)  437, 444 Texas [United States of America]  134, 184, 237, 250, 739, 740, 747 Thailand  59, 186, 226, 300, 375, 692 The Torre‐Bueno Glossary of Entomology 791 thermal oases  28 3I Interactive Keys and Taxonomic Databases  510, 523 ThripsWiki 495 Tibet 199 tospovirus  483, 486, 494 toxic honey  548 toxic plants  251, 268 Triassic Period  517, 519, 591, 712, 726, 729, 734, 735, 749, 760, 771 Trinidad [Trinidad and Tobago]  437, 811

troglobites. See caves and cave dwelling arthropods tropical regions  818 trypanosomal diseases  774 Turkey  258, 397, 400 Turkmenistan  238, 376

u Uganda 322 United Kingdom  714, 745 United Nations Development Programme (UNDP)  60, 76 United States of America  105, 125, 126, 175, 199, 233, 266, 267, 290, 294, 302, 319, 371, 434, 437, 439, 440, 441, 446, 447, 529, 537, 538, 637, 647, 650, 688, 708, 714, 741, 745

v Vanuatu 816 vectors of animal disease, insects as  375, 473, 774, 818 vectors of plant disease, insects as  509, 519, 524, 526, 536, 539, 540, 774 Venezuela  370, 434, 439 venom 633 vicariance 815 Vietnam  59, 236, 283, 296, 512, 546, 647 viruses  509, 524, 540 barley yellow dwarf  769 geminivirus 591 hantavirus 477 Jamestown Canyon  42 myxoma 475 Northway 42 pseudo‐curly top of tomato 526 Snowshoe Hare  42 tospovirus  483, 486, 494 viviparous 315

Voss, Eduard  4

w Wallace, Alfred Russell  81, 691 Waller, Richard  6 Walsh, Benjamin  6 warning coloration. See aposematic coloration Washington State [United States of America]  341, 710 wax production  535, 538, 539, 540, 542, 543, 544, 545, 546 Web Ontology Language (OWL) 792 Western Australia  127 West Indies  85 Wildlife Conservation Society (WCS) 76 wingless insects  269, 435, 440, 441, 445 wing polymorphism  39, 206, 336, 370, 395, 486, 516, 535 Wisconsin [United States of America] 31 World Wide Fund for Nature (WWF)  62, 76 Wrangell Island [Russia]  39 Wyoming [United States of America]  749

x xylem feeding  596

y Yilgarn region [Australia]  128 Yukon Territory [Canada]  27, 29, 31

z Zackenberg [Greenland]  28, 42 Zimbabwe  437, 521, 532, 535, 544 zombification  730, 735, 767, 774

987

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Figure 4.1  Different kinds of insular systems. (a) Oceanic island. The island of Pico in the archipelago of the Azores. This archipelago is one of the best studied for arthropods, as evidenced through the work of Borges and colleagues (Borges 1992, Borges and Brown 1999, Cardoso et al. 2010, Triantis et al. 2010a, Triantis et al. 2010b, Cardoso et al. 2011, Gaspar et al. 2011, Meijer et al. 2011). (b) Cave. View towards entrance from within Algar do Carvão, an ancient lava tube on the Azorean island of Terceira, which harbors a number of endemics, including both spiders and insects (Reboleira et al. 2011). (c) Pleistocene fragment island. Agistri in the Sarconic Islands of Greece. Early on (23–12 million years ago), the Greek islands were all connected in a continuous land mass. Sea transgression (12–5 mya) formed a mid‐Aegean barrier, followed by fragmentation and widening of the Aegean, leading to the Pleistocene, which was characterized by eustatic sea‐level change (Triantis and Mylonas 2009). The long history of connection and isolation has shaped the diversity of arthropods known from the region today (Sfenthourakis and Legakis 2001). (d) Forest fragment. Shown is a “kipuka,” or island of forest surrounded by lava, on the island of Hawaii. Arthropods are often isolated in these fragments and show clear genetic differences among kipukas (Vandergast and Gillespie 2004, Vandergast et al. 2004).

Insect Biodiversity: Science and Society, Volume II, First Edition. Edited by Robert G. Foottit and Peter H. Adler. © 2018 John Wiley & Sons Ltd. Published 2018 by John Wiley & Sons Ltd.

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Figure 4.1 (Cont’d)  (e) Unique habitat islands. Mono Lake is an example of a unique habitat – a saline lake – that is isolated from similar such habitats. It harbors distinct assemblages of organisms, particularly notable being the brine shrimps and alkali flies (Herbst 1999). Other habitat types that hold unique assemblages of arthropods include sand dunes (Van Dam and Matzke 2016) and vernal or desert pools (Ward and Blaustein 1994). (f ) Recent fragment islands. Barro Colorado island, in Gatun Lake of Panama, was formed by flooding of the Chagres River in the creation of the Panama Canal (Leigh 2009). As a result, species numbers declined through relaxation of the supersaturated insular biota as it returns to equilibrium. A similar phenomenon has been documented for oaks, which act as islands for leaf‐mining insects (Opler 1974). (g) Sky islands. The American Madrean sky islands of southeastern Arizona and New Mexico have served to isolate many arthropods on the mountain summits. Particularly well known are the jumping spiders in the Habronattus pugillis complex (Masta 2000), scorpions (Hughes 2011), and beetles (Smith and Farrell 2005, Ober and Connolly 2015). All photographs by George K. Roderick, used with permission.

Figure 9.6  An adult female of Macrembia sp. from Zambia, showing bright colors thought to mimic staphylinid beetles also found in their habitat. Their especially robust silk ejectors are visible on the ventral surface of the front basal tarsus. The silk is unusually wispy for webspinners, which more typically spin silk into cloth‐like coverings. Photograph by author.

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Figure 10.3  Representative families of Ensifera. (a) Grylloidea: Gryllidae: Gryllinae. (b) Grylloidea: Gryllidae: Phalangopsinae. (c) Gryllotalpoidea: Gryllotalpidae. (d) Schizodactyloidea: Schizodactylidae: Comicinae. (e) Stenopelmatoidea: Stenopelmatidae. (f ) Stenopelmatoidea: Anostostomatidae. (g) Stenopelmatoidea: Gryllacrididae. (h) Rhaphidophoroidea: Rhaphidophoridae. (i) Hagloidea: Prophalangopsidae: Cyphoderrinae. (j) Tettigonioidea: Tettigoniidae: Conocephalinae. (k) Tettigonioidea: Tettigoniidae: Pseudophyllinae. (l) Tettigonioidea: Tettigoniidae: Pterochrozinae. (Photographs: Piotr Naskrecki).

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Figure 10.4  Representative families of Caelifera. (a) Tridactyloidea: Tridactylidae. (b) Tetrigoidea: Tetrigidae. (c) Proscopioidea: Proscopiidae. (d) Eumastacoidea: Episactidae. (e) Eumastacoidea: Thericleidae. (f ) Tanaoceroidea: Tanaoceridae. (g) Pneumoroidea: Pneumoridae. (h) Trigonopterygoidea: Trigonopterygidae. (i) Trigonopterygoidea: Xyronotidae. (j) Pyrgomorphoidea: Pyrgomorphidae. (k) Acridoidea: Pamphagidae. (l) Acridoidea: Acrididae. (Photographs: Piotr Naskrecki (a,b,e,g,j–l), Paul Lenhart (c), Robert A. Behrstock (d), Hartmut Wisch (f ), Chien C. Lee (h), and Paolo Fontana (i)).

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Figure 11.1  Similar, but not related: wingless female stick insects with long antennae. (a) Carausius morosus (Lonchodinae) from India (photograph by Christoph Seiler, Altlussheim, Germany). (b) Acanthoxyla inermis (Lanceocercata) from New Zealand (photograph by Mieke Duytschaever, Essen, Belgium). Both species resemble each other strikingly with regard to size, coloration, and lifestyle, both being obligatory parthenogens.

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Figure 11.2  Leaf imitators. (a) Phylliumbioculatum (Phylliinae), a true leaf insect from Borneo (photograph by Christoph Seiler, Altlussheim, Germany). (b) A pair of Malandaniapulchra (Lanceocercata) from Queensland, Australia(photograph by Kathy Hill and David Marshall, Auckland, New Zealand).

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Figure 11.3  Euphasmatodean eggs arranged according to egg‐laying technique (not to scale). The techniques include: (a–e) dropping/flicking away without capitulum; (f–j) dropping/flicking away with capitulum; (k–o) gluing; and (p–t) inserting into soil. (a) Bacillus rossius (Bacillinae), Europe. (b) Phyllium giganteum (Phylliinae), Malaysia. (c) Parapachymorpha spiniger (Clitumninae), Vietnam. (d) Anisomorpha paromalus (Pseudophasmatinae), Mexico. (e) Baculofractum insigne (Lonchodinae), Sumatra. (f) Didymuria violescens (Tropidoderinae), Australia. (g) Phobaeticus serratipes (Pharnaciini), Malaysia. (h) Staelonchodes harmani (Lonchodinae), Borneo. (i) Eurycnema osiris (Lanceocercata), Australia. (j) Alienobostra brocki (Diapheromerinae), Costa Rica. (k) Sipyloidea sipylus (Necrosciinae), Madagascar. (l) Gratidia sp. (Gratidiini), Africa. (m) Marmessoidea rosea (Necrosciinae), Malaysia. (n) Sceptrophasma hispidulum (Gratidiini), Andaman Islands. (o) Trachythorax maculicollis (Necrosciinae), Myanmar. (p) Rhamphophasma spinicorne (Clitumninae), Bangladesh. (q) Sungaya inexpectata (Heteropteryginae), Philippines. (r) Diesbachia tamyris (Necrosciinae), Sumatra. (s) Eurycantha calcarata (Lonchodinae), New Guinea. (t) Creoxylus spinosus (Pseudophasmatinae), Trinidad. After Seiler et al. 2000, photographs by Rainer Koch, Eppelheim, Germany.

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Figure 12.1  (a) Arixenia esau (Arixeniidae) from Borneo, with a unique life history and long, slender legs. (b) Female of Labidura riparia (Labiduridae) in defensive posture in Belgium. Labidura riparia prefers sandy underground habitats such as beaches and riverbanks. (c) A representative of Echinosoma sp. (Pygidicranidae) with uniquely strong, short bristles (i.e., modified setae). (d) Schizoproreus volcanus (Chelisochidae). The Chelisochidae form a small taxon with a preference for warm, humid tropics. The exception is Chelisoches morio, a tramp species of worldwide distribution. (e) Anisolabis maritima (Anisolabididae), a widespread, generalized earwig on which many behavioral and physiological studies have been conducted. Many of the Anisolabididae resemble this species, and identifications require examination of genitalia. (f ) Nymph of Diplatyidae from Brunei, with long, annulated cerci, a plesiomorphic character state in the Dermaptera. (g) Forficula senegalensis (Forficulidae) killed by an unknown fungus in Kenya. (h) Forficula auricularia feeding on grapes after they had been opened by wasps in Germany. (a,c–f ) Photographs by Petr Kocarek, University of Ostrava. (b,g,h) Photographs by Fabian Haas.

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Figure 13.5  (a–d) Grylloblattodea. (a) Female of Grylloblatta campodeiformis occidentalis. (b) Female of Grylloblattella sp. (c) Male of Galloisiana sp. (d) Female of Galloisiana yuasai. (e–h) Mantophasmatodea. (e) Female of Karoophasma biedouwense (Austrophasmatidae). (f ) Male of K. biedouwense. (g) Mating pair of K. biedouwense. (h) Female of Viridiphasma clanwilliamense. Photographs by authors.

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Figure 14.1  Cockroaches. (a) Corydiidae habitus. (b) Ectobiidae (Anallacta) habitus. (c) Blaberidae (Zuluia female) habitus. (d) Blaberidae (Paranauphoeta formosana male) habitus. (e) Blaberidae (Thorax porcellana male) habitus. (f ) Blaberidae (Aptera fusca), female with young, showing brooding behavior. (g) Blattidae (Pseudoderopeltis cf. albilatera), female with ootheca. (h) Cryptocercidae (Cryptocercus), adult and nymph. Images by Z. Varadinova (a–e,g,h) and M. Picker and C. Griffiths (f ).

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Figure 15.1  Live habitus images showing the morphological diversity in the Mantodea. (a) Male of Mantoida sp. from Nicaragua. (b) Female of Acontista sp. from Bolivia. (c) Male of Liturgusa cursor from Nicaragua. (d) Male of Enicophlebia hilara from Madagascar. (e) Male of Compsothespis sp. from Rwanda. (f ) Female of Oxyelaea elegans from Rwanda. (g) Female of Popa spurca from Madagascar. (h) Male of Hymenopus coronatus from Sarawak, Malaysia. (i) Male of Tisma grandidieri from Madagascar. Photographs by authors.

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Figure 16.11  Habitus images of psocopterans. (a) Lithoseopsis hellmani (Mockford) (Amphientomidae). (b) Teliapsocus conterminus (Walsh) (Dasydemellidae). (c) Graphopsocus cruciatus (L.) (Stenopsocidae). (d) Peripsocus subfasciatus (Rambur) (Peripsocidae), female. (e) Ectopsocus meridionalis Ribaga (Ectopsocidae), female. (f ) Lachesilla contraforcepeta Chapman (Lachesillidae). (g) Hyalopsocus sp. (Psocidae), male (left) and female. Images by Diane Young.

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Figure 19.3  Cercopoidea and Myerslopiidae. (a) Aphrophoridae, nymph of Neophilaenus lineatus (Austria, © 2009 Gernot Kunz, used by permission). (b) Aphrophoridae, Philagra cf. parva (Australia). (c) Cercopidae, Cercopis vulnerata (Austria, © 2008 Gernot Kunz, used by permission). (d) Epipygidae, Eicissus sp. (Costa Rica, © 2010 Gernot Kunz, used by permission). (e) Clastopteridae, Clastoptera sp. (Nicaragua, Stephen Cresswell, used by permission). (f ) Machaerotidae, Machaerota coomani (holotype, Vietnam; © 2010 Gernot Kunz, used by permission). (g) Myerslopiidae, Myerslopia rakiuraensis (Stewart Is., New Zealand, Larivière and Larochelle 2013–2015, used by permission).

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Figure 19.4  Cicadellidae. (a) Iassinae, Rugosana querci. (b) Evacanthinae, Boundarus sp. (c) Typhlocybinae, Hymetta balteata. (d) Typhlocybinae, Erythroneura palimpsesta. (e) Ledrinae, Hespenedra sp. (f ) Deltocephalinae, Flexamia pict). (g) Idiocerinae, Idiocerus sp. (h) Cicadellinae, Pawiloma sp. (All photos by ©2015 Christopher H. Dietrich, Illinois Natural History Survey, used by permission).

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Figure 19.5  Membracoidea: (a,b) Aetalionidae; (c) Melizoderidae; and (d–f ) Membracidae. (a) Aetalioninae, Aetalion reticulatum, female guarding eggs mass (©2012 Gernot Kunz, used by permission). (b) Biturritiinae, Tropidaspis sp., female guarding egg mass and tended by ants (Ecuador, ©2010 Kelly Swing, used by permission). (c) Melizoderes sp. (Chile, ©2014 Christopher H. Dietrich, used by permission). (d) Smiliinae, Polyglyptini, Heranice miltoglypta, female guarding eggs (Colombia, ©2011 David Guzman, used by permission). (e) Heteronotinae, Heteronotus sp. (Ecuador, ©2006 Kelly Swing, used by permission). (f ) Endoiastinae, Endoiastus sp. with tending ant (©2012 Gernot Kunz, used by permission).

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Figure 19.6  Membracoidea: Membracidae. (a) Centronodinae, Paracentronodus sp. (Ecuador, ©2010 Kelly Swing, used by permission). (b) Stegaspidinae, Stegaspidini, Umbelligerus woldai (©2011 Gernot Kunz, used by permission). (c) Nicomiinae, Holdgateilla chepuensis (Chile, ©2014 Christopher H. Dietrich, used by permission). (d) Centrotinae, Hypsaucheniini, Jingkara hyalipunctata, female guarding eggs (China, ©2003 Robert L. Snyder, used by permission). (e) Darninae, Hemikypthini, Hemikyptha sp., among the largest of treehoppers (Ecuador, ©2009 Kelly Swing, used by permission). (f ) Membracinae, Hoplophorionini, Metcalfiella vicina, female with nymphs (Colombia, ©2012, David Guzman, used by permission).

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Figure 20.1  Examples of each sternorrhynchan superfamily. (a) Wingless adult of a Cinara species (Aphidoidea: Aphididae). (b) Winged and wingless forms of Aphis sambuci (Aphidoidea: Aphididae). (c) Adult females of Icerya purchasi (Coccoidea: Monophlebidae) with ovisacs. (d) Adult female of Melanaspis obscura (Coccoidea: Diaspididae) with test removed. (e) Adult of Bemisia tabaci (Aleyrodoidea: Aleyrodidae). (f ) Adult of Chamaepsylla hartigii (Psylloidea: Psyllidae). The psyllid photograph was taken by Gabrijel Seljak. Other photographs by author.

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Figure 21.1  Representative adults and larvae of the orders Megaloptera and Raphidioptera. (a) Corydalus sp., adult, Brazil (Megaloptera: Corydalidae). (b) Sialis lutaria, adult, Poland (Megaloptera: Sialidae). (c) Sialis lutaria, larva, Czech Republic (Megaloptera: Sialidae). (d) Ascalaphidae sp., larva, Nicaragua (Neuroptera: Ascalaphidae). (e) Suphalomitus sp., adult, Australia (Neuroptera: Ascalaphidae). (f ) Spermophorella sp., adult, Australia (Neuroptera: Berothidae). (g) Chrysopidae sp., larvae, Colombia (Neuroptera: Chrysopidae). (h) Hypochrysa elegans, adult, Belgium (Neuroptera: Chrysopidae). Photo credits: Arthur Anker (a), Łukasz Prajzne (b), Jan Hamrsky (c), Marshal Hedin (d), Craig Nieminski (e), Shaun Winterton (f ), Robert Oelman (g), Gilles San Martin (h).

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Figure 21.3  Representative adults and larvae of the orders Neuroptera and Raphidioptera. (a) Nymphes myrmeleonoides, eggs and first instar larvae, Australia (Neuroptera: Nymphidae). (b) Nymphes myrmeleonoides, adult, Australia (Neuroptera: Nymphidae). (c) Porismus strigatus, adult, Australia (Neuroptera: Osmylidae). (d) Psychopsis insolens, adult, Australia (Neuroptera: Psychopsidae). (e) Sisyra fuscata, larva, Czech Republic (Neuroptera: Sisyridae). (f ). Sisyra terminalis, adult, Belgium (Neuroptera: Sisyridae). (g). Parainocellia bicolor, larva, Italy (Raphidioptera: Inocelliidae). (h) Agulla sp., adult, United States (Raphidioptera: Raphidiidae). Photo credits: Jim McLean (a), Michael Jefferies (b), Shaun Winterton (c,d,h), Jan Hamrsky (e), Gilles San Martin (f ), Marcello Romano (g).

Figure 23.2  A mating pair of Harpobittacus similis. The female is feeding on the arthropod nuptial gift given to her by the mating male. (Image by W. Bicha.)

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