Handbook of vegetable pests [Second edition] 9780128144886, 0128144882, 9780128144893, 0128144890

Table of contents :
Front Cover......Page 1
Handbook of Vegetable Pests......Page 4
Copyright......Page 5
Dedication......Page 6
Contents......Page 8
Preface......Page 14
Acknowledgments......Page 16
What Are the Major Vegetable Crops?......Page 18
Cabbage and Related Crops (Family Brassicae, Formerly Cruciferae) (Broccoli, Brussels Sprouts, Cabbage, Cauliflower, Chines .........Page 20
Lettuce and Related Crops (Family Compositae) (Artichoke, Celtuce, Chicory, Endive, Escarole, Lettuce, Radicchio)......Page 21
Sweet Corn (Family Poaceae, Formerly Graminae)......Page 22
Frequency of Pests Among the Major Insect Orders......Page 23
Family Chrysomelidae—Leaf Beetles......Page 24
Family Meloidae—Blister Beetles......Page 25
Syrphidae—Flower flies......Page 26
Family Pentatomidae—Stink bugs......Page 27
Pseudococcidae—Mealybugs......Page 28
Family Erebidae—Tiger moths, Woolly Caterpillars, and others......Page 29
Family Sphingidae—Hawk moths and hornworms......Page 30
Order Thysanoptera—Thrips......Page 31
What Is a pest?......Page 32
How damaging Are pests?......Page 33
The types of insect injury......Page 34
Pest management philosophy......Page 35
Physical manipulations......Page 36
Approaches to identification......Page 37
Approaches to Identification......Page 40
Other pests occasionally found on asparagus......Page 60
Other Pests Occasionally Found on Bean and Related Crops......Page 61
Insects Producing Small Mines in Leaves......Page 62
Pests feeding externally on leaves or stems......Page 63
Other pests occasionally found on lettuce and related crops......Page 64
Other pests occasionally found on onion and related plants......Page 65
Other pests occasionally found on squash and related crops......Page 66
Other pests occasionally found on corn......Page 67
Pests producing mines in leaves......Page 68
Other pests occasionally found on tomato and related plants......Page 69
Part I Class Insecta—Insects......Page 70
Natural History......Page 72
Damage......Page 73
Natural History......Page 74
Damage......Page 75
Natural History......Page 76
Management......Page 79
Natural History......Page 80
Natural History......Page 81
Natural History......Page 82
Management......Page 83
Natural History......Page 84
Management......Page 85
Natural History......Page 86
Damage......Page 87
Natural History......Page 88
Natural History......Page 89
Natural History......Page 90
Natural History......Page 91
Damage......Page 92
Natural History......Page 93
Damage......Page 94
Natural History......Page 95
Three-Spotted Flea Beetle......Page 96
Management......Page 97
Natural History......Page 98
Natural History......Page 99
Natural History......Page 100
Damage......Page 101
Natural History......Page 102
Management......Page 103
Natural History......Page 104
Natural History......Page 105
Natural History......Page 106
Natural History......Page 107
Management......Page 108
Natural History......Page 109
Natural History......Page 111
Management......Page 113
Natural History......Page 114
Management......Page 115
Natural History......Page 116
Natural History......Page 118
Management......Page 119
Natural History......Page 120
Damage......Page 121
Natural History......Page 122
Management......Page 125
Natural History......Page 126
Natural History......Page 128
Damage......Page 129
Management......Page 130
Natural History......Page 131
Natural History......Page 132
Natural History......Page 133
Damage......Page 135
Management......Page 136
Natural History......Page 137
Management......Page 138
Natural History......Page 139
Natural History......Page 140
Management......Page 142
Natural History......Page 144
Management......Page 145
Natural History......Page 146
Damage......Page 148
Natural History......Page 149
Damage......Page 150
Natural History......Page 151
Natural History......Page 152
Management......Page 153
Natural History......Page 154
Management......Page 156
Natural History......Page 157
Natural History......Page 158
Natural History......Page 160
Management......Page 162
Natural History......Page 163
Natural History......Page 165
Damage......Page 166
Natural History......Page 167
Natural History......Page 168
Damage......Page 169
Management......Page 170
Natural History......Page 171
Management......Page 172
Natural History......Page 173
Natural History......Page 175
Management......Page 176
Natural History......Page 177
Management......Page 178
Natural History......Page 179
Damage......Page 180
Management......Page 181
Natural History......Page 182
Damage......Page 183
Natural History......Page 184
Management......Page 186
Natural History......Page 187
Damage......Page 188
Natural History......Page 189
Natural History......Page 191
Management......Page 193
Damage......Page 194
Natural History......Page 195
Natural History......Page 196
Management......Page 197
Natural History......Page 198
Management......Page 199
Natural History......Page 200
Natural History......Page 201
Natural History......Page 203
Natural History......Page 204
Management......Page 205
Natural History......Page 206
Management......Page 207
Natural History......Page 208
Management......Page 210
Natural History......Page 211
Management......Page 212
Natural History......Page 213
Natural History......Page 214
Natural History......Page 215
Damage......Page 216
Natural History......Page 217
Management......Page 219
Natural History......Page 220
Management......Page 221
Natural History......Page 222
Natural History......Page 224
Management......Page 226
Natural History......Page 228
Damage......Page 229
Natural History......Page 230
Damage......Page 231
Natural History......Page 232
Natural History......Page 233
Natural History......Page 234
Damage......Page 235
Natural History......Page 236
Natural History......Page 238
Natural History......Page 239
Damage......Page 240
Natural History......Page 241
Management......Page 243
Natural History......Page 244
Damage......Page 246
Management......Page 247
Natural History......Page 248
Natural History......Page 249
Damage......Page 250
Management......Page 251
Damage......Page 252
Natural History......Page 253
Damage......Page 254
Natural History......Page 255
Natural History......Page 257
Management......Page 259
Natural History......Page 260
Natural History......Page 261
Management......Page 263
Natural History......Page 264
Management......Page 265
Natural History......Page 266
Management......Page 267
Natural History......Page 268
Management......Page 269
Natural History......Page 270
Management......Page 271
Natural History......Page 272
Management......Page 273
Natural History......Page 274
Management......Page 275
Natural History......Page 276
Management......Page 278
Natural History......Page 279
Damage......Page 281
Management......Page 282
Natural History......Page 283
Natural History......Page 284
Natural History......Page 285
Management......Page 287
Natural History......Page 288
Natural History......Page 290
Damage......Page 291
Natural History......Page 292
Management......Page 294
Natural History......Page 295
Natural History......Page 296
Natural History......Page 298
Management......Page 299
Natural History......Page 300
Management......Page 301
Natural History......Page 302
Natural History......Page 303
Management......Page 306
Natural History......Page 307
Management......Page 308
Natural History......Page 309
Management......Page 310
Natural History......Page 311
Damage......Page 312
Management......Page 313
Natural History......Page 314
Damage......Page 316
Natural History......Page 317
Damage......Page 319
Natural History......Page 320
Natural History......Page 321
Natural History......Page 323
Natural History......Page 325
Management......Page 326
Natural History......Page 327
Natural History......Page 329
Management......Page 331
Natural History......Page 332
Management......Page 334
Natural History......Page 335
Management......Page 336
Natural History......Page 337
Damage......Page 338
Natural History......Page 339
Natural History......Page 340
Management......Page 342
Natural History......Page 343
Management......Page 345
Natural History......Page 346
Natural History......Page 347
Management......Page 348
Natural History......Page 349
Natural History......Page 350
Natural History......Page 352
Natural History......Page 353
Damage......Page 354
Natural History......Page 355
Natural History......Page 356
Management......Page 358
Natural History......Page 360
Natural History......Page 362
Management......Page 364
Natural History......Page 365
Management......Page 366
Natural History......Page 367
Natural History......Page 368
Damage......Page 370
Natural History......Page 371
Management......Page 372
Natural History......Page 373
Natural History......Page 374
Management......Page 376
Natural History......Page 377
Natural History......Page 378
Management......Page 379
Natural History......Page 380
Management......Page 381
Natural History......Page 382
Damage......Page 384
Natural History......Page 385
Natural History......Page 386
Natural History......Page 388
Natural History......Page 390
Natural History......Page 392
Management......Page 393
Natural History......Page 394
Natural History......Page 396
Natural History......Page 397
Damage......Page 398
Management......Page 399
Natural history......Page 400
Natural history......Page 401
Damage......Page 403
Management......Page 404
Natural History......Page 406
Management......Page 407
Natural History......Page 408
Natural History......Page 409
Natural History......Page 410
Natural History......Page 412
Management......Page 414
Natural History......Page 415
Damage......Page 416
Natural History......Page 417
Management......Page 418
Natural History......Page 419
Natural History......Page 420
Management......Page 423
Natural History......Page 424
Natural History......Page 426
Natural History......Page 427
Damage......Page 429
Natural History......Page 430
Management......Page 432
Natural History......Page 433
Management......Page 434
Natural History......Page 435
Natural History......Page 436
Damage......Page 437
Natural History......Page 438
Natural History......Page 439
Management......Page 441
Natural History......Page 442
Natural History......Page 444
Natural History......Page 445
Natural History......Page 446
Natural History......Page 448
Management......Page 449
Natural History......Page 450
Natural History......Page 451
Natural History......Page 453
Management......Page 454
Natural History......Page 455
Natural History......Page 456
Management......Page 458
Natural History......Page 459
Natural History......Page 460
Natural History......Page 462
Natural History......Page 464
Damage......Page 465
Natural History......Page 466
Natural History......Page 467
Natural History......Page 469
Natural History......Page 472
Natural History......Page 473
Management......Page 475
Natural History......Page 476
Natural History......Page 478
Natural History......Page 479
Damage......Page 481
Life history......Page 482
Natural History......Page 484
Management......Page 487
Natural History......Page 488
Damage......Page 489
Natural History......Page 490
Natural History......Page 491
Management......Page 494
Natural History......Page 496
Natural History......Page 497
Natural History......Page 499
Management......Page 502
Natural History......Page 503
Natural History......Page 504
Natural History......Page 506
Natural History......Page 508
Management......Page 510
Natural History......Page 511
Natural History......Page 513
Natural History......Page 515
Natural History......Page 517
Natural History......Page 519
Management......Page 520
Natural History......Page 521
Natural History......Page 523
Natural History......Page 525
Damage......Page 527
Natural History......Page 528
Damage......Page 529
Natural History......Page 530
Management......Page 532
Natural History......Page 533
Management......Page 534
Natural History......Page 535
Management......Page 536
Natural History......Page 537
Natural History......Page 539
Natural History......Page 540
Natural History......Page 542
Natural History......Page 544
Management......Page 546
Natural History......Page 547
Natural History......Page 548
Natural History......Page 549
Natural History......Page 551
Management......Page 553
Natural History......Page 555
Natural History......Page 556
Management......Page 558
Natural History......Page 559
Natural History......Page 560
Management......Page 562
Natural History......Page 563
Natural History......Page 564
Damage......Page 566
Natural History......Page 567
Natural History......Page 568
Damage......Page 569
Management......Page 570
Natural history......Page 572
Damage......Page 573
Natural history......Page 574
Natural history......Page 576
Management......Page 578
Natural history......Page 579
Management......Page 582
Natural history......Page 584
Natural history......Page 586
Natural history......Page 588
Natural history......Page 590
Natural history......Page 593
Management......Page 596
Natural History......Page 598
Damage......Page 599
Natural History......Page 600
Damage......Page 601
Natural History......Page 602
Natural History......Page 603
Damage......Page 604
Damage......Page 605
Natural History......Page 606
Management......Page 607
Natural History......Page 608
Management......Page 610
Natural History......Page 612
Management......Page 613
Natural History......Page 614
Management......Page 616
Natural History......Page 618
Natural History......Page 619
Natural History......Page 621
Natural History......Page 622
Management......Page 624
Natural History......Page 625
Natural History......Page 626
Damage......Page 627
Natural History......Page 628
Damage......Page 629
Natural History......Page 630
Natural History......Page 632
Management......Page 635
Natural History......Page 636
Natural History......Page 639
Damage......Page 640
Management......Page 641
Key to major orders of insects affecting vegetable crops......Page 644
Key to common stink bugs affecting vegetables......Page 649
Key to common armyworms and cutworms......Page 652
Key to common “cabbageworms”......Page 656
Key to common loopers affecting vegetables......Page 657
Key to common stalk borers affecting corn......Page 658
Guide to common adult flea beetles......Page 659
Key to genera of wireworms commonly affecting vegetables......Page 660
Key to common thrips affecting vegetables......Page 662
Key to common mites affecting vegetables......Page 665
Key to adult slugs commonly affecting vegetables......Page 666
Key to adult snails commonly affecting vegetables......Page 667
Common name, scientific name, and plant family......Page 668
Scientific name, common name, and plant family......Page 671
Plant family, common name, and scientific name......Page 673
Journal Abbreviations and Journal Titles......Page 676
Glossary......Page 682
References......Page 688
Index......Page 806
Back Cover......Page 818

Citation preview

Handbook of Vegetable Pests

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Handbook of Vegetable Pests Second Edition

John L. Capinera

Emeritus Professor Department of Entomology and Nematology Institute of Food and Agriculture Sciences University of Florida United States

Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1650, San Diego, CA 92101, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom © 2020 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN 978-0-12-814488-6 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Charlotte Cockle Acquisitions Editor: Nancy Maragioglio Editorial Project Manager: Billie Jean Fernandez Production Project Manager: Prem Kumar Kaliamoorthi Cover Designer: Christian Bilbow Typeset by SPi Global, India

This book is dedicated to my wife, Marsha, who assisted me in many ways. Her continuing support will be long remembered.

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Contents Preface Acknowledgments

xiii xv

1. Introduction North American Vegetable Crops 1 What Are the Major Vegetable Crops? 1 Characteristics of the Major Vegetable Crops 3 Insects and Insect Relatives 6 Frequency of Pests Among the Major Insect Orders 6 Noninsect Vegetable Pests 7 Order Blattodea—Cockroaches and Termites 7 Order Coleoptera—Beetles, White Grubs, and Wireworms 7 Order Diptera—Flies and Maggots 9 Order Hemiptera—Bugs 10 Order Hymenoptera—Ants, Bees, Sawflies, and Wasps 12 Order Lepidoptera—Moths, Butterflies, and Caterpillars 12 Order Orthoptera—Grasshoppers, Katydids, and Crickets 14 Order Thysanoptera—Thrips 14 Noninsect Invertebrate Pests 15 Pest Management Philosophy and Practices 15 What Is a Pest? 15 How Damaging Are Pests? 16 The Types of Insect Injury 17 Pest Management Philosophy 18 Pest Management Practices 19 Species Identification 20

2. Pest Identification General Considerations Why Identification Is so Important? Approaches to Identification

23 23 23

3. Guides to Pest Identification, Arranged by Plant Taxon Guide to Common Pests Affecting Asparagus Pests Boring into Roots

43 43

Pests Producing Small Mines in Stems Pests Feeding Externally on Leaves (Cladophylls) or Stems Pests Boring into Berries Guide to Common Pests Affecting Bean and Related Crops Pests Boring into Roots or Stems Pests Producing Small Mines in Leaves Pests Feeding Externally on Leaves or Stems Pests Feeding on Flowers, Seeds, or Seedpods Guide to Common Pests Affecting Beet and Related Crops Pests Feeding on Roots Pests Producing Mines in Leaves Pests Feeding Externally on Leaves or Stems Guide to Common Pests Affecting Cabbage and Related Crops Pests Boring into Roots or Stems Insects Producing Small Mines in Leaves Pests Feeding Externally on Leaves or Stems Pests Feeding on Flowers, Seeds, or Seedpods Guide to Common Pests Affecting Carrot and Related Plants Pests Boring into Stems or Feeding on Roots Pests Producing Small Mines in Leaves Pests Feeding Externally on Leaves or Stems Pests Feeding on Flowers or Seeds Guide to Common Pests Affecting Lettuce and Related Crops Pests Feeding on Roots Pests Producing Mines in Leaves Pests Feeding Externally on Leaves or Stems Guide to Identification of Common Insect Pests Affecting Okra Pests Feeding on Roots or Lower Stem Pests Producing Mines in Leaves Pests Boring into Stems Pests Feeding Externally on Leaves Pests Feeding on Blossoms or Fruits Guide to Common Pests Affecting Onion and Related Plants Pests Boring into Bulbs or Roots Pests Producing Small Mines in Leaves Pests Feeding Externally on Leaves or Stems

43 43 43 44 44 44 44 44 45 45 45 45 45 45 45 46 46 46 46 46 46 47 47 47 47 47 48 48 48 48 48 48 48 48 48 48 vii

viii  Contents

Guide to Common Pests Affecting Rhubarb Pests Feeding on Roots Pests Living within Stems and Stalks Pests Feeding Externally on Leaves or Stems Guide to Common Pests Affecting Squash and Related Crops Pests Boring into Roots, Stems, Blossoms, or Fruit Pests Producing Small Mines in Leaves Pests Feeding Externally on Leaves, Stems, Blossoms, or Fruit Guide to Common Pests Affecting Sweet Corn Pests Feeding on Roots Pests Feeding on Seed or Seedling Pests Boring into Stems or Taproot Pests Producing Small Mines in Leaves Pests Feeding Externally on Leaves, Tassel, or Stalk Pests Feeding on Ears or Silk Guide to Common Pests Affecting Sweet Potato Pests Boring into Vines or Roots Pests Producing Small Mines in Leaves Pests Feeding Externally on Leaves or Stems Guide to Common Pests Affecting Tomato and Related Plants Pests Feeding on Roots, Tubers, or Lower Stem Pests Producing Mines in Leaves Pests Boring into Stems Pests Feeding Externally on Leaves or Upper Stems Pests Feeding on Blossoms or Fruits

49 49 49 49 49 49 49 49 50 50 50 50 50 50 50 51 51 51 51 51 51 51 52 52 52

Part I Class Insecta—Insects 4. Order Blattodea––Cockroaches and Termites Family Ectobiidae Asian Cockroach Family Blaberidae Surinam Cockroach Indian Cockroach Family Rhinotermitidae Subterranean Termites

55 55 57 57 57 59 59

5. Order Coleoptera—Beetles, White Grubs, and Wireworms Family Carabidae—Ground Beetles

63

Seedcorn Beetle Slender Seedcorn Beetle Family Chrysomelidae, Subfamily Alticinae—Flea Beetles Cabbage Flea Beetle Corn Flea Beetle Toothed Flea Beetle Crucifer Flea Beetle Desert Corn Flea Beetle Eggplant Flea Beetle Hop Flea Beetle Horseradish Flea Beetle Palestriped Flea Beetle Elongate Flea Beetle Potato Flea Beetle Western Potato Flea Beetle Redheaded Flea Beetle Smartweed Flea Beetle Spinach Flea Beetle Three-Spotted Flea Beetle Yellow-Necked Flea Beetle Striped Flea Beetle Western Striped Flea Beetle Sweetpotato Flea Beetle Tobacco Flea Beetle Southern Tobacco Flea Beetle Tuber Flea Beetle Western Black Flea Beetle Zimmermann’s Flea Beetle Family Chrysomelidae, Subfamily Bruchinae—Pea and Bean Seed Beetles (Pea and Bean Weevils) Bean Seed Beetle (Bean Weevil) Broadbean Seed Beetle (Broadbean Weevil) Cowpea Seed Beetle (Cowpea Weevil) Southern Cowpea Seed Beetle (Southern Cowpea Weevil) Pea Seed Beetle (Pea Weevil) Family Chrysomelidae, Subfamily Cassidinae—Tortoise Beetles Argus Tortoise Beetle Blacklegged Tortoise Beetle Golden Tortoise Beetle Mottled Tortoise Beetle Striped Tortoise Beetle Eggplant Tortoise Beetle Family Chrysomelidae, Several Subfamilies—Leaf Beetles Asparagus Beetle Banded Cucumber Beetle Bean Leaf Beetle Colorado Potato Beetle Grape Colaspis Northern Corn Rootworm

63 63 64 64 65 65 67 69 71 72 73 74 74 76 76 78 78 79 79 80 81 81 82 83 83 85 87 88

89 89 90 92 92 94 97 97 97 97 97 97 99 99 99 101 103 105 109 111

Contents  ix

Red Turnip Beetle Spotted Asparagus Beetle Spotted Cucumber Beetle Striped Cucumber Beetle Western Striped Cucumber Beetle Sweetpotato Leaf Beetle Western Corn Rootworm Yellowmargined Leaf Beetle Family Coccinellidae—Lady Beetles Mexican Bean Beetle Squash Beetle Families Curculionidae and Brentidae—Weevils and Primitive Weevils Cabbage Curculio Cabbage Seedpod Weevil Carrot Weevil Texas Carrot Weevil Cowpea Curculio Maize Billbug Southern Corn Billbug Pea Leaf Weevil Pepper Weevil, Anthonomus eugenii Cano, and Cuban Pepper Weevil Potato Stalk Borer Rhubarb Curculio Sweetpotato Weevil Vegetable Weevil West Indian Sugarcane Rootstalk Borer Weevil West Indian Sweetpotato Weevil Whitefringed Beetle Family Elateridae—Click Beetles and Wireworms Corn Wireworm Oregon Wireworm Dryland Wireworm Great Basin Wireworm Prairie Grain Wireworm Puget Sound Wireworm Southern Potato Wireworm Tobacco Wireworm Gulf Wireworm Sugarbeet Wireworm Eastern Field Wireworm Pacific Coast Wireworm Wheat Wireworm Family Meloidae—Blister Beetles Black Blister Beetle Immaculate Blister Beetle Spotted Blister Beetle Striped Blister Beetle Family Nitidulidae—Sap Beetles Dusky Sap Beetle Four-Spotted Sap Beetle

114 115 116 120 120 122 123 127 129 129 132

134 134 135 137 137 140 141 141 143 146 148 150 151 154 156 158 160 162 162 162 165 165 165 165 167 167 167 170 170 170 172 174 174 177 178 179 181 181 183

Family Scarabaeidae—Scarab Beetles and White Grubs Asiatic Garden Beetle Carrot Beetle Chinese Rose Beetle Green June Beetle Japanese Beetle Oriental Beetle Rose Chafer Western Rose Chafer Spring Rose Beetle Sugarcane Beetle White Grubs Family Tenebrionidae—Darking Beetles and False Wireworms False Wireworms

184 184 186 187 189 191 194 196 196 197 198 200 203 203

6. Order Dermaptera—Earwigs European Earwig Ringlegged Earwig African Earwig

205 207 207

7. Order Diptera—Flies and Maggots Family Agromyzidae—Leafminer Flies American Serpentine Leafminer Asparagus Miner Cabbage Leafminer Corn Blotch Leafminer Pea Leafminer Vegetable Leafminer Family Anthomyiidae—Root and Seed Maggots, Leafminer Flies Bean Seed Maggot Beet Leafminer Spinach Leafminer Cabbage Maggot Onion Maggot Radish Root Maggot Seedcorn Maggot Turnip Root Maggot Family Cecidomyiidae—Gall Midges Swede Midge Family Drosophilidae—Pomace Flies Small Fruit Flies Family Psilidae—Rust Flies Carrot Rust Fly Family Syrphidae—Flower and Bulb Flies Onion Bulb Fly Lesser Bulb Fly Family Tephritidae—Fruit Flies Mediterranean Fruit Fly Melon Fly

211 211 213 215 216 217 219 221 221 222 222 224 227 231 232 235 236 236 238 238 240 240 243 243 243 244 244 247

x  Contents

Oriental Fruit Fly Parsnip Leafminer Pepper Maggot Family Tipulidae—Crane Flies European Marsh Crane Fly Cabbage Crane Fly Family Ulidiidae—Picturewing Flies Cornsilk Flies Sugarbeet Root Maggot

249 251 251 253 253 253 255 255 257

8. Order Hemiptera—Bugs Family Aleyrodidae—Whiteflies Greenhouse Whitefly Sweetpotato Whitefly Family Aphididae—Aphids Artichoke Aphid Asparagus Aphid Bean Aphid Bean Root Aphid Bird Cherry-Oat Aphid Buckthorn Aphid Cabbage Aphid Carrot Root Aphids Coriander Aphid Honeysuckle Aphid Corn Leaf Aphid Corn Root Aphid Cowpea Aphid Koch Foxglove Aphid Green Peach Aphid Lettuce Aphid Lettuce Root Aphid Melon Aphid Pea Aphid Blue Alfalfa Aphid Potato Aphid Rice Root Aphid Sugarbeet Root Aphids Turnip Aphid Willow-Carrot Aphid Family Blissidae—Blissid Bugs Chinch Bug Family Cicadellidae—Leafhoppers Aster Leafhopper Beet Leafhopper Corn Leafhopper Potato Leafhopper Western Potato Leafhopper Family Coreidae––Leaffooted and Squash Bugs Leaffooted Bugs Squash Bugs Family Cydnidae—Burrower Bugs Burrowing Bug

259 259 262 266 266 267 268 271 271 273 275 278 278 278 279 281 283 285 286 290 292 294 297 297 300 303 304 306 308 310 310 312 312 315 318 320 322 323 323 326 329 329

Family Delphacidae—Planthoppers Corn Delphacid Family Lygaeidae—Seed Bugs False Chinch Bug Family Miridae—Plant Bugs Alfalfa Plant Bug Carrot Plant Bug Garden Fleahopper Rapid Plant Bug Tarnished Plant Bug Western Tarnished Plant Bug Pale Legume Bug Tomato Plant Bugs Family Pentatomidae—Stink Bugs Brown Marmorated Stink Bug Brown Stink Bug Consperse Stink Bug Green Stink Bug Harlequin Bug Onespotted Stink Bug Painted Bug Redbanded Stink Bug Redshouldered Stink Bug Say Stink Bug Uhler Stink Bug Southern Green Stink Bug Tomato Stink Bug Family Pseudococcidae—Mealybugs Cotton Mealybug Madeira Mealybug Pink Hibiscus Mealybug Pyrrhocoridae—Cotton Stainers Cotton Stainer Family Psyllidae—Psyllids Potato Psyllid Family Thyrecoridae—Ebony Bugs Little Ebony Bug Family Tingidae—Lace Bugs Eggplant Lace Bug

330 330 332 332 333 333 335 336 338 339 339 339 343 345 345 348 350 351 354 356 357 360 361 363 363 365 368 369 369 371 373 375 375 377 377 379 379 380 380

9. Order Hymenoptera—Ants and Sawflies Family Argidae—Sawflies Sweetpotato Sawfly Family Formicidae—Ants Red Imported Fire Ant

383 383 384 384

10. Order Lepidoptera—Caterpillars, Moths, and Butterflies Family Acrolepiidae—False Diamondback Moths Leek Moth

389 389

Contents  xi

Family Bedellidae—Bedellid Moths Morningglory Leafminer Sweetpotato Leafminer Family Depressariidae—Depressariid Moths Parsnip Webworm Family Crambidae—Borers, Budworms, Leaftiers, Webworms, and Snout Moths Alfalfa Webworm Beet Webworm Cabbage Budworm Cabbage Webworm Oriental Cabbage Webworm Celery Leaftier False Celery Leaftier Cross-Striped Cabbage Worm European Corn Borer European Pepper Moth Garden Webworm Hawaiian Beet Webworm Spotted Beet Webworm Southern Beet Webworm Melonworm Cucumber Moth Pickleworm Purplebacked Cabbage Worm Sod and Root Webworms Southern Cornstalk Borer Southwestern Corn Borer Sugarcane Borer Sweetpotato Leaf Folder Sweetpotato Vine Borer Family Erebidae—Woollybear Caterpillars, Tiger Moths, and Others Banded Woollybear Green Cloverworm Okra Caterpillar Saltmarsh Caterpillar Yellow Woollybear Family Gelechiidae—Leafminer Moths Eggplant Leafminer Potato Tuberworm Tomato Leafminer Tomato Pinworm Family Hesperiidae—Skippers Bean Leafroller Family Lycaenidae—Hairstreak Butterflies Gray Hairstreak Family Noctuidae—Armyworms, Cutworms, Loopers, Stalk Borers, and Noctuid Moths Alfalfa Looper Army Cutworm Armyworm Bean Leafskeletonizer Beet Armyworm

391 391 391 392 392 393 393 395 398 398 398 400 400 402 403 407 409 410 410 410 413 413 416 418 419 421 422 425 427 428 429 429 431 433 434 436 438 438 439 442 443 445 445 447 447 449 449 450 452 455 456

Bertha Armyworm Bilobed Looper Black Cutworm Bronzed Cutworm Cabbage Looper Celery Looper Clover Cutworm Corn Earworm Darksided Cutworm Dingy Cutworm Fall Armyworm Glassy Cutworm Granulate Cutworm Old World Bollworm Pale Western Cutworm Plantain Looper Potato Stem Borer Hop Vine Borer Redbacked Cutworm Southern Armyworm Soybean Looper Spotted Cutworm Stalk Borer Striped Grass Looper Sweetpotato Armyworm Velvet Armyworm Sweetpotato Leaf Folder Tobacco Budworm Variegated Cutworm Western Bean Cutworm Winter Cutworm Yellow-Striped Armyworm Western Yellow-Striped Armyworm Zebra Caterpillar Family Papilionidae—Celeryworms and Swallowtail Butterflies Black Swallowtail Anise Swallowtail Family Pieridae—Cabbage Worms, White, and Sulfur Butterflies Alfalfa Caterpillar Imported Cabbage Worm Mustard White Southern Cabbage Worm Southern White Family Plutellidae—Diamondback Moths Diamondback Moth Family Pterophoridae—Plume Moths Artichoke Plume Moth Family Pyralidae—Pyralid Moths Lesser Cornstalk Borer Limabean Pod Borer Family Sesiidae—Vine Borers and Clearwing Moths

459 461 462 465 467 471 473 474 479 480 482 486 487 489 491 494 494 494 496 498 500 502 504 506 508 508 510 511 513 516 518 520 520 522 523 523 523 525 525 527 530 531 532 534 534 538 538 539 539 542 543

xii  Contents

Squash Vine Borer Southwestern Squash Vine Borer Family Sphingidae—Hornworms and Sphinx Moths Sweetpotato Hornworm Tobacco Hornworm Tomato Hornworm Whitelined Sphinx Family Tortricidae—Leafroller Moths Pea Moth

543 543 546 546 547 547 550 551 551

11. Order Orthoptera—Grasshoppers, Crickets, and Katydids Family Acrididae—Grasshoppers American Grasshopper Differential Grasshopper Eastern Lubber Grasshopper Migratory Grasshopper Redlegged Grasshopper Southern Redlegged Grasshopper Twostriped Grasshopper Family Gryllidae—Field Crickets Fall Field Cricket Spring Field Cricket Southeastern Field Cricket Family Gryllotalpidae—Mole Crickets Shortwinged Mole Cricket Southern Mole Cricket Tawny Mole Cricket Family Tettigoniidae—Shield-Backed Katydids Mormon Cricket Coulee Cricket

555 555 557 559 562 567 567 569 571 571 571 571 573 573 573 573 576 576 576

12. Order Thysanoptera—Thrips American Bean Thrips Bean Thrips Chilli Thrips

581 581 583

Common Blossom Thrips Florida Flower Thrips Grass Thrips Melon Thrips Onion Thrips Tobacco Thrips Western Flower Thrips

585 586 588 589 591 595 597

13. Other Invertebrate Pests Class Acari—Mites Banks Grass Mite Broad Mite Bulb Mites Twospotted Spider Mite Strawberry Spider Mite Tumid Spider Mite Tomato Russet Mite Class Collembola—Springtails Garden Springtail Class Diplopoda—Millipedes Garden Millipede Class Isopoda—Woodlice (Pillbugs and Sowbugs) Common Pillbug Dooryard Sowbug Class Gastropoda—Slugs and Snails Slugs Snails Class Symphyla—Symphylans Garden Symphylan

601 601 602 604 605 605 605 608 609 609 611 611

Appendix A: Keys to Selected Groups of Pests Appendix B: Vegetable Plant Names Journal Abbreviations and Journal Titles Glossary References Index

627 651 659 665 671 789

613 613 613 615 615 619 622 622

Preface The Handbook of Vegetable Pests was produced as a reference for professionals working in agriculture. Although used mostly as a reference for entomologists, this manual also is handy as a guide for horticulturalists, cooperative extension service personnel, consultants, and others. This book is an update for material accumulating since 2001, consisting mostly of information that has accumulated since that date. About 50 new treatments of pests are included, consisting mostly of recent invaders. The key elements of the Handbook of Vegetable Pests are the introduction, which provides an overview of vertebrate pests that cause injury to crop plants in North America; identification guides, which list the major and minor pests known to attack each crop plant; pest profiles, which describe the appearance, life history, and management of the pests; a­ ppendices, which provide some simple keys to important taxa; glossary, which defines many scientific terms used in the Handbook; references, a list of journals, books, and other publications cited in the pest profile section; and index, which will assist in finding entries in the Handbook. Particularly noteworthy is the abundance of new journals that have proliferated in recent years, many of which are available only in electronic form.

xiii

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Acknowledgments I would like to acknowledge Lyle Buss for the photographic support he provided, which proved to be extremely valuable.

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Chapter 1

Introduction

NORTH AMERICAN VEGETABLE CROPS The production of vegetable crops is a fundamental industry, a popular hobby, and an important element of our culture. The farm gate value of North American vegetable crops exceeds $13 billion annually (2016 estimate), and fuels several other industries such as equipment manufacture, transportation, and retail and restaurant food sales worth many times the value of the raw materials. Vegetable production also has immense value as a form of recreation and relaxation. Surveys of recreational habits consistently reveal gardening to be the number one American hobby. Also, food is an important element of all cultures, and to be able to access diverse and freshly harvested produce, and perhaps to share it with family and friends, remains a desirable and cherished behavior among North Americans. Because the United States and Canada are populated primarily by people whose ancestors migrated from another country in relatively recent times, there are many specialty crops desired by one ethnic group or another that are not yet part of the mainstream of commerce. Thus, local or home garden production of specialty vegetables flourishes in many ethnic communities. For example, Asian vegetables are too small a component of vegetable commerce to be tabulated separately by statisticians concerned with vegetable production. This will change, however, as increase in Asian and Latin American vegetables is evident in American markets. Broccoli was rare outside Italian-American communities until the 1950s, and has since become one of the most popular vegetables, so it is safe to predict that other changes are imminent as North American consumers become more cosmopolitan in their dietary habits.

What Are the Major Vegetable Crops? Numerous types of vegetables are available to the people of the United States and Canada. Availability has changed radically during the 20th century. Early in the century, a Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00001-7 © 2020 Elsevier Inc. All rights reserved.

great deal of vegetable production was local, people ate what was seasonally available, and in the winter months ate only what stored well in root cellars or could be preserved through canning. With the improvement in transportation in the middle of the century, southern vegetable production areas could keep northern consumers supplied with fresh vegetables through most of the winter season. Now fresh produce not only is produced in southern states, but increasing quantities are being imported from Mexico, Central America, and even Southern South America. Complete agreement is lacking on what plants should be called vegetables. In government statistics, potato and sweet potato tend to be grouped with field or staple crops such as wheat, barley, and rice. However, most people think of potato and sweet potato as vegetables. Dry beans such as pinto bean and navy bean are also usually treated as field crops, whereas the same species grown for edible pods as bush beans and pole beans are considered vegetables, and even dry beans are thought of as vegetables by many. Strawberry is treated as a vegetable in some statistical compilations, but most people think of strawberry as a fruit. Also, there are plants such as parsley and coriander that might be considered herbs rather than vegetables because they are used mostly for seasoning. What vegetables do North Americans prefer to eat? Despite an increase in importation of vegetables from other countries, examination of the commercial production data (area planted and value of crops) provides a good indication of what is popular. Following are estimates of fresh market vegetable production data for the most important vegetable crops grown in the United States for 2016–17 (Table 1.1). The relative importance of processed (canned and frozen) vegetables varies considerably from the fresh market values. In some cases (e.g., pea, beet, lima bean, sweet corn, tomato), a substantial portion of the crop is diverted to processing, whereas with other vegetables (e.g., potato, sweet potato, cucumber, cabbage) nearly all is sold as fresh produce.

1

2  Handbook of Vegetable Pests

TABLE 1.1  Summary of fresh market vegetable production data from United States. Area planted Crop

Acres

TABLE 1.2  Leading fresh market producing states (United States). % of nation

Value Hectares

$ millions

State

Area harvested

Value

Artichoke

6800

2747

54

California

43.7

53.4

Asparagus

23,300

9413

73

Florida

10.0

15.1

Beans, lima

28,000

11,300

23

Georgia

6.5

6.6

Beans, snap

236,000

95,344

441

Arizona

6.4

3.8

Broccoli

129,400

52,277

750

Texas

4.4

3.1

Cabbage

57,300

23,149

450

Carrot

71,550

28,906

758

Cauliflower

37,000

14,948

389

Celery

25,100

10,140

665

Corn, sweet

482,500

194,700

231

Cucumber

118,000

47,670

344

7000

2828

122

Garlic

33,000

13,332

390

Lettuce

166,800

67,387

1500

Melon, cantaloupe

51,600

20,850

261

Melon, honeydew

13,600

5494

80

138,000

55,752

971

Pepper, bell

43,300

17,493

642

Pepper, chilli

18,900

7635

143

Pumpkin

67,000

27,070

207

1,050,500

516,108

3710

163,300

65,970

705

Spinach

47,500

19,190

282

Squash

37,400

15,109

149

Tomato

311,500

125,846

1670

Watermelon

113,000

45,652

578

Eggplant

Onion

Potato, Irish Potato, sweet

Most vegetable crops can be grown in all areas of the United States and Canada, though some crops are clearly favored by long seasons and hot summers, and others by cooler summers. Home gardeners are remarkable in their ability to nurture vegetables through all sorts of climatic difficulties, but commercial producers tend to gravitate to the ideal climatic conditions for a particular vegetable. Thus, at least in commercial vegetable production, crops are concentrated in a few areas. The leading fresh market vegetable producing states are illustrated in Table 1.2.

The southern or warm-weather states tend to dominate the fresh market vegetable industry because vegetables can be grown only in these locations during the winter months. In contrast, northern states contain a high proportion of the processed vegetable industry, though California leads in processed vegetables as well. The leading producers of processed vegetables in the United States are illustrated in Table 1.3.

TABLE 1.3  Leading producers of processed vegetables (United States). % of nation State

Area harvested

Value

California

20.9

43.7

Wisconsin

17.3

9.8

Minnesota

14.8

7.8

Washington

11.2

7.9

6.9

5.4

Oregon

In Canada, commercial vegetable production similarly is concentrated in a few provinces as presented in Table 1.4. TABLE 1.4  Commercial vegetable production in Canada. Province

Value, % of nation

Ontario

35.6

Quebec

29.8

British Columbia

15.5

Prince Edward Island

7.2

Manitoba

6.5

Introduction Chapter | 1  3

Not evident from these data are the historical trends in consumption. Although a detailed description of dietary changes is beyond the scope of this treatment, salad vegetables such as lettuce, cucumber, and tomato have gained increasing importance. Some vegetables such as potato and sweet corn remain popular, though in the case of sweet corn there has been a shift from canned and frozen to fresh produce. Although some of the cool-weather vegetables such as cabbage, pea, rutabaga, and turnip have decreased in popularity, some such as Brussels sprouts, broccoli, and cauliflower have increased in popularity.

Characteristics of the Major Vegetable Crops Asparagus (Family Asparagaceae, Formerly Liliaceae) Asparagus is a hardy perennial plant, and once established remains productive for many years. It is grown in all except the warmest regions of North America, and is winter-hardy in most climates. It is usually grown from crowns (roots), but seeds may be used. Seeds are spread freely by birds, and it is common to find asparagus growing wild along roadsides, fences and irrigation ditches. California, Washington, and Michigan are leaders in asparagus production, though commercial production is also important in Eastern Canada. Asparagus is popular in home gardens grown in northern areas, probably because it is one of the first crops available for harvest in the spring. It has been an important vegetable since ancient times, and originated in the Mediterranean region. The spears or stems as they first push up from the soil are harvested; once they begin to branch they become tough and inedible. The most serious pest is asparagus aphid, Brachycorynella asparagi (Mordvilko), though its economic impact is mostly a western phenomenon. In Eastern North America, asparagus beetle, Crioceris asparagi (Linnaeus), and spotted asparagus beetle, C. duodecimpunctata (Linnaeus), can be a nuisance.

Bean and Related Crops (Family Fabaceae, Formerly Leguminosae) (Bush Bean, Chickpea, Cowpea, Dry Bean, Faba Bean, Lentil, Lima Bean, Pea, Snap Bean, and Other Beans) Legumes are known for their ability to harbor nitrifying bacteria; nitrogen enhances soil productivity. The cultivated legume vegetable crops are not particularly efficient as a source of nitrogen for plant growth, however, so fertilization is still required. Most of the leguminous vegetable crops are warmweather crops, and are killed by light frosts. Pea is a notable exception, thriving under early season and cool weather conditions, though it is killed by heavy frost. Also, though not often grown in the United States, and principally a home garden crop in Canada, faba bean is commonly grown in the northern regions of Europe. The legume vegetables are annual crops. The most commonly grown bean crops, snap bean and lima bean, are native to Central America. Pea and cowpea

(blackeyed pea) likely originated in Asia. In the United States, snap beans are quite popular, both in fresh and processed form. Florida and California lead in the production of fresh market beans, Minnesota and Wisconsin in processing pea. Cowpea production occurs principally in the southeastern states where historically it was quite important because it remained productive through the long summer months when most other vegetable crops failed. In Canada, pea is a more important crop than is bean, and Ontario and Quebec lead in the production of these crops. The legumes are cultivated for their seeds or seed pods. The below-ground seed attacking maggots, Delia spp., can be important pests under cool weather conditions. The key pest of bean in many areas is Mexican bean beetle, Epilachna varivestris Mulsant. Pea is quite susceptible to infestation by pea aphid, Acyrthosiphon pisum Shinji. Cowpea is plagued by cowpea seed beetles (weevils), Callosobruchus spp., and cowpea curculio, Chalcodermus aenus Beheman, and southern green stink bug, Nezara viridula (Linnaeus). Locally, a number of other pests can be important, particularly thrips, leafminers, leafhoppers, and flea beetles.

Beet and Related Crops (Family Amaranthaceae, Formerly Chenopodiaceae) (Beet, Beetroot, Chard, Spinach, Swiss Chard, Quinoa) Beet apparently originated in the Mediterranean region, and spinach in Iran. Beet and its relatives are biennial crops, requiring more than 1 year but less than 2 to complete their natural life cycle. They are grown as annuals when cultivated as vegetables. Though originally grown entirely for its foliage, cultivars of beets with edible below-ground portions (the edible “root” is mostly thickened stem material) became popular in the Europe beginning about 1800. Beet, once popular in North America, has declined greatly in popularity. Chard has never been an important crop, and remains relatively obscure. Spinach is not very popular, but commercial production is stable. Most commercial cultivation of beets in the United States occurs in the Great Lakes area, but spinach production is more western in distribution, from Arkansas and Texas to California. In Canada, these crops are grown primarily in Ontario, Quebec, and British Columbia. In home gardens, these crops thrive nearly everywhere. The principal pests are green peach aphid, Myzus persicae (Sulzer) and beet and spinach leaf miner, Pegomya spp. In the west, beet leafhopper, Circulifer tenellus (Baker), can be very damaging.

Cabbage and Related Crops (Family Brassicae, Formerly Cruciferae) (Broccoli, Brussels Sprouts, Cabbage, Cauliflower, Chinese Cabbage, Collards, Kale, Kohlrabi, Mustard, Radish, Rutabaga, Turnip) Brassicaceous or cruciferous vegetables, often called “cole” crops, are generally grown for their leaves. These are coolseason crops, and tolerate light freezes and even brief heavy

4  Handbook of Vegetable Pests

freezes, but prolonged deep freezes are fatal. Though naturally biennials, they are grown as annuals. Cabbage and its many forms originated along the shores of Europe; mustard and radish are from Asia. Some are popular foods in North America, others are of regional significance. Perhaps the most interesting vegetable in this group is broccoli, which has become popular only since the 1950s. Cauliflower is a moderately important crop that is increasing in popularity. Cabbage and turnip, once popular in the United States, have declined in importance, though cabbage remains a significant crop. Rutabaga is an important dietary element in Canada, but not in the United States. Collards, and to a lesser degree kale, are popular vegetables in the southeastern states. In the United States, commercial production of broccoli, cauliflower, and Brussels sprouts is concentrated in California. California and Florida are the primary producers of radish. Cabbage is produced widely. In Canada, Ontario, and Quebec are the important producers of these crops. In home gardens, these crops are produced throughout North America. The principal pests of cabbage and its closest relatives include the root maggots, Delia spp.; cabbage aphid, Brevicoryne brassicae (Linnaeus); diamondback moth, Plutella xylostella (Linnaeus); cabbage looper, Trichoplusia ni (Hübner); and imported cabbageworm, Pieris rapae (Linnaeus). Mustard and radish tend to be plagued more by green peach aphid, Myzus persicae (Sulzer).

Carrot and Related Crops (Family Apiaceae, Formerly Umbelliferae) (Carrot, Celery, Celeriac, Chervil, Cilantro, Fennel, Parsley, Parsnip) The apiaceous (umbelliferous) vegetables are biennial, but are grown as annuals. They require cool weather to develop properly. Most survive heavy frost but are killed by prolonged freezing weather. The major crops of this group, carrot and celery, originated in the Mediterranean region. Carrot and parsnip are grown for their root, celery, celeriac and fennel for the swollen stem bases, and parsley and cilantro for their foliage. These crops are grown widely in the United States, in both northern and southern locales, but California dominates commercial production. In Canada, commercial production occurs predominately in Quebec and Ontario. Carrot and celery have long been popular vegetables. Parsnip once was an important crop because it could be stored well during the winter months, and could even be left in the soil during freezing weather. While Canadians and Europeans retain a fondness for this crop, Americans rarely consume it. Parsley retains its utility in the American diet, but cilantro is rapidly growing in popularity as Mexican and Asian recipes gain broader acceptance. Fennel is similarly growing in popularity, but is yet a minor crop awaiting discovery by the American palate. Chervil

is rarely grown in North America, and different types are grown for their root and foliage. In some areas, carrot weevil, Listronotus oregonensis (LeConte), and carrot rust fly, Psila rosae (Fabricius), are important pests, and leafhoppers sometimes transmit aster yellows disease. American serpentine leafminer, Liriomyza trifolii (Burgess), is often the most serious threat to commercially grown celery.

Lettuce and Related Crops (Family Compositae) (Artichoke, Celtuce, Chicory, Endive, Escarole, Lettuce, Radicchio) Lettuce is an immensely popular vegetable. Among vegetables grown in the United States, it surpasses all other vegetables except for potato in area of land devoted to production and in crop value. Canada lacks the warm-weather production areas found in the United States, so it is not as important a commercial crop there. It is popular among Canadian consumers and much lettuce is imported from the United States. California and Arizona dominate commercial production in the United States; Quebec is the major producer in Canada. Lettuce apparently originated in Europe or Asia, and has been grown for over 2000 years. Lettuce is grown everywhere in North America for home consumption, and despite the apparent concentration of lettuce production in few states or provinces, specialty lettuces are grown around many large metropolitan areas for local consumption. Lettuce and most related crops are grown for their leaves, though in the case of celtuce the stem is eaten. Lettuce and related crops are ­cool-season annuals. Although killed by heavy frost, these crops are also susceptible to disruption by excessive heat. Hot weather causes lettuce to flower and become bitter tasting. Several insects are important pests of lettuce. Aster leafhopper, Macrosteles quadrilineatus Forbes, is an important vector of aster yellows in some production areas. Several species of aphids may be damaging, though green peach aphid, Myzus persicae (Sulzer), generally is most important. Numerous caterpillars such as corn earworm, Helicoverpa zea (Boddie), and cabbage looper, Trichoplusia ni (Hübner), threaten the lettuce crop. Artichoke, more correctly known as globe artichoke, is a thistle-like plant grown for the edible blossom bud. It is one of only a few vegetable plants grown in North America as a perennial, with new growth arising from the roots annually. Like some other perennials, it can be grown with some success as an annual crop by planting roots, but this is not a common practice. Although artichoke can be grown over a broad geographic area it is not cold-hardy. Commercial production is limited to the California coast, where the cool, moist climate favors its growth. It is not a popular vegetable, and considered by many to be a “luxury” vegetable. The origin of artichoke appears to be the Western Mediterranean region of

Introduction Chapter | 1  5

Europe. A similar plant grown for the leaf stalks is cardoon; this plant is not grown commercially in North America. The key pest of artichoke is artichoke plume moth, Platyptilia carduidactyla (Riley), though aphids such as artichoke aphid, Capitophorus elaeagni (del Guercio); and bean aphid, Aphis fabae Scopoli, can be quite damaging at times.

where summers are cool. The stalks or leaf petioles are used as food, though this vegetable is infrequently consumed. Most commercial production occurs in northern states such as Michigan, Oregon, and Washington, but it remains ­principally a home garden crop. There are few important pests of rhubarb, with rhubarb curculio, Lixus concavus Say, perhaps the most serious pest.

Okra (Family Malvaceae)

Squash and Related Crops (Family Cucurbitaceae) (Cucumber, Pumpkin, Squash, Cantaloupe, Watermelon, and Other Melons)

Okra is thought to be native to Africa and is an important crop in tropical countries. It is also an important element of southern cooking because it is one of the few vegetables that remains productive throughout the long summer of the southeast. It is an annual plant and is killed by light frost. Okra is grown for the seed pod which, like snap bean, is harvested before it matures. It is unusually tall for a vegetable crop, often attaining a height of 2 m. It is a relatively minor vegetable crop from a national perspective and so production statistics are infrequent. Commercial production in the United States occurs in the southeast from South Carolina to Texas. The pods are subject to attack by several pests, with the most damaging being red imported fire ant, Solenopsis invicta Buren; southern green stink bug, Nezara viridula (Linnaeus); and leaffooted bugs, Leptoglossus spp.

Onion and Related Plants (Family Amaryllidaceae, Formerly Alliaceae) (Chive, Garlic, Leek, Onion, Shallot) Onion and its relatives are biennial or perennial plants, but cultivated as annuals. These have long been important crops, with use of onion documented for nearly 5000 years. Their origin is thought to be Asia. Onion is grown principally for the below-ground leaf bases, where a bulb is formed, but tops are also edible. They are tolerant of cool weather, but also thrive under hot conditions. Onion is cultivated widely in North America, though commercial production of the sweet varieties tends to be concentrated in southern areas, whereas the pungent varieties are cultivated at more northern latitudes. In the United States, leading producers are California, Washington, Texas, Colorado, New York, and Georgia. California also leads the nation in garlic production. Ontario and Quebec produce most of the onions grown in Canada. The key pests of onion are onion thrips, Thrips tabaci Lindeman, and onion maggot, Delia antiqua (Meigen).

Rhubarb (Family Polygonaceae) Rhubarb is one of the few perennial vegetable plants that is cultivated as a perennial. Its origin is Northern Asia, and its use can be traced back about 5000 years. Rhubarb thrives

The cucurbit crops are important vegetables, though they vary in economic importance. They are annual plants, and are warmth-loving crops. All are cultivated for their fruit. Light frost will kill cucurbits, and even cool weather will permanently disrupt growth. Squash and pumpkin originated in Central and South America; cucumber, melons, and watermelon are from Africa or perhaps Asia. The summer squashes and cucumbers are stable in production or increasing slightly in importance, though squash is considered to be a minor vegetable crop and statistics are scant. The winter squashes are of decreasing significance in diet and commerce. Pumpkin is grown mainly as an ornamental plant, though some is processed for food. All types of melons are increasing in popularity and economic significance. Cucumbers are produced in many states, but Georgia and Florida lead in fresh market production and Florida in cucumbers for pickles. Texas, Florida, and Georgia are the leading producers of watermelon, but commercial production is widespread. Cantaloupe and other melon production is centered in California and Arizona. Cucurbit crops have some serious insect pests. Home garden plants are plagued by squash vine borer, Melittia cucurbitae (Harris); squash bug, Anasa tristis (De Geer); and in the southeastern states by pickleworm, Diaphania nitidalis (Stoll). Commercial crops are affected by whitefly and aphid plant virus vectors; Diabrotica spp. and Acalymma sp. cucumber beetles in the north central states; and pickleworm in the southeast.

Sweet Corn (Family Poaceae, Formerly Graminae) Corn, which is usually known as maize outside of the United States, apparently was domesticated in Mexico, and perhaps has descended from a similar grain, teosinte. Corn originally was cultured because it was productive, a good source of carbohydrates and other nutrients, and the grain stored well. Sweet corn is a recent innovation that was first developed in the mid-1700s, and lacks the storage characteristics of the older types, or grain corn. Corn is grown for the seed, which are clustered in a structure called the “ear.” Sweet corn is a popular vegetable, though

6  Handbook of Vegetable Pests

canned sweet corn is less popular than it once was due to the availability of frozen corn. The more recent availability of fresh corn that does not quickly lose its sweetness (supersweet cultivars) has increased demand for wholeyear by fresh or frozen. Sweet corn is cultured widely in North America. In the United States, Florida, California, New York, and Georgia lead in fresh sweet corn production, whereas Minnesota and Wisconsin lead in processed sweet corn. In Canada, the provinces producing most of the sweet corn are Ontario and Quebec. There are many insects that feed on grasses, and many of them have moved from feeding on wild grasses to feeding on corn. Thus, native species such as corn earworm, Helicoverpa zea (Boddie); fall armyworm, Spodoptera ­frugiperda (J.E. Smith); and Diabrotica rootworms are the most serious pests. However, some introduced insects have also become frequent pests, including European corn borer, Ostrinia nubilalis (Hübner); Japanese beetle, Popillia ­japonica Newman; and corn leaf aphid, Rhopalosiphum maidis (Fitch).

Sweet Potato (Family Convolvulaceae) Sweet potato is an immensely important crop in some parts of the world, but not in North America, where acreage and consumption are declining. Sweet potato probably originated in Mexico and is well adapted to tropical growing conditions. Some moist-fleshed varieties of sweet potato are called yams, but yams are a separate species normally found in Polynesia, and infrequently seen in North America. A perennial crop, sweet potato is normally grown as an annual. It is cultivated for its tuber. Sweet potato cannot tolerate prolonged cool weather and perishes if exposed to light frost. Most of the domestically produced sweet potatoes are grown in North Carolina and Louisiana. Sweetpotato weevil, Cylas formicarius (Fabricius), is the most damaging pest of this crop, but anything that damages the tubers, including wireworms and flea beetle and cucumber beetle larvae, is economically threatening.

Tomato, Potato, and Related Plants (Family Solanaceae) (Eggplant, Pepper, Potato, Tomatillo, Tomato) The solanaceous crops are among the most popular vegetable crops in North America. Potato ranks as the most valuable vegetable crop grown in the United States. In Canada, potato and tomato are first and second in importance, but warm-season crops such as pepper do not thrive in Canada. Among home gardeners, tomato is the most popular crop. Potato is grown for its tuber, and the other crops for their fruit. The origins of the solanaceous crops are diverse; eggplant originated in India, potato and tomato in Peru, tomatillo and pepper in Mexico or

Guatemala. Tomato, tomatillo, pepper, and eggplant are warm-season perennials that are cultivated as annuals. Potato is a cool-season perennial cultured as an annual. Potato in most of North America is at risk from Colorado potato beetle, Leptinotarsa decemlineata (Say), but several aphids also are commonly damaging, particularly green peach aphid, Myzus persicae (Sulzer). In the west, beet leafhopper, Circulifer tenellus (Baker), can be damaging. Tomato is affected by silverleaf whitefly, Bemisia argentifolii Bellows and Perring; corn earworm, Helicoverpa zea (Boddie); thrips-transmitted plant diseases, and many other pests. Home garden production, but not commercial production, is at risk from tobacco and tomato hornworms, Manduca spp. In southern production areas, pepper weevil, Anthonomus eugenii Cano, is the key pest of pepper.

INSECTS AND INSECT RELATIVES With over 90,000 insect species inhabiting the United States and Canada, it might seem to be an impossible task to identify one specimen among so many possibilities. However, a surprisingly small number of insects injure plants, and even fewer damage vegetable crop plants. About 300 species are documented to be pests, though a few more are capable of damage, or feed on vegetables only rarely. Insects are members of the group of animals collectively classified as Insecta. Insecta is a major class of animals, much like other major groups such as Reptilia (reptiles), Mammalia (mammals), Aves (birds), Pices (fish), and Gastropoda (snails and slugs). Insecta is subdivided into major groups called orders, and further subdivided into smaller groups of like organisms called families. Occasionally, further subdivisions are noted, such as subfamilies, but the most important designations, other than the order and family of insects, are the genus and species names. The genus and species designation (sometimes along with the original describer or species “author”) is also called the scientific name, and is used to provide universal recognition. Common names often vary among different regions of the world, and insects even acquire different names depending on the crop attacked, but scientific names are universally accepted and are changed rarely.

Frequency of Pests Among the Major Insect Orders The occurrence of vegetable pests among the orders of insects is more or less proportional to the total number of species in each order. The approximate number of species in each major order inhabiting the United States and Canada, and the number of species within the corresponding order that are considered in this book are presented in Table 1.5.

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TABLE 1.5  Frequency of species and coverage in this book. Number of species in United States and Canada

Number treated in this book

Coleoptera (beetles)

24,000

106

Hymenoptera (ants, bees, and wasps)

18,000

3

Diptera (flies)

17,000

30

Lepidoptera (moths and butterflies)

12,000

95

Hemiptera (bugs)

10,500

77

1000

15

Insect order

Orthoptera (grasshoppers and crickets) Thysanoptera (thrips)

700

10

Blattodea (cockroaches and termites)

100

5

20

3

Dermaptera (earwigs)

The order Hymenoptera, and to a lesser degree the order Diptera, contain many beneficial species (parasitoids and pollinators), accounting for their underrepresentation as vegetable pests.

Noninsect Vegetable Pests There are several other groups of invertebrate animals that act like insects, sometimes inflicting chewing injury to plant foliage, fruit, and roots. Some even resemble insects in general appearance. Therefore, they are included in this treatment of “insect” pests. Best known among these other invertebrates are the snails, slugs, mites, millipedes, sowbugs, and pillbugs.

Order Blattodea—Cockroaches and Termites Traditionally, the cockroaches are placed in the order Blattodea (or a similar name) whereas the termites are placed in the order Isoptera. However, it is now believed that they had common ancestors, even though they look quite different. Indeed, they have some common features. We think of termites as social insects that feed on wood. Some cockroaches also display elements of social behavior and can digest cellulose quite efficiently, thanks to m ­ icroorganisms

in their digestive system. So they are quite similar functionally, despite looking quite different. It is useful to think of cockroaches as primitive termites, or termites as a highly evolved group of cockroachs. In North America, there are about 10 species that we would consider to be pest termites of vegetables, and 5 that would be considered pest species of cockroaches. They are basically tropical insects, so they are more commonly found in warmer regions. Cockroaches and termites are not nomally considered to be vegetable pests, but it would be more correct to consider them not to be major vegetable pests. In the tropics, termites do quite a lot of damage to crops, so we should not be entirely surprised to occasionally see damage to our crops by termites. Likewise, some cockroaches are known to nibble on leaves of plants, especially damaging seedlings.

Order Coleoptera—Beetles, White Grubs, and Wireworms Coleoptera is the largest order of insects, containing about 40% of the known species in the class Insecta. The most distinctive feature is the structure of the wings. The front pair of wings, called elytra, are normally thickened and hard, providing protection for the thinner, membranous hind pair of wings. The hind wings are longer than the front wings, and fold beneath the forewings when the insect is not in flight. Beetles have chewing mouthparts, with well-­ developed mandibles. The immatures undergo complete metamorphosis, displaying egg, larval, and pupal stages before attaining adulthood. Their behavior and ecology are diverse. Larvae are soft bodied, and usually bear three pairs of legs on the thorax, and no prolegs on the abdomen. Larvae often develop in protected habitats such as within stems and soil, but some are found feeding on foliage. About 24,000 species are known from the United States and Canada.

Family Carabidae—Ground Beetles Although the pest species are somber brown or black in color, carabids are sometimes brightly colored and metallic. They tend to be slightly flattened and long legged, with the elytra marked with long grooves and ridges. They vary widely in size. The antennae are thread-like. Nearly all carabids are beneficial, with both the larval and adult stages feeding on other insects. Larvae generally inhabit the soil. They have a thin, elongate body with a large head. About 2300 species are known from the United States and Canada.

Family Chrysomelidae—Leaf Beetles This is a large and diverse family. Many are brightly colored. They range in length from 1.5 to 13 mm. The body tends to be oval and strongly convex. The antennae and legs are moderately long to short. The antennae are thread-like or slightly expanded apically, but never clubbed. About 1500

8  Handbook of Vegetable Pests

species of chrysomelids occur in the United States and Canada. As an aid to identification, two of the easily recognizable subfamilies are treated separately in the following. Subfamily Alticinae—Flea Beetles Flea beetles are a distinctive group of leaf beetles. They derive their name from the impressive leaping behavior of adults, which is made possible by muscular, greatly enlarged hind femora. These dark-colored beetles are small, usually less than 5 mm in length, sometimes considerably smaller. They may be metallic, or even brightly colored on occasion. They tend to leave small round holes in leaves when they feed, sometimes consuming only one surface of the leaf and leaving the other intact. Most larvae live belowground, feeding on the roots of plants. Subfamily Bruchinae—Pea and Bean Seed Beetles Adult bruchids have a short, broad downturned snout bearing mouthparts at the tip. These beetles are small, measuring no more than 6.5 mm in length, and generally less than 5 mm. Their body tapers toward the head. The antennae are slightly expanded at the tips, or serrate. The elytra extend only about two-thirds the length of the abdomen, exposing the terminal segments. The larvae live within the seeds of legumes. They possess legs only during the first instar; thereafter they are legless, plump, and humpbacked. There are over 130 species of bruchids known from the United States and Canada. Subfamily Cassidinae—Tortoise Beetles Tortoise beetles are distinguished by the flaring of the pronotum and elytra, which largely cover the head and legs when viewed from above. This protected, armored, or turtle-like appearance is the basis for the common name of this group of leaf beetles. Many species are brightly colored, often orange or gold. Some species can be confused with lady beetles, family Coccinellidae. However, in lady beetles the head is clearly visible. Also, in lady beetles there are three tarsal segments on each leg, whereas tortoise beetles appear to have four tarsal segments. Larvae are surface feeders, found on leaves alongside adults. They bear numerous branched spines, and often carry debris or fecal material on their backs. Other Subfamilies—Leaf Beetles A few other subfamilies of Chrysomelidae contain pests, but they are not so readily distinguished as are the aforementioned subfamilies. For example, subfamily Criocerinae contains the asparagus-feeding species, Galucerinae contains the corn rootworms and cucumber beetles, and Chrysomelinae contains Colorado potato beetle. As with the other chrysomelids, the adults feed on above-ground plant tissue; larval-feeding behavior is variable. The adults are often brightly colored and moderate in size, often measuring 4–12 mm in length. The antennae are usually moderately long in length.

Family Coccinellidae—Lady Beetles This family is best known for its beneficial, predatory members. However, a small number are plant feeders. Most lady beetle species are brightly colored and spotted, but their general pattern is misleading; not only are some important predators spotless, but the spot pattern varies greatly even within a species, reducing its diagnostic value. The legs are short or moderately long, and bear only three tarsal segments. The antennae are clubbed. The body is strongly convex, almost hemispherical in shape. There are about 400 species known from the United States and Canada. The plant feeding species have larvae which bear large branched spines.

Families Curculios, Weevils, and Brentidae—Weevils The members of this family are distinguished by their elongated snout, which bears mouthparts at the tip. The elbowed, clubbed antennae are attached at about the midpoint of the snout. The body form varies among species, but is usually elongate-oval. These beetles tend to be moderate in size, ranging from 3 to 12 mm, but sometimes considerably larger. They usually are dark in color, with the elytra attaining the tip of the abdomen. This latter character is useful for distinguishing curculionids from bruchids. The plump larvae lack legs, and normally are found burrowing within plant tissue. Over 2600 species are known from the United States and Canada.

Family Elateridae—Click Beetles and Wireworms Click beetles are elongate, parallel-sided, and usually relatively flattened in appearance. The pronotum is relatively large, and as wide as the thorax. They bear a structural mechanism that allows them to flex rapidly, producing a clicking noise. The head is usually hidden below the pronotum. The antennae are usually serrate, but not clubbed. The elytra bear ridges and grooves. These beetles usually are obscurely colored, often brown and black. The beetles are usually moderate or large in size, measuring 12–30 mm in length. Although the adults are phytophagous, it is the larval stage, known as wireworms, that is most destructive. The larvae are slender (hence the name wireworm), round in cross section, hard-bodied, and shiny. They feed on the roots of plants. About 900 species are known from the United States and Canada.

Family Meloidae—Blister Beetles These elongate, narrow-bodied beetles are unusual in that their elytra are soft and flexible. The pronotum is narrower than the head and the thorax. The tips of the elytra tend to diverge. The legs and antennae are long and thread-like. Blister beetles are moderate to large in size, measuring 12–25 mm in

Introduction Chapter | 1  9

length. The adults feed on foliage and blossoms but the larvae live below-ground where they feed on insects; the common species feed on the egg pods of grasshoppers. About 310 species are known from the United States and Canada.

Family Nitidulidae—Sap Beetles These beetles tend to be small to moderate in size, less than 12 mm in length. They are oval in shape, and often distinguished by short elytra which expose the terminal abdominal segments. Also, the antennae are distinctly clubbed. Both larvae and adults feed on decaying fruit. About 185 species occur in the United States and Canada.

Family Scarabaeidae—Scarab Beetles and White Grubs The most distinctive feature of scarab beetles is the shape of the antennae. The three terminal segments are expanded into plate-like or finger-like structures that can be closed to form a club. Many species have long, spiny legs. Scarab beetles vary widely in size, from 2 to over 100 mm. The May and June beetles are oval and strongly convex. Others, however, are more flattened, with a wide prothorax and an abdomen that tapers acutely toward the posterior end. Most species are drab in color, usually brown or black. Some, however, such as Japanese beetle, are brightly colored and metallic. The adults feed on above-ground foliage, blossoms, and fruit. Larvae are plump and C-shaped in general form, usually living below-ground on roots of plants. About 1400 species of scarabs live in the United States and Canada.

Family Tenebrionidae—Darkling Beetles and False Wireworms Darkling beetles typically inhabit arid environments, particularly the western regions of North America. Adults tend to be long-legged with a hump-backed appearance. They generally are black in color. Their antennal structure is variable. Darkling beetles are easily confused with carabid beetles, but can be distinguished by their tarsal structure. Darkling beetles vary from 2 to 35 mm in length. About 1000 species occur in the United States and Canada. Although adults feed on plants, larvae are more destructive due to their tendency to attack seedlings. The larvae are called false wireworms because they are narrow and elongate in form, resembling wireworms in general appearance.

Order Diptera—Flies and Maggots The flies are notable in possessing only a single pair of wings. The wings are membranous. The second pair is reduced to a pair of small knobbed structures, called halteres, that function to provide balance. Flies tend to be small and fragile. The mouthparts are variable, but usually function to sponge up liquids or are needle-like and

are used to pierce and suck up liquids. The antennae are variable. They undergo complete metamorphosis, displaying egg, larval, and pupal stages in addition to the adult, or fly, form. The larvae generally are worm-like and legless, and are called maggots. Maggots are usually quite featureless, but the anterior end is equipped with hook-like structures that are used to tear host tissue. Pupation occurs within the old larval exoskeleton (outer covering of the maggot) and is called a puparium to distinguish it from the pupa found in most other insects, and which lacks the larval covering. About 17,000 species of flies are known from the United States and Canada.

Agromyzidae—Leaf Mining Flies These small black or grayish flies usually measure less than 2 mm in length, and are often marked with yellow. The wings lack color. Leafminers are best known for the winding mines or tunnels created by larvae as they feed between the upper and lower surfaces of leaf tissue. The typical mine is narrow, grows wider as the larva increases in size, but some species cause irregular, blotchy mines. About 190 species of Agromyzidae are known from the United States and Canada.

Drosophilidae—Pomace Flies These small fruit flies are also known as vinegar flies because they are attracted to fermenting fruit and vegetables. They are small in size, rarely exceeding 3 mm in length. The wings are colorless, broad, and have few veins. The antenna bears a hair (brista) that is generally plumose. About 120 species of Drosophilidae are known from the United States and Canada.

Ulidiidae—Picture-winged Flies These medium-sized flies attain a length of up to 12 mm. They have wings that are marked with broad bands of color. The fly’s body is often shiny or metallic in appearance. Larvae are found in fruits and roots, or on decaying tissue. Many species are not capable of injuring healthy plants, but are incorrectly implicated as being damaging because they are found in rotten tissue. About 125 species occur in the United States and Canada.

Psilidae—Rust Flies These small- to medium-sized flies measure 3–8 mm in length. They tend to be rather slender, and bear long antennae. Their wings are not colored. The only pest species is carrot rust fly, Psila rosae (Fabricius), which is covered with a dense coat of hairs, and larvae burrow in roots. Only 34 species are known from the United States and Canada.

Syrphidae—Flower Flies The adults of many species in this common family of flies are brightly colored, often black and yellow, and are sometimes mistaken for bees. Most species measure 10–20 mm in length,

10  Handbook of Vegetable Pests

and their wings are not marked. The larvae of some species are predatory, and are often found feeding within colonies of aphids. A few species, however, feed on the ­below-ground portions of plants. The number of syrphid species inhabiting the United States and Canada is about 875.

Tephritidae—Fruit Flies These medium-sized flies often attain a length of 10–12 mm. Their wings are spotted or banded. Their larvae usually develop within fruit, and some species are severe fruit pests, especially in tropical areas. A few species mine leaves. About 280 species occur in the United States and Canada.

Family Cydnidae—Burrower Bugs Burrower bugs are oval, resembling stink bugs in general body form, though they lack the elongate lateral lobes of the pronotum usually present in true stink bugs, family Pentatomidae. The scutellum of burrower bugs is enlarged, though not as large as found in the negro bugs, family Thyrecoridae. Perhaps the most distinctive feature is the numerous spines on the tibiae, which likely aid in burrowing through the soil, its major habitat. There are about 35 species in North America. Most are small, less than 8 mm in length, and blackish in color. They are nocturnal, and uncommonly observed except at lights. Only one species, Pangaeus bilineatus (Say), is known as a pest.

Order Hemiptera—Bugs The suborders of Hemiptera include Heteroptera (water bugs, coreids, lygaeids, stink bugs, etc.), Cicadomorpha (cicadas, spittlebugs, leafhoppers, etc.), and Sternorrhyncha (white flies, aphids, scales, mealybugs, psyllids, etc.). Other taxa are possible. The Hemiptera are distinguished from other group of four-winged insects by the presence of long piercing-sucking mouthparts in the form of a “beak” that originates at the front of the head. In addition, the basal portion of the front pair of wings (the corium) is thickened and usually without veins, whereas the apical portion is thin, membranous, and normally bears veins. The hind wings are entirely membranous and also bear veins. The wings normally are held flat over the back when the insects are at rest, not angled or roof-like. Insects in the order Hemiptera can be confused with insects in the order Cicadomorpha and Sternorrhyncha which also possess tubular piercing-sucking mouthparts. In these taxa, however, the mouthparts originate at the back of the head rather than the front of the head, and the entire front wing is membranous. Many Hemiptera have scent glands that produce strong odors. Some species possess short wings, or are wingless. Not all Hemiptera feed on plants; many are predatory, feeding on other insects. Metamorphosis in Hemiptera is incomplete, with the adult stage preceded by the egg and nymphal stage, but the pupal stage lacking. The immatures, or nymphs, greatly resemble the adults in form, though lacking fully developed wings. Their food and habitat is the same as in the adult stage.

Family Lygaeidae—Seed Bugs

Family Coreidae—Leaffooted and Squash Bugs

The stink bugs derive their name from disagreeable odors which are produced when the bugs are handled. They are also known as shield bugs, a reflection of their overall shape, and a more distinguishing character than their odor. These bugs are moderate to large in size, usually 6–23 mm in length. Stink bugs possess a triangular structure behind the prothorax and at the base of the wings known as a scutellum; it reaches to the midpoint of the abdomen or beyond. The number of mouthpart (beak) segments is 4, and the number of antennal segments is 5. Stink bugs may be

These bugs tend to be fairly large, usually 10–30 mm, and dark colored. If disturbed they tend to produce a foul-­ smelling odor. The antennae and mouthparts each bear four segments. The legs are long, and are flattened in some species. The membranous portion of the forewings bear six or more veins. They tend to be dull colored, usually brown or black. Only a few of the approximately 120 species ­occurring in the United States and Canada are destructive to vegetables.

Lygaeid bugs resemble coreids, but usually are small, often 4–12 mm in length. Sometimes they are colorful. As suggested by the common name, most species are seed feeders, though some suck plant sap or are predatory. The species damaging vegetables are less than 5 mm in length. The antennae and mouthparts (beak) each bear four segments. The membranous portion of the front wing bears only 4–5 veins, a character that is useful for distinguishing them from the Coreidae. In some species, the femora of the front legs are enlarged. About 300 species are known from the United States and Canada.

Family Miridae—Plant Bugs These common bugs tend to be narrow-bodied and elongate. They are moderate in size, measuring 4–10 mm in length. They are soft-bodied relative to most other bugs. The antennae and mouthparts (beak) each consist of four segments. The veins at the tips of the forewings are few in number, forming semicircular loops originating and ending in the thickened basal portion of the wing (the corium), rather than divergent or parallel veins terminating at the wing margin. Also, the apical portion of the corium is triangular, and separated from the rest of the corium by a groove. Mirids are normally plant feeders, but some feed opportunistically on insects.

Family Pentatomidae—Stink Bugs

Introduction Chapter | 1  11

plant feeders, especially seed feeders, or predators. The predatory species tend to have a thick beak, and the phytophagous species have a thin beak. There are about 250 species in the United States and Canada.

certainly not resembling the adult. The nymph often produces waxy filamentous secretions; it generally is sedentary. About 100 species are known from the United States and Canada.

Family Thyreocoridae—Ebony Bugs

Family Aphididae—Aphids

These small oval bugs resemble stink bugs, and are often confused with beetles. They measure only 3–6 mm in length and possess a greatly enlarged scutellum that covers most of the abdominal segments and wings, superficially resembling the elytra found on beetles. They usually are black in color. The antennae consist of five segments. Only about 30 species are known from the United States and Canada.

Aphids are small to moderate in size, normally measuring 2–3.5 mm in length. They undergo complex life cycles, often alternating between winged and wingless generations and between perennial and annual host plants. Aphids usually occur in colonies, sometimes attaining very high densities. Most bear a pair of tubular structures called cornicles near the tip of the abdomen. They secrete from the anus sugary secretions that cause stickiness and discoloration on foliage. The thread-like antennae are moderate in length, and consist of three to six segments. Some species produce waxy secretions that obscure their general appearance, causing them to resemble tufts of cotton or other inanimate objects. Occasionally they live below-ground on roots, though generally they feed on leaves and young stem tissue. They are important as direct pests of vegetable crops due to the plant sap they ingest, but they also transmit plant viruses very effectively, which greatly exacerbates their economic impact. Nearly 1400 species are known from the United States and Canada.

Family Tingidae—Lacebugs Lacebugs are probably the most distinctive of the Hemiptera. The adults possess ornate, lacy wings consisting of many small cells. In some species, the pronotum is expanded and similarly ornate. The immatures are spiny. These are small insects, usually 3–5 mm in length. The antennae and mouthparts (beak) each consist of four segments. Most species feed on trees, and about 160 species are known from the United States and Canada. The suborders Cicadomorpha and Sternorrhyncha consists of a diverse assemblage of insects that are difficult to characterize because superficially they are so different in general appearance. They are closely related to Hemiptera, and treated by some as members of that order. The mouthparts of these taxa are the piercing-sucking type, but instead of arising at the front of the head arise in basally, at the back of the head. Sometimes the mouthparts appear to originate between the base of the front pair of legs. Normally there are four wings of relatively uniform texture throughout, though in some the front wings are slightly thickened. When at rest, the wings are usually held angled or roof-like over the body, not flat over the back as is typically found in Hemiptera. The antennae may be short or bristlelike, or long and thread-like. Metamorphosis is incomplete, with the egg, nymphal, and adult stages present. Nearly 7000 species are known from the United States and Canada.

Family Cicadellidae—Leafhoppers and Family Delphacidae—Planthoppers

Family Aleyrodidae—Whiteflies

These small- to medium-sized insects rarely exceed 12 mm in length and are narrow-bodied in shape. They generally have a sharp or bluntly pointed head, The wings are normally fully formed, extending the length of the abdomen, but occasionally they are short-winged. The front wings are slightly thickened. The antennae are thread-like, originating between the eyes in Cicadellidae and beneath the eyes in Delphacidae. These insects produce sound, but it is barely audible to humans. The leafhoppers and planthoppers are second only to aphids as important plant disease vectors. Leafhoppers are extremely diverse in North America, with over 2500 species known from the United States and Canada. Planthoppers, on the other hand, are few in number, with only about 145 species known from the same geographic area.

Whiteflies are small but distinctive insects. They rarely exceed 2–3 mm in length, and usually are entirely white in color. Veins are not apparent on the wings because they are covered with white scales. The antennae are usually fairly long, thread-like, and consist of seven segments. Whiteflies are tropical species, and most abundant in warm climates and in greenhouses. Because they reproduce rapidly and attain very high densities, and can transmit plant viruses, they can be quite damaging to vegetable crops. The nymphal stage is flattened, often not really resembling an insect, and

Pseudococcidae—Mealybugs Mealybugs are oval, flattened insects that secrete waxy filamentous material over their body, sometimes making their identity difficult to determine. Many species also produce filaments around the periphery of the body, including particularly long filaments in the anal region that resemble “tails.” About 280 species are known from the United States and Canada, but are usually known as pests of weedy plants and ornamentals in greenhouses. Only pink hibiscus mealy-

12  Handbook of Vegetable Pests

bug, Maconnellicoccus hirsutus (Green), is known to cause serious injury to vegetable crops in the field. Psyllidae—Psyllids Psyllids are small insects, measuring 2–5 mm in length. They resemble aphids superficially, and cicadas upon close examination. In adults the antennae are long, consisting of 9–11 segments. The hind legs are stout and capable of producing long jumps. Their leaping ability is an important diagnostic character. The body of nymphs is flattened, with a short fringe of filaments along the periphery. This stage is sedentary. In the United States and Canada, about 260 species are known to occur. Few species are important to crops.

Order Hymenoptera—Ants, Bees, Sawflies, and Wasps The order Hymenoptera is diverse in appearance and biology. Hymenopterans bear two pairs of membranous wings. Most have chewing mouthparts. Often a significant constriction occurs between the thorax and the abdomen, producing a thread-like “waist.” A distinct ovipositor is present in many females. The order Hymenoptera contains many social insects such as ants and bees, which live in colonies, and also many solitary species. Some hymenopterans such as cicada killer wasps attain a large size, sometimes over 100 mm. In contrast, hymenopterans such as egg and thrips parasitoids are among the smallest insects, measuring about 0.2 mm in length. Most members of this order are not plant pests, rather serving to pollinate plants, scavenge detritus, recycle nutrients, or parasitize other insects. Note that although only a single species of ant is treated as a plant pest in this book, many other species are indirectly detrimental because they tend aphids, affecting their location and abundance, and thus indirectly influence the amount of damage to crops. The order Hymenoptera has complete metamorphosis, displaying egg, larval, pupal, and adult stages. About 18,000 species are known from the United States and Canada, but some authorities believe that many species await discovery.

Family Argidae—Argid Sawflies There are several families of sawflies, but most sawflies feed only on trees and shrubs. Only one species, in the family Argidae, attacks vegetable crops, and it is limited to feeding on sweet potato. The adults are broad-bodied, lacking the constriction or narrowing of the abdomen found in most Hymenoptera. Larvae of sawflies are elongatecylindrical in form and resemble caterpillars, but bear six or more pairs of prolegs along their abdomen, more than is found on caterpillars. The prolegs of sawflies also lack the terminal hooks or crochets that occur on caterpillar prolegs. There are only about 60 species of argid sawflies in the

United States and Canada, though the number of sawflies in the several sawfly families totals nearly 1000 species in this geographic area. Adult argid sawflies measure 8–15 mm in length and bear antennae with only three segments; the third segment is disproportionately long.

Family Formicidae—Ants Ants are social insects, generally living in large colonies. They also display division of labor, which is reflected in the appearance of different “castes” within a colony which differ in their appearance and behavior. Thus, we find some ants specialized for reproduction, but others for food collection (workers) or colony defense (soldiers). Ants measure 1–20 mm in length. The anterior portion of their abdomen is markedly constricted, but the first one to two segments bear bumps. The antennae are elbowed, an important diagnostic feature, and consist of 6–13 segments. Few ants can be considered to be direct pests of vegetables, and only red imported fire ant, Solenopsis invicta Buren, is regularly damaging. However, because ants attend aphids, they influence the abundance of these other plant pests, and exacerbate damage. This is somewhat offset by the predatory behavior of ants, as they can be voracious predators of caterpillars and other insect pests.

Order Lepidoptera—Moths, Butterflies, and Caterpillars This large group contains many destructive pests. The most important diagnostic character is the presence of flattened scales on the wings and often other parts of the adult’s body. Adult lepidopterans have four wings, are often rather heavy bodied, and bear medium-sized to long antennae. The antennae vary in appearance; in butterflies the tip is usually expanded to form a knob, whereas in moths the apical expansion is lacking. Adults also have long, tubular mouthparts that uncoil to collect nectar from flowers. This structure is not capable of piercing plant tissue. Lepidopterans undergo complete metamorphosis, possessing egg, larval, pupal, and adult stages. The immature stage, or caterpillar, is cylindrical and bears three pairs of thoracic legs and two to five pairs of abdominal prolegs. The prolegs bear small hooks, called crochets, at their tips. The head capsule is well developed and hardened. The head bears chewing mouthparts and small, obscure antennae. The caterpillar may be naked or covered with short or long hairs. Larvae usually feed on plant foliage, but sometimes on other plant parts. About 12,000 species are known from the United States and Europe.

Family Erebidae—Tiger Moths, Woolly Caterpillars, and Others Erebids are heavy-bodied, medium-sized moths, usually measuring 25–30 mm in wingspan. They are often strikingly

Introduction Chapter | 1  13

marked. The forewings are unusually long and pointed. The caterpillars bear thick and long coats of hairs, giving rise to the “woolly” designation. Few species are vegetable pests, and those that are tend to be associated primarily with weeds. There are about 265 species of erebids in the United States and Canada.

Family Gelechiidae—Gelechid Moths and Leafminers These moths are small, measuring only 8–25 mm in wingspan. The wings usually bear a fringe of long hairs. The larvae of vegetable-infesting species are primarily leafminers, though potato tuberworm larvae also enter potato tubers. Approximately 630 species of gelechids are known from the United States and Canada.

Family Hesperiidae—Skipper Butterflies Skippers are not usually considered to be plant pests, and only one species affects vegetable crops, bean leafroller. Skippers are small to moderately sized, heavy-bodied insects. Many species, including bean leafroller, Urbanus proteus (Linnaeus), have elongate extensions or “tails” on their hind wings. They tend to be brownish in color, with antennae bearing thickened and hooked tips. Larvae generally feed within rolled leaves. Larvae have an enlarged head, and the thorax immediately behind the head is constricted. About 300 species of hesperiids are known from the United States and Canada.

Family Lycaenidae—Hairstreak, Copper, and Blue Butterflies These small brightly colored butterflies are not often thought of as pests. However, gray hairstreak, Strymon ­melinus (Hübner), feeds on legumes. The hairstreaks generally bear 2–3 narrow extensions or “tails” trailing from the back of the hind wing. The larvae generally are short and broad, with the head largely hidden by the thorax. About 140 members of this family occur in the United States and Canada.

Family Noctuidae—Noctuid Moths, Armyworms, Cutworms, Loopers, and Stalk Borers This is one of the largest and most important groups of vegetable pests. The moths are mostly nocturnal and obscurely marked. They are moderate in size, measuring 20–45 mm in wingspan, and heavy bodied. The front wings are rather narrow, the hind wings broad. The hind wings are usually paler in color than the forewings. The larvae are variable in appearance, but generally dull colored and lacking long hairs or stout bristles. Many larvae are nocturnal, seeking shelter during the day in soil and beneath debris. Cutworm and armyworm larvae normally bear

five pairs of abdominal prolegs. Armyworms are so named because of their tendency to migrate in groups, but this is a transient property that is usually associated with high densities, and even some insects called cutworms take on such migratory tendencies at times. Looper larvae bear only three to four pairs of abdominal prolegs, allowing them to arch their back as they crawl. Most larvae are leaf feeders or stem borers. Nearly 3000 species of noctuids occur in the United States and Canada.

Family Papilionidae—Swallowtail Butterflies and Celeryworms Swallowtail butterflies are large insects, often 60–100 mm in wingspan. They usually are brightly colored, and have elongate extensions or “tails” trailing from the hind wings. Few attack vegetables, with only umbelliferous crops being damaged. The larvae are large, heavy bodied, and colorful. When disturbed, larvae evert colorful, scented glands from behind the head. Only 35 species of swallowtails occur in the United States and Canada.

Family Pieridae—White, Sulfur and Orange Butterflies, and Cabbageworms Pierid butterflies are among the most common butterfly species in gardens, meadows, and roadsides. They are medium in size, usually 40–60 mm in wingspan. The white or yellowish color of their wings is normally marked with black spots or borders. The larvae of several species affect crucifers and are often called “cabbageworms.” About 60 species inhabit the United States and Canada.

Family Crambidae—Snout Moths, Borers, Budworms, Leaftiers, and Webworms This family is second only to Noctuidae in importance among the Lepidoptera. The adults are small to medium in size, with a wingspan of 10–40 mm. The head appears to protrude forward, providing the basis for the “snout moth” designation, though it is enlarged mouthparts (labial palps) that account for the extension. Because of their manner of holding the wings when at rest, pyralid moths generally have a distinctive triangular appearance. The feeding behavior of larvae is quite diverse, though stem borers, leaf tiers, and webworms predominate. The larvae are not easily distinguished from noctuid larvae and some other caterpillars, but many pyralids have a light spot or a ring on the body at the base of body hairs. Nearly 1400 species of Pyralidae are known from the United States and Canada.

Family Sphingidae—Hawk Moths and Hornworms Hawk moths are among the largest moths, with a wingspan sometimes exceeding 160 mm. The front wings are ­longer

14  Handbook of Vegetable Pests

than the hind wings, sometimes twice as long. Hawk moths are strong fliers, often hovering like hummingbirds and sometimes mistaken for these small birds. Though sometimes active during daylight, hawk moths are most commonly observed near dusk, usually hovering at flowers or darting from flower to flower. The larvae are large and heavy bodied, and usually bear a single prominent tapered appendage or “horn” at the posterior tip. This horn is the basis for the common name of larvae, and is incorrectly thought by some to be harmful to humans. Hornworms feed on a variety of plants, but only a few injure vegetables. Approximately 125 species of hawk moth are known from the United States and Canada.

antennae tend to be about one-half the length of the body or less, which serves to differentiate this group from the Tettigoniidae. Females lack the large ovipositors found in many other groups of orthopterans. The Acrididae is the largest and most destructive group of orthopterans, though they typically reach their greatest abundance in arid grassland environments where vegetables are not grown. They are quite dispersive, however, with some species migrating long distances and causing great crop destruction. Many species are indiscriminate feeders, accepting a broad range of host plants. About 550 species live in the United States and Canada.

Family Tortricidae—Leafrollers and Borers

Family Gryllidae—Field Crickets

Tortricid moths are generally small, but range from 10 to 35 mm in wingspan. They are obscure moths, usually mottled brown or gray. The larvae often roll leaves or bore into plant tissue, and many are important plant pests. Over 1000 species of tortricids are known from the United States and Canada. However, among vegetable crops, only pea is affected by a tortricid; pea moth, Cydia nigricana (Fabricius), burrows into pea pods and feeds on seeds.

Crickets are heavier bodied than grasshoppers, and have longer antennae. The ovipositor of females is conspicuous. Sound production is an important aspect of cricket biology. Among the many types of crickets, only field crickets are crop pests, and then only infrequently. About 40 species of field crickets are known from the United States and Canada.

Order Orthoptera—Grasshoppers, Katydids, and Crickets The order Orthoptera consists principally of grasshoppers, crickets, and katydids. Most are medium (20–50-mm body length) but some are large (up to 80 mm) in size. They tend to be relatively thin and elongate in body form, and possess long legs, particularly the hind legs. Most species have two pairs of wings, though some have abbreviated wings and a few are wingless. The front wings are narrow and slightly thickened, but not very hard. They serve mostly to protect the hind wings from damage rather than for flight. The hind wings are large but thin and membranous. The hind legs are enlarged and can be used for leaping, though leaping is a defense reaction and grasshoppers normally move about by walking. The mouthparts of grasshoppers are the chewing type. The antennae are variable in length, often quite long, and usually thread-like. Grasshoppers undergo incomplete metamorphosis, with egg, nymphal and adult stages, but not a pupal stage. The immatures, or nymphs, greatly resemble the adults, with nymphs differing primarily due to small body size and poorly developed wings. Nymphs occur in the same habitat as adults and, like adults, feed principally on plant foliage. Some orthopterans produce sound. About 1000 species of Orthoptera are known from the United States and Canada.

Family Acrididae—Grasshoppers This family of Orthoptera is also known as “short-horned grasshoppers” due to the length of the antennae. Acridid

Family Gryllotalpidae—Mole Crickets Mole crickets are among the most distinctive orthopterans. These medium-sized insects, about 20–40 mm in length, have front legs that are modified for digging through soil. Their antennae are relatively short. The female’s ovipositor is not pronounced. As pests, they are limited to the southern states. Only seven species are known from the United States and Canada.

Family Tettigoniidae—Shieldbacked Crickets and Katydids This family is best known from the large green leaf mimics known as katydids. Katydids are not vegetable pests, but a few related insects known as shieldbacked crickets can cause damage. Within the Rocky Mountain region, some species can be very destructive. The large (40–60 mm), humpbacked, short-winged, shieldbacked crickets resemble field crickets more than katydids, though they can be green in color. The antennae of tettigoniids are longer than the body. The female’s ovipositor is large. Though flightless, shieldbacked crickets are quite dispersive and may migrate into crop-growing areas. About 250 species of tettigoniids occur in the United States and Canada.

Order Thysanoptera—Thrips These very small insects usually measure only 1–2 mm in length. They have two pairs of wings, with both pairs consisting largely of fringe hairs. The antennae are medium in length and thread-like. The mouthparts pierce plant tissue and remove plant sap, though the mouthparts are often described as rasping rather than piercing-sucking. These i­ nsects

Introduction Chapter | 1  15

e­ xhibit a form of incomplete development in which the first two instars feed actively but the final two instars are inactive. Tarsal claws usually are not present. The nymphal instars generally resemble the adults in body form, the principal exceptions being the smaller size and undeveloped wings of nymphs. Thrips can be found on foliage and in blossoms. Despite their small size they can be quite destructive due to their ability to transmit plant viruses. About 700 thrips species inhabit the United States and Canada.

Noninsect Invertebrate Pests Other classes of animals resemble insects, or inflict similar damage to vegetable crops. There are four basic groups of such pests: 1. Class Acari (phylum Arthropoda)—mites. These animals are smaller than most insects, and usually bear eight legs. 2. Class Collembola—springtails, and class Symphyla— symphylans (both phylum Arthropoda). These small soil-dwelling animals have been considered to be insects in the past and share many morphological characteristics with insects. 3. Class Isopoda—pillbugs and sowbugs, and class Diplopoda—millipedes (both phylum Arthropoda). These many-legged animals can be distinguished by their many-segmented bodies and numerous legs. 4. Class Gastropoda (phylum Mollusca)—slugs and snails. These slimy animals may or may not bear shells, but the lack of body segmentation and legs provides for easy recognition.

PEST MANAGEMENT PHILOSOPHY AND PRACTICES What Is a Pest? A pest is not a biological phenomenon, it is an anthropomorphic designation. For example, we consider termites to be beneficial organisms when they live in forests, converting dead trees into soil organic matter. The same insects are pests when they feed on wood associated with human structures. In both cases, the termite behavior is the same, but in one case we place no value on the wood being consumed, and in the other we assign high economic value. Sometimes the mere presence of insects causes concern or alarm. When simple insect occurrence or very minor feeding is the basis for designating an insect as a pest, the insect is said to be an esthetic or cosmetic pest. However, when insects decrease the value of a commodity they are said to be economic pests. There is no absolute distinction between

esthetic and economic injury, especially with respect to vegetable crops. A home gardener may consider a dimple on a tomato fruit caused by the feeding of a stink bug to be an insignificant blemish, an esthetic injury, but the same type of blemish can cause a produce buyer to downgrade the value of a crop, causing significant economic loss to a tomato farmer. Similarly, a few holes in the leaves of cabbage or lettuce plants caused by flea beetles is of no significance early in the life of a crop because the affected foliage is not harvested. The same type of injury, should it be more frequent or appear late in the development of the crop, could constitute injury either by reducing the growth rate of the plant or by affecting the appearance of the harvested commodity. It is very useful to differentiate between direct and indirect pests. Direct pests are those that attack the portion of the vegetable plant that is harvested for food. Indirect pests attack some other portion of the plant, not the harvested portion. A stink bug feeding on a tomato fruit is a direct pest; a leaf miner burrowing in tomato foliage and a wireworm feeding on tomato roots are indirect pests. Ultimately, the presence of large numbers of indirect pests can be damaging, but on a per individual insect basis indirect pests are less injurious than direct pests. Knowing the identity of pests, and their feeding behavior, allows us to distinguish between a severe and modest threat to a vegetable crop, and determines our course of action in dealing with the threat. In commercial vegetable production, economic considerations are very important. Commercial vegetable producers normally are aware of the “economic injury level,” the point at which pest suppression is economically feasible. There is little value to spending more money on production costs such as pest suppression than can recovered in increased yield. Therefore, a grower may not be very concerned about indirect pests, relative to direct pests. Similarly, growers may be unconcerned about direct pests that are few in number unless they burrow into the produce and are not easily detectable; such cryptic damage can dramatically lower the value of a crop because consumers are intolerant of even slight insect contamination of food products. Commercial growers often make decisions that affect large acreages, and have a huge financial investment at stake. Under such circumstances, it is commonplace to assume a worst-case scenario, and to act decisively to protect crops from insect damage. This approach is viewed as a form of crop insurance, and in many cases is a good investment. However, in many cases good crop monitoring can establish precisely how abundant and damaging insects might be, and because they often are not numerous enough to cause injury, significant financial savings can be realized by treating crops for pests only when treatment is actually needed. It is important to bear in mind that unnecessary treatments for pests also are a financial loss. Such unnecessary treatments also encourage the development of insecticide resistance in pest populations. In the home garden environment there is greater latitude for risk, and

16  Handbook of Vegetable Pests

it is often possible to abstain from pest suppression actions unless pests are observed. In either the commercial or home garden situation, we can make sound decisions about the need for pest control only when we have information about pest identity, biology, and damage potential.

How Damaging Are Pests? Despite our best efforts to prevent injury, vegetable crops do sustain damage by insects and other pests. The level of loss varies among crops, locations, and years. Also, some pests are consistently injurious, others rarely so. Crop losses were estimated by Metcalf and Metcalf (1993), using various sources, to average about 12.5% for commercially grown vegetable crops in the United States. Undoubtedly, entomologists could quibble about the validity of any estimate, particularly because losses vary so much from region to region and from year to year, but the loss clearly is significant. The estimated crop losses and the economic value of the insect damage (based on figures from the late 1980s) are illustrated in Table 1.6.

TABLE 1.6  Estimated crop losses in United States. Crop

% loss

Asparagus

15

22.8

Beans, snap

12

11.8

Broccoli

17

49.7

Cabbage

17

3.8

2

6.9

Cauliflower

17

34.4

Celery

14

32.3

Cucumber

20

26.0

Lettuce

7

73.1

Melons

10

9.7

Onion

18

76.7

Pea

10

7.3

Potato

12

230.2

Sweet corn

14

50.5

Carrots

$ loss (millions)

Not surprisingly, the cost of preventing injury varies considerably among locations and crops, and from year to year. The cost of insecticide use in the central Florida tomato production region during 1995–96 was high, $380/ acre ($950/ha) in the spring production period and $488/ acre ($1220/ha) in the autumn period. This does not include application costs because machinery costs are often spread over several practices; for example, fungicides and insecticides are often applied simultaneously. Similarly, fumigation practices are directed primarily to disease and nematode management, though weed and insect suppression also accrues. Crop scouting costs cannot be assigned to any particular pest group, and are shown separately. Note that insect control costs are appreciably can be higher in the autumn period. This is due to better survival of pests during the summer intercrop period than the winter intercrop period, resulting in greater abundance and greater risk of damage. Also noteworthy is that even in the relatively insecticide-intensive Florida cropping system, insecticide costs represent no more than 13.8% of the operating costs, 8.3% of total preharvest costs, and 4.7% of total tomato production costs. Farmers tend to focus their cost-cutting efforts on the higher priced elements of the crop production system, often labor and packing operations. The prevalent attitude among most farmers, as long as insecticides are available, effective, and affordable, is that their use will minimize an element of risk at relatively low cost. However, if all the pest-related costs are aggregated, the total cost is greatly enhanced, and the true cost of “crop insurance” can be better appreciated. In the Florida tomato production example cited above, the impact of pest management practices is estimated to cost about 49% of operating costs, 29% of total preharvest costs, and 17% of total costs. Insecticide use is expensive in many crops other than tomatoes. For the period 1995–96, insecticide costs on some other vegetable crops grown in Florida are presented in Table 1.7.

TABLE 1.7  Estimated crop loses for vegetable production in Florida, United States. Crop

Region of Florida

$/acre

$/hectare

Sweet potato

5

8.0

Tomato

7

98.5

Beans, snap

South

108

270

15

11.3

Cabbage

North

113

282

Celery

South

352

880

Central

201

502

South

211

527

Central

122

305

Watermelon

These losses do not include the cost of preventing insects from causing even greater injury. The financial cost of managing vegetable pests is difficult to estimate, but the value of insecticides alone is measured in hundreds of millions of dollars.

Corn, sweet

Continued

Introduction Chapter | 1  17

TABLE 1.7  Estimated crop loses for vegetable production in Florida, United States—cont’d Crop

Region of Florida

$/acre

Cucumber

South

172

430

Eggplant

Central

457

1142

Pepper

South

427

1067

Central

387

1769

Central

119

297

North

24

60

Squash

South

67

167

Watermelon

South

131

327

Central

48

120

North

9

23

Potato

$/hectare

As is evident from this example, insecticide costs tend to be higher in Southern Florida than in Central and Northern Florida (Smith and Taylor, 1996). This is due to the better survival and greater abundance of insects in the subtropical regions of Southern Florida. Such variation in insect abundance, and costs associated with insect suppression, are not restricted to Florida, nor are they due only to weather-­related factors. The presence of certain prevailing wind patterns, culture of alternate crop hosts, and the abundance of weeds or plant disease also influence the severity of pest damage. Sometimes insect problems are so severe or expensive that farmers abandon culture of certain crops in a particular area. Home gardeners similarly avoid culture of certain crops if they regularly encounter difficulty. Both farmers and home gardeners sometimes lack the sophistication, motivation, and economic resources to manage pests with the latest, most effective technology. On the other hand, if the garden plot is sufficiently small and the gardener sufficiently motivated, extraordinary efforts are sometimes applied. The culture of vegetables under row cover material or screening is a good example of strong desire to overcome pest pressure, or sometimes a strong desire to avoid insecticide use.

The Types of Insect Injury Insects cause several different types of injury to vegetables, with some species capable of causing more than one type of damage. Recognition of the nature of injury is often an important diagnostic feature in the process of determining pest identity. Because the mouthparts of insects are principally useful for chewing, or for piercing and sucking liquid plant contents, the principal forms of injury reflect these major behaviors. The most common form of injury occurs when insects live externally on a plant, biting off portions of tissue and

swallowing the small pieces. Such insects, known generally as chewing insects, are usually foliage feeders or root feeders. There are numerous variations in behavior among chewing insects. Some insects, particularly at young stages, feed only on the leaf surface or avoid consumption of leaf veins. This selective feeding results in the destruction of leaves, though the veins remain and sometimes the outline of the leaf is still quite apparent. This damage is called ­skeletonizing because the “skeleton,” or veins, are left intact. Leaf rollers or leaftiers web together leaves to provide shelter for the insect. They may remain within the shelter to feed, or venture forth only to return to the shelter when not actively feeding. Leaves may also provide shelter for leaf miners, which feed on the internal tissues of leaves while leaving the outermost layers intact. Leaf miners normally spend their entire larval life within a single leaf, leaving only when it is time to pupate. Some chewing insects are stem borers, chewing into leaf petioles, stalks, or roots. In many respects, stem borers are similar to leaf miners, differing mostly in the site of feeding. However, they sometimes move considerable distances within the stem, and tend to be more damaging than leaf miners. By burrowing through root, stalk, and petiole tissues, borers disrupt the transport of water and nutrients for large sections of the plant, or even the entire plant. This not only inhibits normal growth, but sometimes leads to complete loss in productivity or death of the plant. Perhaps the most damaging of the chewing insect pests are blossom or fruit feeders, insects that feed on or within the reproductive tissues of the plant. The fruit is often the most valuable portion of the plant, and even a small amount of feeding can cause rejection for cosmetic (esthetic) reasons, or can lead to invasion by plant pathogens. In addition to the direct injury caused by chewing mouthparts, chewing insects sometimes cause damage indirectly. Chewing insects sometimes vector plant diseases, though they are much less efficient than piercing-sucking insects, or open a potential site of infection for plant pathogens borne in the soil, water, air, or insect fecal material. Many insects have long tubular mouthparts that allow them to pierce the conducting elements of plants, or individual cells, and to extract the liquid contents. These insects, called piercing-sucking insects, may be found on leaves, stems, or roots, but most often are associated with the terminal or youngest leaf and stem tissue. Feeding by piercingsucking insects often results in discoloration and deformity, apparently because insects not only extract liquids but secrete digestive enzymes and toxins while feeding. A common response is chlorosis, or yellowing of leaf tissue; this may occur generally, locally, or even in individual cells. Premature leaf drop sometimes occurs following feeding by piercing-sucking insects. Leaf curling or cupping is a common form of deformity, usually associated with feeding by aphids. Galls, or swellings caused by an increase in plant cell size or number, often envelop feeding insects and

18  Handbook of Vegetable Pests

provide them with both protection and enhanced nutrition. Piercing-sucking insects often cause stunting, or decrease in stem and leaf elongation. Reduced stem elongation but normal lateral growth results in dense, bushy vegetation called witch’s broom. Perhaps the most important form of injury caused by piercing-sucking insects is pathogen (disease) transmission. Piercing-sucking insects are particularly adept at transmitting diseases caused by viruses and similar organisms.

Pest Management Philosophy There are several ways to approach the management of vegetable pests. In some instances, profit is the major concern, though in other cases maximum yield or pesticide-free vegetables are desired. In some cropping systems, or for some particular pests, effort is made to prevent pest populations from establishing by preventative application of insecticide. On the other hand, often crops are monitored and not sprayed until the first appearance of a particular pest. In still other situations, small or moderate numbers of pests are tolerated but the density is carefully monitored and suppression initiated when densities reach some threshold of abundance. The philosophy behind commercial growers’ pest management practices is quite variable, despite the common objective of making a profit. Part of the difference in approach to pest management stems from the uncertainty of production and profit. When a particular vegetable is in relatively short supply the price is high and very good profits are made. Under such circumstances, the cost of pest management practices seem insignificant and maximization of yield pays handsome dividends. On the other hand, when a vegetable is in good supply relative to demand, the price received by the grower is low, perhaps barely covering costs of production. Under these circumstances, farmers who cut costs where they can, such as unnecessary insecticide use, make greater profit. Thus, a farmer’s decision to spend money on pest management practices is tempered by the future outlook of supply and profit. One thing is certain, however; any vegetable crop must be high in quality and free of blemishes if it is to be marketed at a profit. Thus, it is not surprising that producers invest heavily in insecticides. Home gardeners are less sensitive to appearance of the produce. They are often motivated as much by the quality of the produce, which often means freshness and absence of pesticide residues, as they are by the possibility of saving money by growing their own vegetables. Because profit is not a key element, home gardeners can engage in practices that would be cost-prohibitive on a commercial scale. Home gardeners also tend to have less ready access to highly effective but very toxic pesticides. Commercial producers of organically grown vegetables are faced with the difficult

challenge of producing blemish-free produce with a very limited array of “organic” insecticides. It is always desirable to prevent pest populations from developing because this eliminates the need for corrective action and the risk of crop loss. Certain actions can be taken, such as growing a crop in an isolated location, cultivating a pest-resistant cultivar, or modifying the date of planting to avoid the major flight period of a pest. Such practices often virtually eliminate problems with certain pests. Unfortunately, there are many potential pests for any vegetable crop, and such extraordinary efforts can only be directed to one or a few pests per crop. Therefore, such manipulations are usually directed only at the most severe, regularly occurring pests that must be eliminated if a crop is to be successful. That leaves other pests to be managed in a curative or corrective mode when, and if, they appear. Insecticides are very useful for this purpose, although they are not necessarily the only option. Residual, broad-spectrum insecticides, if used f­ requently, can prevent invasive insects from establishing, as well as eliminate most pest problems that do exist. By using such materials on a regular basis, growers can avoid concern about regularly occurring pests as well as sporadic pests. Historically, such broad-spectrum insecticides were readily available, economic, unfailingly effective, and perceived to be relatively safe for people and the natural environment. Although conditions and perceptions relative to the characteristics of insecticides have changed, it can be easily seen how the use of insecticides would have great appeal to producers, and how insecticide use would become an integral part of commercial vegetable production. Increasingly, growers cannot depend on the availability of broad-spectrum insecticides to rid their fields of pest problems. There are three primary constraints on use of insecticides: cost, effectiveness, and availability. The cost of insecticides and their application can be quite high. Insecticides are applied to some crops at 2–3-day intervals during periods of growth when the crop is particularly vulnerable to damage. This can result in considerable cost to the producer. If material and application costs increase faster than the revenue farmers receive for their produce, it is no longer costeffective to grow the crop. Also, insecticides are not always effective. Some vegetable crops are attacked by insects that are inherently difficult to control. It is also possible to exacerbate the problem caused by certain pests such as mites, leaf miners, and whiteflies by excessive use of insecticides. The principal causes for increased problems with pests following insecticide application are insensitivity of the target pest to the insecticide (i.e., resistance), and destruction of potentially effective natural enemies by the insecticide. Lastly, availability of pesticides is decreasing. The high cost of developing new products discourages insecticide manufacturers from developing new products. Concern over the health of farm workers, consumers, and

Introduction Chapter | 1  19

nontarget wildlife such as birds and fish also has lead to regulatory constraints on the availability and use of insecticides. Some insecticides are quite hazardous to fish and birds, but others have few deleterious effects, disrupting the metabolism or behavior of insects but not other animals. The use of more selective pesticides, or the use of pesticides more selectively, is increasingly encouraged. Selectivity implies targeting only pests that need to be eliminated, and use of insecticides only when they are truly needed. To accomplish the objective of shifting from broadspectrum to selective pest suppression materials, it is imperative that the identity, biology, and damage potential of pests be known. Vegetable producers are also encouraged to use alternatives to pesticides, or a combination of tactics, rather than depending solely on insecticides for their pest suppression needs. They attempt to integrate pest management practices such as using cultural practices that limit pest potential, with careful monitoring of pest abundance, and application of selective insecticides only when necessary.

Pest Management Practices There are numerous types of practices that can be implemented to protect a crop from pests, or to eliminate pests that have invaded. These practices can be organized into four broad categories: biological control, cultural manipulations, physical manipulations, and insecticides. They differ greatly in effectiveness, ease of implementation, cost of implementation, time interval required before becoming effective, and reliability. No single practice is effective for all pest problems, though insecticides come closest to being universally suitable. Unfortunately, the benefits of insecticide use are sometimes offset by health or environmental hazards.

Biological Control The use of natural enemies to suppress pest insects is one of the oldest and most effective approaches known. Although using natural organisms to our advantage is highly desirable, it is often difficult to implement and difficult to predict the outcome. Biological control tends to work best when the pest species is an invader from another region, and is maintained at a low or moderate level of abundance in its native land by natural enemies. In this case, we import the native enemies and then culture and release them where they are needed. Once established, if the native enemies are well adapted to the new environment they provide permanent, no-cost suppression. This approach is called classical biological control. However, sometimes natural enemies cannot persist at a high enough level in the new environment to provide effective suppression, or effective enemies cannot be located. In this case, we must culture and release natural enemies that have some promise of providing s­ uppression

in large enough numbers to overwhelm the biotic potential of the pest, and drive their population down. This often requires regular, timely release of natural enemies, and is called augmentive biological control because we are augmenting the natural population of natural enemies with supplemental biological control agents. The final major approach to using biological control organisms is to modify the environment or otherwise preserve and favor existing natural enemies. This is called conservation biological control, and usually involves preserving some habitat or food resource, including alternate host insects, or protecting the beneficial insects from the deleterious effects of pesticides. The conservation approach is especially appealing when the insect pest is a native species and no source of exotic natural enemies is apparent, or when the economics of producing the commodity do not favor mass culture and release of natural enemies—often an expensive undertaking.

Cultural Manipulations Modification of planting, crop maintenance, and harvesting practices can sometimes affect pests. For example, a common practice is to delay planting so as not to have the plant present when the insect pest becomes active, or to plant early so the crop plants are well established and able to withstand some root feeding or defoliation and remain healthy. This is most effective when insects occur synchronously and for a relatively brief period of time, and when working with a crop that will not suffer from such manipulations. Similarly, tillage of the soil and destruction of crop residue often reduces the overwintering survival of insects. Other cultural practices that often are manipulated in an effort to manage pests are crop irrigation, rotation between dissimilar crops, crop isolation from sources of pests, fallow or crop-free periods, and cultivation of crop varieties that are less attractive or suitable for pests. Cultural manipulations are often the most cost-effective approach to pest management, but it is difficult to manage an entire complex of pests using these techniques. Resistance of host plants to injury by key pests is increasingly an important mechanism of cultural manipulations. The inclusion of resistant genomes increasingly provides plants with full or partial resistance, though the resistance may be limited to only some of the key pests. Normally, suppression of injury is limited to Lepidoptera, although in some cases other taxa may be affected.

Physical Manipulations Physical manipulations sometimes are useful, especially for small-scale or home garden vegetable production. For example, metal and tar paper barriers around the base of seedlings can provide protection against the feeding of cutworms and oviposition by root maggots, respectively. Also, reflective or colored mulch sometimes will repel

20  Handbook of Vegetable Pests

certain flying insects, including vectors of important plant viruses. Screen and floating row covers often can be used to protect insects from reaching susceptible crops, though allowance may need to be made to provide entry by pollinators. Occasionally, traps can be used for collection and ­destruction of pests. Traps can be baited with food-based or chemical lures, they may be attractive due to an insect’s natural orientation to a particular color, or can take advantage of the pest’s tendency to seek a certain type of shelter. Some large insects can be observed readily and simply collected by hand, not necessitating any type of attractive device. Physical manipulations tend to be material intensive, and sometimes labor intensive, so their use is most common in high value crops or in small plots.

Insecticides The most common approach to vegetable crop protection is to use insecticides. Most insecticides are derived from synthetic organic chemicals, though some are derived from naturally occurring minerals or plants. Insecticides disrupt the physiology of the insect. Most of the currently available synthetic organic insecticides, particularly the broad-spectrum insecticides, disrupt the nervous system of insects. Some are quite specific to insects, but many are biocides, general poisons that can affect fish, birds, and mammals if they are exposed to sufficiently high levels. The botanical insecticides are favored by organic gardeners because they are perceived to be “natural.” Botanicals degrade quickly in the environment, but some are quite toxic to humans and should be handled as carefully as synthetic insecticides. A few products are derived from, or consist of, insect-­pathogenic microorganisms. These microbial insecticides tend to be very specific, and safe to most nontarget organisms. The best example of this is the bacterium Bacillus thuringiensis. Increasing in popularity is the use of soaps (detergents) and oils (both mineral and vegetable). Though their effectiveness is generally limited to small organisms, they pose few hazards to humans and other animals. Insecticides are formulated in many different ways. This flexibility as well as long shelf life help account for the popularity of insecticides. Most insecticides are mixed with water and applied as a spray, but sometimes dust or granule formulations are most efficient, or aerosol applicators are easier to use. Least frequent of the application methods is bait formulations, wherein the toxicant is mixed with an attractive, edible material.

Species Identification General Considerations When an unknown pest is found on vegetables, it is important to reduce the number of potential identities to only a few by assigning the unknown to a small group, usually an

insect order and family. Each order and family contains only a fraction of the approximately 300–400 species that are capable of causing damage to vegetables. You can greatly simplify identification by knowing the order, and perhaps the family. Usually only a few characteristics such as wing number and leg or mouthpart type are needed to assign an unknown pest to order and family. The “quick guide” in this section and a key in Appendix A will aid you in this process.

Why Identification is so Important There is a great wealth of information about vegetable pests. Hundreds of careers have been devoted to determining which species are damaging, the nature of their damage, the life cycle of the pest, and how the pest is best managed to eliminate or minimize injury to plants. It is easy to overestimate the damage potential of some large, leaffeeding pests, and similarly easy to underestimate the damage caused by small, sap-sucking species. Failure to recognize the presence of certain pests such as plant virus vectors early in the season can prove to be a grievous error, resulting in severe crop injury. On the other hand, sometimes vegetable plants are treated with an insecticide because someone has observed lady beetles or another beneficial insect, and mistaken them for pests. Therefore, the collective knowledge and experience of generations of entomologists are available once you identify the pest—but are useless unless correct identification is made of the pest at hand.

Approaches to Identification There are three guiding principles to identification that, if followed, enhance correct identification. Remember that some species can be identified only by authorities, and some cannot be distinguished based only on appearance, but most serious pests are readily distinguished with but little effort. You may need to use some low-level magnification such as a 10 × hand lens, and such equipment is inexpensive and easy to use. 1. Although there are numerous pests, most are easily assigned to groups based on easy-to-discern characters such as the number of legs or wings, and the type of mouthparts. With little effort it is easy to distinguish such common groups as aphids, caterpillars, whiteflies, and wireworms. 2. Behavior is as important as appearance in distinguishing among pests. How or where a pest feeds is critical knowledge. The presence of silk webbing, the positioning of eggs, and other aspects of pest biology often are key elements in distinguishing among pests. When attempting to identify an unknown pest, try to observe as much as possible about pest behavior and plant damage.

Introduction Chapter | 1  21

3. Details are important; look carefully at the pest and its damage, and do not trust your memory. Collect the pest you want to identify, do not just kill it. No one who is unfamiliar with the pest’s identity can anticipate what characters will be critical in distinguishing it from its relatives. It is entirely too easy not to notice the number of wings or the color of the legs, or not to remember to count the stripes or spots. Compare the written descriptions with the physical evidence carefully. Do not ignore aspects of the description because you are unfamiliar with the terminology. Entomologists, like members of all scientific disciplines, use technical words to describe the physical appearance and behavior of their subjects of study. In this book, the use of terminology is minimized, and terms are described in the glossary and illustrated with diagrams.

If a pest poses a major economic threat to a crop and you are uncertain of its identity, there are several sources of assistance available. The principal source in the United States is the Cooperative Extension System associated with the Land-grant University found in each state. A cooperative extension service office is normally found in each county, and the employees are commonly called county agents. The county agents often can assist with identification, and if they cannot they will forward the pest on to the state university for identification. Research centers and experiment stations are other good sources of expertise, both in Canada and the United States. Crop consultants provide identification and pest management recommendations on a for-fee basis, and usually can be found in important agricultural areas. You can access considerable additional information to aid in identification if you have access to the internet.

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Chapter 2

Pest Identification

GENERAL CONSIDERATIONS

APPROACHES TO IDENTIFICATION

When an unknown pest is found on vegetables, it is important to reduce the number of potential identities to only a few by assigning the unknown to a small group, usually to an insect order and family. Each order and family contains only a fraction of the approximately 300–400 species that are capable of causing damage to vegetables. You can greatly simplify identification by knowing the order, and perhaps the family. Usually, only a few characteristics such as wing number and leg or mouthpart type are needed to assign an unknown pest to order and family. The “quick guide” in this section and a key in Appendix A will aid you in this process.

There are three guiding principles to identification that, if followed, enhance correct identification. Remember that some species can be identified only by authorities, and some cannot be distinguished based only on appearance, but most serious pests are readily distinguished but little effort. You may need to use some low-level magnification such as a 10 × hand lens, and such equipment is inexpensive and easy to use.

WHY IDENTIFICATION IS SO IMPORTANT? There is a great wealth of information about vegetable pests. Hundreds of careers have been devoted to determining which species are damaging, the nature of their damage, the life cycle of the pest, and how the pest is best managed to eliminate or minimize injury to plants. It is easy to overestimate the damage potential of some large, leaf-feeding pests, and similarly easy to underestimate the damage caused by small and sap-sucking species. Failure to recognize the presence of certain pests, such as plant virus vectors early in the season, can prove a grievous error, resulting in severe crop injury. On the other hand, sometimes vegetable plants are treated with an insecticide, because lady beetles or other beneficial insects are mistaken for pests. Therefore, the collective knowledge and experience of generations of entomologists are available to identify the pest—but are useless unless correct identification is made of the pest at hand.

Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00002-9 © 2020 Elsevier Inc. All rights reserved.

1. Although there are numerous pests, most are easily assigned to groups based on easy-to-discern characters such as the number of legs or wings and the type of mouthparts. With a little effort, it is easy to distinguish such common groups as aphids, caterpillars, whiteflies, and wireworms. 2. Behavior is as important as appearance in distinguishing the pests. How or where a pest feeds is critical knowledge. The presence of silk webbing, the positioning of eggs, and other aspects of pest biology are often key elements in distinguishing the pests. When attempting to identify an unknown pest, try to observe as much as possible about the pest behavior and plant damage. 3. Details are important; look carefully at the pest and its damage, and do not trust your memory. Collect the pest you want to identify, do not just kill it. No one who is unfamiliar with the pest’s identity can anticipate what characters will be critical in distinguishing it from its relatives. It is entirely too easy to fail to notice the number of wings or the color of the legs, or not to remember to count the stripes or spots. Compare the written descriptions with the physical evidence carefully. Do not ignore aspects of the ­ description

23

24  Handbook of Vegetable Pests

b­ecause you are unfamiliar with the terminology. Entomologists, like members of all scientific disciplines, use technical words to describe the physical appearance and behavior of their subjects of study. In this book, the use of terminology is minimized, and terms are described in the glossary and illustrated with diagrams. If a pest poses a major economic threat to a crop and you are uncertain of its identity, there are several available sources of assistance. The principal source in the United States is the Cooperative Extension System associated with the Land-Grant University found in each state. A cooperative extension service office is normally found in each

county, and the employees are commonly called county agents. The county agents often can assist with identification, and if they cannot they will forward the pest to the state university for identification. Research centers and experiment stations are other good sources of expertise, in both Canada and the United States. Crop consultants provide identification and pest management recommendations on a for-fee basis and usually can be found in important agricultural areas. You can access considerable additional information to aid in identification if you have access to the Internet. Suggested sources of supplementary information and assistance are found in Appendix B.

Quick guide to orders of vegetable-feeding insects. Order of insects

Common name

Blattodea

Termites, cockroaches

Example of adult

FIG. 2.1  Termite

FIG. 2.2  Cockroach

Front wings

Hind wings

Various

Various

Pest Identification Chapter | 2  25

Quick guide to orders of vegetable-feeding insects (Continued) Order of insects

Common name

Coleoptera

Beetles, weevils, grubs

Example of adult

FIG. 2.3  Weevil

Front wings

Hind wings

Thickened, hard, meeting in straight lines on back

Membranous, folding beneath front wings

Short, thickened (if present)

Large, membranous, folding fan-like under front wings (if present)

Membranous

Absent

FIG. 2.4  Beetle

Dermaptera

Earwigs

FIG. 2.5  Earwig

Diptera

Files, maggots

FIG. 2.6 (Continued)

26  Handbook of Vegetable Pests

Quick guide to orders of vegetable-feeding insects (Continued) Order of insects

Common name

Hemiptera

Plant, seed, stink, lace, and leaf-footed bugs

Example of adult

Front wings

Hind wings

Membranous

Membranous

FIG. 2.7

Antennae

Mouthparts

Other features

Short to moderately long

Chewing

Three pairs of legs

Example of immature

Immature

Various FIG. 2.8  Termite

FIG. 2.9  Cockroach

Pest Identification Chapter | 2  27

Quick guide to orders of vegetable-feeding insects (Continued) Antennae

Mouthparts

Other features

Short to moderately long, thread-like or clubbed

Chewing

Usually with 3 paris of legs

Example of immature

FIG. 2.10  Beetle grub

Immature

Usually with distinct head capsule and three pairs of thoracic legs

FIG. 2.11  Beetle grub

Moderately long, thread-like

Chewing

Large forceps-like cerci

Resemble adults except for reduced wings

FIG. 2.12  Earwig Short, variable in from

Lappingsucking

Small balancing organs (halteres) in place of hind wings

Head reduced; legs absent FIG. 2.13  Maggot (Continued)

28  Handbook of Vegetable Pests

Quick guide to orders of vegetable-feeding insects (Continued) Antennae

Mouthparts

Other features

Example of immature

Usually moderately long four to five segments

Piercingsucking, originating at front of head

Sometimes resembling beetles

Immature

Resemble adults except for reduced wings

FIG. 2.14  Bug nymph

Quick guide to the important orders of vegetable-feeding insects. Order of insects

Common name

Example of adult

Front wings

Hind wings

Membranous

Membranous

Membranous

Membranous, smaller than front wings

Broad, with scales

Broad, with scales

FIG. 2.15  Aphid

Hemiptera

Aphids, leafhoppers, whiteflies, psyllids

FIG. 2.16  Leafhopper

Hymenoptera

Sawflies, ants

FIG. 2.17  Ant

Lepidoptera

Moths, butterflies, caterpillars, cutworms, loopers, webworms FIG. 2.18  Moth

Pest Identification Chapter | 2  29

Quick guide to the important orders of vegetable-feeding insects (Continued) Order of insects

Common name

Orthoptera

Grasshoppers, crickets, katydids

Example of adult

Front wings

Hind wings

Thickened (if present), many veins

Large, membranous, folding fan-like under front wings (if present)

Very slender, with fringe of long hairs

Very slender, with fringe of long hairs

FIG. 2.19  Grasshopper

Thysanoptera

Thrips

FIG. 2.20  Thrips

Antennae

Mouthparts

Other features

Example of immature

Immature features

FIG. 2.21  Immature

Usually short

Piercingsucking, originating at back of head

Extremely diverse, sometimes with waxy or threadlike exudates

Often resemble adults, but sometimes flattened

FIG. 2.22  Adult (Continued)

30  Handbook of Vegetable Pests

Quick guide to the important orders of vegetable-feeding insects (Continued) Antennae

Mouthparts

Thread-like

Chewing

Other features

Example of immature

Immature features Leg number variable, sawflies with seven pairs of prolegs in addition to thoracic legs

FIG. 2.23  Slug caterpillar Usually moderately long, threadlike, feathery, or clubbed

Coiled sucking tube

Moderately long to long, thread-like

Chewing

Two to five pairs of prolegs in addition to thoracic legs FIG. 2.24  Caterpillar

Hind legs well developed

Resemble adults except for reduced wings FIG. 2.25  Hopper

Short

Minute, piercing-sucking

Minute in size

Resemble adults except for reduced wings

FIG. 2.26  Immature thrips

Pest Identification Chapter | 2  31

Quick guide to the adults of important families of vegetable-feeding beetles. Family

Common name

Example of adult

Flea beetles

Antennae

Other features

Moderately long, thread-like

Small, often metallic

Moderately long, weekly clubbed and branched

Tip of abdomen exposed, not covered by etytra

Short to moderately long, thread-like

Pronotum and elytra expanded, flaring over legs; colorful

Moderately long

Elongate body form; colorful

FIG. 2.27  Flea beetles

Pea and bean seed beetles

Chrysomelidae

FIG. 2.28  Seed beetles

Tortoise beetles

FIG. 2.29  Tortoise

Leaf beetles

FIG. 2.30  Leaf (Continued)

32  Handbook of Vegetable Pests

Quick guide to the adults of important families of vegetable-feeding beetles (Continued) Family

Common name

Curculionidae

Weevils

Example of adult

Antennae

Other features

Short to moderately long, elbowed and clubbed

Elongate head forming shout

Short, clubbed

Conves; colorful, spotted

Short, branched

Elongate

Moderately long, thread-like

Colorful or metallic

FIG. 2.31  Weevils

Coccinellidae

Lady beetles

FIG. 2.32  Ladies

Elateridae

Click beetles

FIG. 2.33

Meloidae

Blister beetles

FIG. 2.34  Blister beetles

Pest Identification Chapter | 2  33

Quick guide to the adults of important families of vegetable-feeding beetles (Continued) Family

Scarabaeidae

Common name

Example of adult

Scarab and June beetles

Antennae

Other features

Short, branched, or clubbed

Convex; legs spiny

FIG. 2.35  Scarabs

Quick guide to the adults of important families of vegetable-feeding flies. Family

Common name

Agromyzidae

Leafminers

Example of adult

Antennae

Other features

Short

Small; wings transparent

Short

Medium size; wings transparent; legs medium length

Short to moderately long

Patterned wings

FIG. 2.36  Leafminers

Anthomyiidae

Root maggots and leafminers

FIG. 2.37  Root maggots

Ulidiidae

Picture-wing flies

FIG. 2.38  Ulidiids (Continued)

34  Handbook of Vegetable Pests

Quick guide to the adults of important families of vegetable-feeding flies (Continued) Family

Common name

Tephritidae

Fruit flies

Example of adult

Antennae

Other features

Short

Patterned wings

FIG. 2.39  Fruit flies

Quick guide to the adults of important families of vegetable-feeding bugs. Family

Common name

Coreidae

Squash and leaffooted bugs

Example of adult

Antennae

Other features

Moderately long, four segments

Front wing veins highly branched

Moderately long, four segments

Small size

FIG. 2.40  Squash bugs

Pseudococcidae

Mealybugs

FIG. 2.41  Mealybugs

Pest Identification Chapter | 2  35

Quick guide to the adults of important families of vegetable-feeding bugs (Continued) Family

Common name

Miridae

Plant bugs

Example of adult

Antennae

Other features

Moderately long, four segments

Slender body

Moderately long, five segments

Shield-shaped body

FIG. 2.42  Mirids

Pentatomidae

Stink bugs

FIG. 2.43  Stink bugs

Quick guide to the adults of important families of vegetable-feeding Hemiptera. Family

Common name

Aleyrodidae

Whiteflies

Example of adult

Antennae

Other features

Thread-like, seven segments

Opaque white wings

FIG. 2.44  Whiteflies (Continued)

36  Handbook of Vegetable Pests

Quick guide to the adults of important families of vegetable-feeding Hemiptera (Continued) Family

Common name

Aphididae

Aphids

Example of adult

Antennae

Other features

Thread-like, three to six segments

Wings transparent; cornicles usually present

Small

Front wings thickened and opaque

Long, 9 to 11 segments

Wings transparent; cornicles absent

FIG. 2.45  Aphids

Cicadellidae

Leafhoppers

FIG. 2.46  Leafhopper

Psyllidae

Psyllids

FIG. 2.47  Psyllids

Quick guide to the adults of important families of vegetable-feeding moths and butterflies. Family

Common name

Erebidae

Tiger and wooly moths

Example of adult

FIG. 2.48  Erebid moth

Wings

Other features

Front wings much longer than hind wings

Nocturnal; robust body, moderately large

Pest Identification Chapter | 2  37

Quick guide to the adults of important families of vegetable-feeding moths and butterflies (Continued) Family

Common name

Gelechiidae

Leafminer moths

Example of adult

Wings

Other features

Fringed

Nocturnal; small size

Muted colors, usually brownish

Nocturnal; robust body, moderately large

Wings form triangle when at rest

Nocturnal; palps protrude markedly from head, smallto medium-size

Front wings long and pointed

Nocturnal; large size

Hind wing with protruding lobe or “tail”

Diurnal; large size

White or yellow with black spots

Diurnal; moderate size

FIG. 2.49  Gelechid moth

Noctuidae

Armyworms, cutworms, and loopers

FIG. 2.50  Armyworm moth

Crambidae

Borers, webworms

FIG. 2.51  Webworm

Sphingidae

Hawk or sphinx moths

FIG. 2.52  Sphinx moth

Papilionidae

Swallowtails

FIG. 2.53  Swallowtail

Pieridae

White and sulfur butterflies

FIG. 2.54  White

38  Handbook of Vegetable Pests

Quick guide to the adults of important families of vegetable-feeding grasshoppers and crickets. Family

Common name

Acrididae

Grasshoppers

Example of adult

Antennae

Other features

About onehalf length of the body

Medium- to largebody size; usually colorful

Length of the body or longer

Medium-body size; color black

About length of the pronotum

Front legs enlarged for digging; color brownish

Length of the body or longer

Large body size; color variable

FIG. 2.55  Hoppers

Gryllidae

Field crickets

FIG. 2.56  Field crickets

Gryllotalpidae

Mole crickets

FIG. 2.57  Mole crickets

Tettigoniidae

Mormon crickets

FIG. 2.58  Mormon crickets

Pest Identification Chapter | 2  39

Quick guide to major noninsect groups of pests affecting vegetables. Class of noninsects

Common name

Example of adult

Front wings

Hind wings

None

None

None

None

FIG. 2.59  Mite Acari

Mites

FIG. 2.60  Mite

Collembola

Springtails FIG. 2.61  Springtails

(Continued)

40  Handbook of Vegetable Pests

Quick guide to major noninsect groups of pests affecting vegetables (Continued) Class of noninsects

Common name

Diplopoda

Millipedes

Example of adult

Front wings

Hind wings

None

None

None

None

None

None

FIG. 2.62  Mite

FIG. 2.63  Slug Gastropoda

Slugs, snails

FIG. 2.64  Snail

Isopoda

Pillbugs, sowbugs

FIG. 2.65  Bug

Pest Identification Chapter | 2  41

Quick guide to major noninsect groups of pests affecting vegetables (Continued) Class of noninsects

Common name

Symphyla

Symphylans

Example of adult

Front wings

Hind wings

None

None

FIG. 2.66  Symphylan

Antennae

Mouthparts

Other features

Immature features

Minute

Piercing-sucking

Usually four pairs of legs

Resemble adults

Short to moderately long

Chewing

Three pairs of legs

Resemble adults

Minute

Chewing

Two pairs of legs per body segment

Resemble adults

Extrudable

Hidden

Produce mucus; shell present or absent

Resemble adults

Minute

Chewing

Seven pairs of legs

Resemble adults

Moderately long

Chewing

12 pairs of legs

Resemble adults

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Chapter 3

Guides to Pest Identification, Arranged by Plant Taxon

The following are some guides to assist in identification of common vegetable pests. The guides are based on the characters that are easy to observe, and thus consist of obvious attributes such as the portion of the plant is affected, general morphological characteristics such as the presence or the absence of legs or wings, the mouth parts or feeding habits (chewing or sucking insects), and the general category of pest (commonly orders or families). To use the guides effectively, the reader must have some basic knowledge of pests, but with only a little experience it is relatively easy to narrow down a candidate pest to just a few possibilities. Once you have some idea of the identity, the individual descriptions and keys should aid in positive identification. Some pests are commonly associated with certain crops, and often cause damage. The identification guides are designed to assist you in identifying these common pests. However, there are other pests that are found only occasionally or infrequently. These are not integrated into the main identification guides, but follow the main list as “other pests.” This arrangement or prioritization of pests is not perfect because at a certain location or time the minor pests or “other pests” may assume significance. However, you are unlikely to find the “other” pests regularly damaging. The crops in the pest guides are arranged by plant family rather than by individual crop. This reduces redundancy when listing pests. The clustering of crops works quite well because insects usually make their food selection based on the plant chemistry, and plants in the same family share similar chemistry. Though it may not be obvious that crops such as Brussels sprouts and radish are related, insects detect their similar chemistry and thus these crops share many common pests. If you are uncertain of crop plant relationships, consult Appendix B.

Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00003-0 © 2020 Elsevier Inc. All rights reserved.

GUIDE TO COMMON PESTS AFFECTING ASPARAGUS Pests Boring into Roots Leafminers: Asparagus Symphylans: Garden

Pests Producing Small Mines in Stems Leafminers: Asparagus

Pests Feeding Externally on Leaves (Cladophylls) or Stems Pests with chewing mouthparts Armyworms and cutworms: Red-backed cutworm Caterpillars: Saltmarsh Leaf beetles: Asparagus, spotted asparagus Scarab beetles: Japanese Pests with piercing-sucking mouthparts Aphids: Asparagus Thrips: Bean, flower, onion

Pests Boring into Berries Leaf beetles: Spotted asparagus

Other Pests Occasionally Found on Asparagus Aphids: Bean, cowpea, green peach, melon, potato Armyworms and cutworms: Beet, fall, southern, yellowstriped and western yellowstriped armyworm; corn earworm; zebra caterpillar; black, spotted, sweetpotato, variegated, and velvet cutworm, Old World bollworm

43

44  Handbook of Vegetable Pests

Blister beetles: Black, immaculate, spotted, striped Borers: Stalk Grasshoppers: Differential, migratory, twostriped Loopers: Soybean Mites: Bulb Other caterpillars: Yellow woollybear Plant bugs: Alfalfa, pale legume, tarnished ,western tarnished; garden fleahopper Scarab beetles: Chinese rose Stink bugs: Green, harlequin, onespotted, Say Wireworms: False

GUIDE TO COMMON PESTS AFFECTING BEAN AND RELATED CROPS (Bean, Chickpea, Cowpea, Faba Bean, Lentil, Lima Bean, Pea, and Others)

Pests Boring into Roots or Stems Pests with legs Borers: Lesser cornstalk Flea beetles: Palestriped Leaf beetles: Bean; banded, spotted, striped, and western striped cucumber; grape colaspis Scarab beetles: Green June Symphylans: Garden Pests without legs Maggots: Bean seed, seedcorn Weevils: Pea leaf, vegetable; whitefringed beetle

Pests Producing Small Mines in Leaves Leafminers: American serpentine, pea, vegetable

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and cutworms: Armyworm, beet ­armyworm, and zebra caterpillar Leafrollers: Bean Leafminer: Tomato Loopers: Alfalfa, cabbage, and soybean looper, bean leafskeletonizer, green cloverworm Other caterpillars: Saltmarsh caterpillar, yellow woollybear, gray hairstreak Flea beetles: Palestriped, potato, western potato Leaf beetles: Bean; banded, spotted, striped, and western striped cucumber Other beetles: Mexican bean Weevils: Pea leaf, vegetable, whitefringed beetle Pests with piercing-sucking mouthparts Aphids: Bean, cowpea, green peach, pea Leafhoppers: Beet, potato, western potato Mites: Broad; strawberry, tumid, and twospotted spider

Other bugs: Leaffooted Plant bugs: Garden fleahopper, tarnished, western tarnished Stink bugs: Brown, green, onespotted, Say, southern green, brown marmorated Thrips: Bean, melon, onion, western flower Whiteflies: Chilli, common blossom, silverleaf, sweetpotato, greenhouse

Pests Feeding on Flowers, Seeds, or Seedpods Armyworms and cutworms: Corn earworm, western bean cutworm Borers: Pea moth, limabean pod, European corn, gray hairstreak Loopers: Bean leafskeletonizer Stink bugs: Brown, green, onespotted, redbanded, Say, southern green, brown marmorated Other bugs: Leaffooted, tarnished plant Weevils: Bean, broadbean, cowpea, pea, and southern cowpea weevil, cowpea curculio

Other Pests Occasionally Found on Bean and Related Crops Ants: Red imported fire Aphids: Bean root, blue alfalfa, buckthorn, foxglove, melon, potato Armyworms and cutworms: Bertha, fall, southern, sweetpotato, velvet, yellowstriped, and western yellowstriped armyworms; army, black, clover, darksided, dingy, glassy, granulate, pale western, redbacked, spotted, and variegated cutworms; tobacco budworm Blister beetles: Black, immaculate, spotted, striped, Old World bollworm Borers: Stalk Crickets: Fall, spring and southeastern field; Mormon Earwig: European Flea beetles: Redheaded, smartweed, tobacco, tuber, western black Flies: Melon Grasshoppers: American, differential, migratory, redlegged, twostriped Loopers: Bilobed and plantain Maggots: Radish root Mealybug: Jack Beardsley Millipedes: Garden Other caterpillars: Alfalfa caterpillar, banded woollybear Pillbug: Common Plant bugs: Rapid, superb, legume, pale legume Scarab beetles: Asiatic, oriental, spring rose, Japanese, white grubs Slugs

Guides to Pest Identification, Arranged by Plant Taxon Chapter | 3  45

Snails Springtails: Garden Stink bugs: Harlequin Thrips: Tobacco Webworms: Alfalfa, beet, garden; celery leaftier Wireworms: Eastern field, false, Great Basin, Pacific Coast, sugarbeet, wheat

GUIDE TO COMMON PESTS AFFECTING BEET AND RELATED CROPS (Beet, Chard, Spinach, Swiss Chard)

Pests Feeding on Roots Aphids: Sugarbeet root Maggot: Sugarbeet root Scarab beetles: Green June Symphylans: Garden

Pests Producing Mines in Leaves Leafminer: Beet, spinach

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and cutworms: Beet and southern ­armyworm, clover and variegated cutworm, winter cutworm zebra caterpillar Blister beetles: Black, immaculate, spotted, striped Flea beetles: Palestriped, potato, tuber Other caterpillars: Saltmarsh Webworms: Alfalfa, beet, garden, Hawaiian beet, southern beet, spotted beet Pests with piercing-sucking mouthparts Aphids: Bean, green peach Leafhoppers: Beet Thrips: Common blossom

Other Pests Occasionally Found on Beet and Related Crops Aphids: Bean, buckthorn, foxglove, melon, potato Armyworms and cutworms: European pepper moth, armyworm, bertha, fall armyworm, glassy, pale western, redbacked, sweetpotato, velvet, yellowstriped, and western yellowstriped armyworm; corn earworm; army, black, granulate, and spotted cutworm Borer: European corn, lesser cornstalk, stalk Crickets: Fall, southeastern, and spring field; shortwinged, southern, and tawny mole; Mormon Earwig: European Flea beetles: Eggplant, hop, redheaded, spinach, smartweed, sweetpotato, threespotted, western black, western potato, yellownecked

Flies: European crane Grasshoppers: Differential, migratory, redlegged, twostriped Leaf beetles: Banded cucumber, grape colaspis, spotted, and western spotted cucumber Leafhopper: Western potato Leafminer: American serpentine, pea Loopers: Alfalfa, cabbage, celery Maggots: Seedcorn Mites: Bulb, broad, twospotted spider Other bugs: False chinch Other caterpillars: Banded and yellow woollybear; whitelined sphinx Plant bugs: Alfalfa, garden fleahopper, tarnished, western tarnished, pale legume Scarab beetles: Asiatic garden, carrot, Japanese, oriental; white grubs Slugs Snails Springtails: Garden Stink bugs: Harlequin Thrips: Bean, onion, tobacco Webworms: Celery and false celery leaftier Weevil: Vegetable Wireworms: Eastern field, Gulf, Pacific Coast, southern potato, sugarbeet, tobacco

GUIDE TO COMMON PESTS AFFECTING CABBAGE AND RELATED CROPS (Broccoli, Brussels Sprouts, Cabbage, Cauliflower, Chinese Cabbage, Collards, Kale, Mustard, Radish, Turnip, and Others)

Pests Boring into Roots or Stems Pests with legs Flea beetles: Cabbage, crucifer, horseradish, striped, western black, western striped Scarab beetles: Green June, white grubs Termites: Subterranean Pests without legs Maggots: Cabbage, radish, seedcorn, Swede midge, turnip root Weevils: Cabbage curculio, vegetable, whitefringed beetle

Insects Producing Small Mines in Leaves Pests with legs Flea beetles: Horseradish and Zimmermann’s Webworms: Cabbage, oriental cabbage Pests without legs Leafminers: Cabbage, pea

46  Handbook of Vegetable Pests

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Caterpillars Armyworms and cutworms: Zebra caterpillar Cabbageworms: Cross-striped, imported, purplebacked, southern, mustard and southern white, winter cutworm Loopers: Alfalfa, cabbage Other caterpillars: Cabbage budworm, diamondback moth Webworms: Cabbage and oriental cabbage Beetles Flea beetles: Cabbage, crucifer, horseradish, striped, western black, western striped, Zimmermann’s Leaf beetles: Red turnip, yellowmargined Weevils: Vegetable Cockroaches Asian, Surinam Pests with piercing-sucking mouthparts Aphids: Cabbage, green peach, turnip Stink bugs: Harlequin, painted Thrips: Onion Whiteflies: Silverleaf, sweetpotato

Pests Feeding on Flowers, Seeds, Or Seedpods Cabbageworms: Southern Weevils: Cabbage seedpod

Other Pests Occasionally Found on Cabbage and Related Crops Ant: Red imported fire Aphids: Buckthorn, potato Armyworms and cutworms: Beet, bertha, fall, southern, yellowstriped, and western yellowstriped armyworm; corn earworm; army, black, clover, darksided, dingy, glassy, granulate, redbacked, spotted, and variegated cutworm, Old World bollworm Blister beetles: Black, immaculate, spotted, striped Borers: European corn, lesser cornstalk, stalk Cockroach: Asian Crickets: Fall, spring, and southeastern field; shortwinged, southern, and tawny mole; Mormon Earwig: European, ringlegged Flea beetles: Hop, palestriped, potato, redheaded, southern tobacco, smartweed, tobacco, tuber, western potato Flies: European crane, small fruit Grasshoppers: Differential, eastern lubber, migratory, twostriped Leaf beetles: Banded and spotted cucumber Leafhoppers: Aster, western potato Loopers: Bean leafskeletonizer; bilobed, celery, plantain, soybean

Millipedes: Garden Other bugs: False chinch Other caterpillars: Saltmarsh, whitelined sphinx, yellow woollybear Pillbugs: Common Plant bugs: Garden fleahopper, pale legume, tarnished, western tarnished Scarab beetles: Asiatic garden, Chinese rose, Japanese Slugs Springtails: Garden Snails Stink bugs: Brown, green, Say, southern green Thrips: Bean, melon Webworms: Celery and false celery leaftier; alfalfa, beet, garden Weevils: Cabbage curculio Whiteflies: Greenhouse Wireworms: Corn, eastern field, false, Gulf, Oregon, Pacific Coast, potato, southern, sugarbeet, tobacco, wheat

GUIDE TO COMMON PESTS AFFECTING CARROT AND RELATED PLANTS (Carrot, Celery, Celeriac, Chervil, Coriander, Fennel, Parsley, Parsnip)

Pests Boring into Stems or Feeding on Roots Aphids: Carrot root Flies: Carrot rust Scarab beetles: Green June Symphylans: Garden Weevils: Carrot, Texas carrot, vegetable

Pests Producing Small Mines in Leaves Leafminers: American serpentine, parsnip, and pea

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and cutworms: Beet, fall, and southern armyworm; black and granulate cutworm, winter cutworm Flea beetles: Potato, western potato Loopers: Alfalfa, cabbage, celery Other caterpillars: Black and anise swallowtails, saltmarsh caterpillar Webworms: Celery and false celery leaftier Weevils: Vegetable Pests with piercing-sucking mouthparts Aphids: Coriander, green peach, honeysuckle, melon, willow carrot Leafhopper: Aster

Guides to Pest Identification, Arranged by Plant Taxon Chapter | 3  47

Mites: Twospotted spider Plant bugs: Carrot, pale legume, tarnished, western tarnished plant; garden fleahopper Stink bugs: Southern green

Pests Feeding on Flowers or Seeds Webworms: Parsnip Plant bugs: Carrot, tarnished

Other Pests Occasionally Found on Carrot and Related Crops Aphids: Bean, bean root, buckthorn, cowpea, foxglove, potato Armyworms and cutworms: European pepper moth, yellowstriped and western yellowstriped armyworm; army, dingy, pale western, and spotted cutworm; ­zebra caterpillar Blister beetles: Black, immaculate, spotted, striped Crickets: Fall, spring, and southeastern field; shortwinged, southern and tawny mole; Mormon Earwigs: European Flea beetles: Palestriped, redheaded, smartweed Grasshoppers: Differential, eastern lubber, migratory, redlegged, twostriped Leafhoppers: Western potato Leaf beetles: Banded cucumber Looper: Bean leafskeletonizer, plantain, soybean Maggots: Seedcorn Mealybug: Jack Beardsley Millipedes: Garden Mites: Bulb Other bugs: False chinch, leaffooted, little negro Other caterpillars: Yellow woollybear Plant bugs: Rapid, superb Scarab beetles: Asiatic garden, carrot, white grubs Slugs Symphylans: Garden Thrips: Onion, tobacco Webworms: Alfalfa, beet, garden Weevils: Whitefringed beetle Whiteflies: Silverleaf, sweetpotato Wireworms: Eastern field, Great Basin, Gulf, southern potato, tobacco, wheat

GUIDE TO COMMON PESTS AFFECTING LETTUCE AND RELATED CROPS (Artichoke, Celtuce, Chicory, Endive, Escarole, Lettuce, Radicchio)

Pests Feeding on Roots Aphids: Lettuce root

Scarab beetles: Green June, white grubs Symphylans: Garden

Pests Producing Mines in Leaves Leafminers: American serpentine, pea, vegetable

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and cutworms: Armyworm, beet, yellowstriped, western yellowstriped armyworm; corn earworm; zebra caterpillar; black, granulate, and variegated cutworm, winter cutworm Cockroaches: Asian, Surinam Flea beetles: Palestriped, spotted, and western spotted cucumber Grasshoppers: American, differential, eastern lubber, migratory, redlegged, twostriped Loopers: Alfalfa, cabbage, celery Webworms: Celery leaftier Other caterpillars: Saltmarsh, artichole plume Slugs Snails Pests with piercing-sucking mouthparts Aphids: Artichoke, bean, cowpea, green peach, lettuce, potato Leafhoppers: Aster, potato, western potato Plant bugs: Garden fleahopper; tarnished, western tarnished, and pale legume Stink bugs: Southern green Thrips: Western flower Other bugs: Leaffooted Whiteflies: Common blossom, greenhouse, silverleaf, sweetpotato Mites: Twospotted spider

Other Pests Occasionally Found on Lettuce and Related Crops Aphids: Bean, cowpea, foxglove Armyworms and cutworms: European pepper moth; bertha armyworm; black, clover, dingy, glassy, granulate, redbacked, and spotted cutworm; tobacco budworm; corn earworm; saltmarsh Crickets: Fall, spring, and southeastern field; shortwinged, southern, and tawny mole; Mormon Earwigs: European, redlegged Flea beetles: Palestriped, potato, redheaded, smartweed, tuber, western black, western potato Flies: European crane Leaf beetles: Banded and spotted cucumber Leafhoppers: Western potato Loopers: Bilobed, soybean Maggots: Seedcorn

48  Handbook of Vegetable Pests

Millipedes: Garden Other bugs: False chinch Other caterpillars: Whitelined sphinx Pillbug: Common Springtail: Garden Stink bugs: Harlequin, Say Thrips: Bean, melon and western flower Webworm: Alfalfa, beet, garden; celery and false celery leaftier Weevils: Vegetable Wireworms: Corn, eastern field, false, Great Basin, Oregon, Pacific Coast, sugarbeet

GUIDE TO IDENTIFICATION OF COMMON INSECT PESTS AFFECTING OKRA Pests Feeding on Roots or Lower Stem Leaf beetles: Banded and striped cucumber Termites: Subterranean

Pests Producing Mines in Leaves Leafminers: Vegetable

Pests Boring into Stems Borers: European corn

Pests Feeding Externally on Leaves Pests with chewing mouthparts Armyworms and cutworms: Corn earworm; ­yellowstriped and southern armyworm Leaf beetles: Spotted cucumber Loopers: Okra caterpillar Other caterpillars: Gray hairstreak Scarab beetles: Japanese Cockroach: Asian Pests with piercing-sucking mouthparts Aphids: Green peach, melon Plant bugs: Garden fleahopper Mealybug: Cotton Mites: Twospotted and tumid spider Thrips: Melon Whiteflies: Silverleaf, sweetpotato

Pests Feeding on Blossoms or Fruits Ants: Red imported fire Armyworms and cutworms: Corn earworm Bugs: Leaffooted Other caterpillars: Gray hairstreak Stink bugs: Brown, brown marmorated, green, southern green

Other Pests Occasionally Found on Okra Plants Armyworms and cutworms: Black cutworm Blister beetles: Black Flea beetles: Palestriped, redheaded, smartweed Grasshoppers: American Leaf beetles: Grape colaspis, striped, and western striped cucumber Leafhoppers: Potato, western potato Leafminers: Pea Loopers: Soybean Mealybug: Madeira Scarab beetles: Chinese rose, Japanese, white grubs Stainer: Cotton Stink bugs: Harlequin Weevils: Whitefringed beetle Whiteflies: Greenhouse

GUIDE TO COMMON PESTS AFFECTING ONION AND RELATED PLANTS (Chive, Garlic, Leek, Onion, Shallot)

Pests Boring into Bulbs or Roots Flies: Onion and lesser bulb Maggots: Onion, seedcorn Mites: Bulb

Pests Producing Small Mines in Leaves Leafminers: American serpentine, pea

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and caterpillars: Beet armyworm, leak moth, winter cutworm Other caterpillars: Saltmarsh Pests with piercing-sucking mouthparts Thrips: Common blossom, onion, tobacco, western flower

Other Pests Occasionally Found on Onion and Related Plants Aphid: Bean Armyworms and cutworms: Armyworm, fall, yellowstriped, and western yellowstriped armyworm; army, black, clover, darksided, dingy, granulate, pale western, redbacked, spotted, and variegated cutworm, Old World bollworm Blister beetles: Black, immaculate Borers: Potato stem Crickets: Shortwinged, southern, and tawny mole; Mormon

Guides to Pest Identification, Arranged by Plant Taxon Chapter | 3  49

Flea beetles: Palestriped Grasshoppers: Differential, eastern lubber, migratory, twostriped Leaf beetles: Banded cucumber Loopers: Alfalfa Maggots: Bean seed Plant bugs: Pale legume, tarnished, western tarnished Scarab beetles: Asiatic garden, oriental, white grubs Springtails: Garden Stink bugs: Onespotted Symphylans: Garden Thrips: Bean, melon Webworms: Alfalfa, beet, garden Weevils: Vegetable Wireworms: Eastern field, false, Great Basin

GUIDE TO COMMON PESTS AFFECTING SQUASH AND RELATED CROPS (Cucumber, Pumpkin, Squash, Cantaloupe, Watermelon, and Other Melons)

Pests Boring into Roots, Stems, Blossoms, or Fruit Pests with legs Borers: Pickleworm; melonworm; squash and southwestern squash vine Leaf beetles: Banded, spotted, striped, and western striped cucumber Pests without legs Flies: Melon and oriental fruit; small fruit

Pests Producing Small Mines in Leaves

GUIDE TO COMMON PESTS AFFECTING RHUBARB Pests Feeding on Roots Leaf beetles: Spotted cucumber beetle Weevils: Rhubarb curculio

Pests Living within Stems and Stalks Borers: Potato stem, stalk Weevils: Rhubarb curculio

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and cutworms: Spotted, variegated, and winter cutworm Flea beetles: Hop, potato Scarab beetles: Japanese Pests with piercing-sucking mouthparts Aphids: Bean, green peach, potato Leafhopper: Potato

Other Pests Occasionally Found on Rhubarb Armyworms and cutworms: Bertha, southern, yellowstriped, and western yellowstriped armyworm; army cutworm Borers: European corn Earwig: European Flea beetles: Palestriped Looper: Alfalfa Maggots: Seedcorn Other caterpillars: Yellow woollybear Scarab beetles: Asiatic garden, oriental, white grubs Symphylan: Garden Webworms: Alfalfa, beet, garden

Leafminers: American serpentine, pea, vegetable

Pests Feeding Externally on Leaves, Stems, Blossoms, or Fruit Pests with chewing mouthparts Armyworms and cutworms: Southern armyworm; granulate cutworm Cockroaches: Surinam Leaf beetles: Banded, spotted, striped, and western striped cucumber; western corn rootworm Other beetles: Squash Other caterpillars: Melonworm Pests with piercing-sucking mouthparts Aphids: Green peach, melon Leafhoppers: Potato; western potato Mealybug: Cotton, Jack Beardsley Mite: Broad; twospotted spider Other bugs: Garden fleahopper; leaffooted; squash and horned squash Thrips: Florida flower, melon, western flower Whiteflies: Chilli, common blossom, silverleaf, sweetpotato, greenhouse

Other Pests Occasionally Found on Squash and Related Crops Ants: Red imported fire Aphids: Bean, buckthorn, potato, rice root Armyworms and cutworms: European pepper moth, beet, fall, yellowstriped, and western yellowstriped armyworm; black, darksided, dingy, redbacked, and variegated cutworm; corn earworm, tobacco budworm Blister beetles: Black, immaculate Borers: Lesser cornstalk, stalk Crickets: Fall, spring, and southeastern field; shortwinged, southern, and tawny mole Earwig: European

50  Handbook of Vegetable Pests

Flea beetles: Palestriped, potato, tuber, western potato Flies: Oriental fruit Grasshoppers: American, differential, eastern lubber, migratory, twostriped Leaf beetles: Grape colaspis Leafhoppers: Beet Loopers: Alfalfa, cabbage, soybean Maggots: Bean seed, seedcorn Millipedes: Garden Other beetles: Whitefringed Other bugs: False chinch Other caterpillars: Whitelined sphinx, yellow woollybear Pillbugs: Common Plant bugs: Garden fleahopper, pale legume, tarnished, western tarnished Sap beetles: Dusky and fourspotted Scarab beetles: Carrot, Chinese rose, green June, white grubs Slugs Snails Springtails: Garden Stink bugs: Brown, green, onespotted, southern green, harlequin Symphylans: Garden Thrips: Onion, tobacco Webworms: Alfalfa, beet, garden Wireworms: Eastern, false, field, gulf, Pacific Coast, southern potato, sugarbeet, tobacco, wheat

GUIDE TO COMMON PESTS AFFECTING SWEET CORN Pests Feeding on Roots Aphids: Corn root Flea beetles: Corn, desert corn, redheaded, toothed Leaf beetles: Northern and western corn rootworms; spotted and banded cucumber Scarab beetles: Green June, white grubs Termites: Subterranean Webworms: Sod and root Weevils: Whitefringed beetle Wireworms: Corn, eastern field, false, Great Basin, Gulf, Oregon, Pacific Coast, southern potato, sugarbeet, tobacco, wheat

Pests Feeding on Seed or Seedling Armyworms and cutworms: Black, darksided, granulate, spotted, variegated, and winter cutworm Borers: Hop vine, potato stem Flea beetles: Corn, desert corn, redheaded, toothed Maggots: Bean seed, seedcorn Other beetles: Seedcorn, slender seedcorn

Webworms: Sod and root Weevils: Maize and southern corn

Pests Boring into Stems or Taproot Borers: European corn, lesser cornstalk, southern cornstalk, southwestern corn, stalk, sugarcane Weevils: Maize and southern corn billbug

Pests Producing Small Mines in Leaves Borer: Stalk Leafminer: Corn blotch Cutworm: Winter

Pests Feeding Externally on Leaves, Tassel, or Stalk Pests with chewing mouthparts Armyworms and cutworms: Armyworm, fall armyworm, corn earworm, winter cutworm Borers: European corn, southern cornstalk, southwestern corn, stalk, sugarcane Grasshoppers: American, differential, migratory, twostriped, redlegged Leaf beetles: Grape colaspis, western corn rootworm Scarab beetles: Japanese, sugarcane Webworms: Sod and root Weevils: Maize and southern corn billbug Pests with piercing-sucking mouthparts Aphids: Corn leaf, bird cherry-oat Mealybug: Cotton Mites: Twospotted spider, Banks grass Other bugs: Chinch Leaf and planthoppers: Corn delphacid, corn leafhopper Thrips: Florida flower, chilli, grass

Pests Feeding on Ears or Silk Armyworms and cutworms: Corn earworm, fall armyworm, western bean cutworm, Old World bollworm Borers: European, southwestern corn Earwigs: European Flies: Cornsilk Leaf beetles: Northern and western corn rootworm; spotted and banded cucumber Other beetles: Dusky and fourspotted sap Scarab beetles: Green June, Japanese

Other Pests Occasionally Found on Corn Ant: Red imported fire Aphids: Bean, green peach, potato Armyworms and cutworms: European pepper moth, army, beet, bertha, bronzed, glassy, and pale ­western

Guides to Pest Identification, Arranged by Plant Taxon Chapter | 3  51

cutworm; sweetpotato, velvet, yellowstriped, and western yellowstriped armyworm; zebra caterpillar Blister beetles: Black, striped Crickets: Mormon Flies: European crane Leaf beetles: Grape colaspis Leafhoppers: Aster and western potato leafhopper Loopers: Soybean Other caterpillars: Banded and yellow woollybear; saltmarsh Mealybug: Jack Beardsley Millipedes: Garden Plant bugs: Tarnished Scarab beetles: Asiatic, carrot, Chinese rose Symphylans: Garden Stink bugs: Brown, green, onespotted, southern green Slugs Thrips: Bean, chilli, tobacco Webworms: Alfalfa, beet, garden Weevils: Whitefringed beetle

GUIDE TO COMMON PESTS AFFECTING SWEET POTATO Pests Boring into Vines or Roots Borers: Sweetpotato vine Flea beetles: Sweetpotato Leaf beetles: Banded and spotted cucumber; sweetpotato Scarab beetles: Spring rose, sugarcane Wireworms: Corn, gulf, southern potato, tobacco Weevils: Sweetpotato and West Indian sweetpotato; whitefringed beetle

Pests Producing Small Mines in Leaves Leafminers: Morningglory, sweetpotato

Pests Feeding Externally on Leaves or Stems Pests with chewing mouthparts Armyworms and cutworms: Fall, southern, and yellowstriped armyworm; black, darksided, and granulate cutworm Cockroaches: Surinam Flea beetles: Sweetpotato Leaf beetles: Sweetpotato Other caterpillars: Sweetpotato hornworm Tortoise beetles: Argus, blacklegged, golden, mottled, striped Pests with piercing-sucking mouthparts Leafhoppers: Potato, western potato Whiteflies: Chilli, common blossom, silverleaf, sweetpotato

Other Pests Occasionally Found on Sweet Potato Ant: Red imported fire Aphids: Green peach, foxglove, melon, potato Armyworms and cutworms: Beet, sweetpotato, and velvet armyworm; army, dingy, and variegated cutworm; corn earworm Borers: Lesser cornstalk Blister beetles: Black, striped Crickets: Fall, spring, and southeastern field; shortwinged, southern, and tawny mole Earwing: Ringlegged Flea beetles: Elongate, palestriped, redheaded, smartweed Grasshopper: Differential, migratory, twostriped Loopers: Cabbbage, soybean Mites: Tumid, twospotted spider Maggots: Seedcorn Mealybug: Jack Beardsley Other bugs: Little negro Other caterpillars: Yellow woollybear Plant bugs: Garden fleahopper, pale legume, tarnished, western tarnished Sawflies: Sweetpotato Scarab beetles: Asiatic garden, carrot, Chinese rose, white grubs Stainer: Cotton Stink bugs: Southern green Thrips: Onion Weevils: Vegetable Whiteflies: Greenhouse

GUIDE TO COMMON PESTS AFFECTING TOMATO AND RELATED PLANTS (Eggplant, Pepper, Potato, Tomatillo, Tomato)

Pests Feeding on Roots, Tubers, or Lower Stem Borers: Potato tuberworm Bugs: Tomato Crickets: Shortwinged, southern, and tawny mole Armyworms and cutworms: Black, granulate, and variegated cutworm Flea beetles: Eggplant, potato, tuber, western potato Maggots: Seedcorn Symphylans: Garden Scarab beetles: Green June, white grubs Termites: Subterranean Weevils: Vegetable weevil, whitefringed beetle Wireworms: Corn, eastern field, Great Basin, southern potato, sugarbeet, tobacco

Pests Producing Mines in Leaves Leafminers: American serpentine, tomato, vegetable

52  Handbook of Vegetable Pests

Other caterpillars: Eggplant leafminer, potato tuberworm, tomato pinworm

Pests Boring into Stems Borers: European corn, potato stalk, potato stem, and stalk borer; potato tuberworm; tomato pinworm, tomato leafminer

Pests Feeding Externally on Leaves or Upper Stems Pests with chewing mouthparts Armyworms and cutworms: Beet, fall, southern, yellowstriped, and western yellowstriped armyworm, tobacco budworm, corn earworm; variegated and winter cutworm Cockroaches: Surinam Flea beetles: Eggplant, potato, tuber, western potato, tobacco, southern tobacco Leaf beetles: Colorado potato Leafminer: Tomato Loopers: Alfalfa, cabbage Other caterpillars: Tobacco and tomato hornworm, saltmarsh Tortoise beetles: Eggplant Weevils: Vegetable; whitefringed beetle Pests with piercing-sucking mouthparts Aphids: Buckthorn, foxglove, green peach, potato Lacebugs: Eggplant Leafhoppers: Aster, beet, potato, western potato Mealybug: Cotton Mites: Broad, tomato russet, twospotted spider Other bugs: Leaffooted Plant bugs: Garden fleahopper, tarnished, western tarnished Psyllids: Potato Stink bugs: Southern green Thrips: Florida flower, chilli, common blossom, melon, onion, western flower Whiteflies: Greenhouse, silverleaf, sweetpotato

Pests Feeding on Blossoms or Fruits Armyworms and cutworms: Beet, yellowstriped, southern armyworm, and western yellowstriped armyworm; corn earworm; variegated cutworm, Old World bollworm

Borers: Corn earworm, tobacco budworm, tomato pinworm Flies: Small fruit Maggots: Pepper Other bugs: Leaffooted Other caterpillars: Tobacco and tomato hornworm Sap beetles: Dusky Scarab beetles: Green June Stainer: Cotton Stink bugs: Brown, brown marmorated, consperse, green, onespotted, Say, southern green, tomato Weevils: Pepper

Other Pests Occasionally Found on Tomato and Related Plants Ant: Red imported fire Aphids: Bean root, rice root, melon Armyworms and cutworms: European pepper moth, army, bertha, black, bronzed, clover, darksided, dingy, granulate, pale western, redbacked, and spotted cutworm; zebra caterpillar, Old World bollworm Blister beetles: Black, immaculate, spotted, striped Borer: Lesser cornstalk Crickets: Fall, spring, and southeastern field; shortwinged, southern, and tawny mole; Mormon Earwigs: European, ringlegged Flea beetles: Hop, palestriped Flies: European crane, Mediterranean fruit Grasshoppers: American, differential, migratory, twostriped, redlegged Leaf beetles: Grape colaspis; spotted, striped, and western striped cucumber Leafminers: Pea Loopers: Soybean Mealybug: Jack Beardsley, Madeira Millipedes: Garden Mites: Bulb Other bugs: Little negro Other caterpillars: Whitelined sphinx, yellow woollybear Pillbugs: Common Plant bugs: Alfalfa, pale legume, rapid, superb Scarab beetles: Carrot, Chinese rose, Japanese Slugs Springtails: Garden Stink bugs: Harlequin Symphylans: Garden Thrips: Bean, tobacco Webworms: Alfalfa, beet, garden Wireworms: False, Gulf, Oregon, Pacific Coast, wheat

Part I

Class Insecta—Insects

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Chapter 4

Order Blattodea––Cockroaches and Termites

FAMILY ECTOBIIDAE Asian Cockroach

Blattella asahinai Mizukubo (Blattodea: Ectobiidae)

Natural History Distribution. Blattella asahinai cockroach was described only in 1981 from Okinawa, Japan. It closely resembles German cockroach, Blattella germanica (Linnaeus) (Blattodea: Ectobiidae), which is a recent discovery. Asian cockroach is found widely in southern Asia, and occasionally in tropical and subtropical environments. It apparently attained the United States in about 1986, when it was found near Tampa, Florida. It is expected eventually to inhabit all of the southern US and presently is found from South Carolina to Texas. Host Plants. Asian cockroach is considered to be omnivorous. Though not generally known as a plant pest, Asian cockroaches have long been known to damage young leaves of citrus trees and ripe strawberries in Florida, warranting chemical suppression. Brenner (1991) reported that Asian cockroach can completely consume the tassels of sweet corn, feed on cabbage, and consume rose and petunia flowers. They can attain high densities in organically maintained gardens, and infest nursery plants. In laboratory studies, it was possible to demonstrate that these cockroaches will nibble on leaves of several vegetable crop plants such as romaine and leaf lettuce, tomatillo, okra, cabbage, Brussels sprouts, and broccoli (Capinera, Unpublished). Their damage to vegetable plants generally is quite minor, and their

Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00004-2 © 2020 Elsevier Inc. All rights reserved.

greatest impact is as a contaminant of lettuce, as these insects hide between the layers of leaves, only to emerge after harvest. Natural Enemies. The natural enemies of Asian cockroach have not been studied, but in tropical environments, ants are very important predators of cockroaches, and determinants of cockroach distribution (Tarli et al., 2014). Life Cycle and Description. Asian cockroach requires about 66  days in the case of males, and 68  days in the case of females, to mature from an egg to an adult. The new adult then mates and produces an egg capsule in about 8  days. The female carries the egg capsule for about 19 days before the eggs hatch. Thus, the complete life cycle (egg hatch to egg hatch) at 25°C requires nearly 100 days, though temperature affects the rate of development (Atkinson et al., 1991). In Alabama and Georgia, the Asian cockroach population increases dramatically between May, when a small number of overwintering adults are present, and AugustSeptember, when immature cockroaches make up a large proportion of the population (Snoddy and Appel, 2008). Apparent abundance decreases during dry or cold weather, as the cockroaches burrow into the leaf litter and soil to escape adverse environmental conditions. All stages of this cockroach were found to overwinter in the substrate, though the overwintering population was highly skewed (about 10% adults and 90% nymphs), shifting to about 100% nymphs by May. During the summer months, the proportion of adults increased as the nymphs matured, but the absolute number of immatures increased markedly due to reproduction.

55

56  Handbook of Vegetable Pests

FIG. 4.1  Adult Asian cockroach. (Photo by L. Buss.)

Egg. The female of Asian cockroach carries an egg case, or ootheca, until just before egg hatch occurs. The ootheca protrudes from the posterior end (genital chamber) of the female. Nymphs will often hatch from the ootheca while still being carried by the female. Oothecae contain, on average, 30–40 eggs. The egg case is an elongate, quadrate capsule, with the corners rounded. It measures about 8 mm long, 3 mm high, and 2 mm wide. The females are capable of producing 5 or 6 egg capsule during her life, but under normal mortality situations may produce only 2 egg capsules, resulting in about 80 eggs, before perishing. Viable oothecae require about 20 days of incubation before hatching, whereas nonviable oothecae are dropped by the female within 4 days. Nymph. The number of instars varies, but six is normal. The nymphs are brown to black in color, with distinct dark dorsolateral stripes along the thorax that are separated by a light-brown central area. Short cerci protrude from the posterior end of the insect, and the head bears elongate antennae. Adult. The adult is 10–15 mm long, brown, and with two darker stripes running dorsolaterally along the length of the pronotum. The males are relatively slender, with the posterior abdomen tapered, the terminal segments of the abdomen covered by the wings, but the elongate cerci protruding. The females are relatively stout, with the posterior region of the abdomen rounded. The wings barely cover the sides of the abdomen but extend well beyond the tip. Cerci are evident in the female, as well as in the male. Females require 13  days after emergence to produce the first ootheca. Development and reproduction of Asian cockroach are given by Atkinson et al. (1991). Asian cockroaches are very similar to German cockroaches, and positive confirmation is usually made using chemical or molecular methods, or by dissection of the male tergal glands. However, there are some minor but consistent differences between the two species, as described and

pictured by Ross and Mullins (1988) and Richman (2017). Adult Asian cockroaches have longer and narrower wings and produce smaller egg capsules. The first instars of Asian cockroaches have 23 antennal segments whereas the corresponding instar of German cockroach has 24 or 25 segments. When the ootheca is being carried by the adult female of Asian cockroach, it does not extend as far beyond the tip of the abdomen as it does in the German cockroach. Also, the head capsule widths of first instar nymphs of Asian cockroach are slightly smaller (mean ± SE = 0.64 ± 0.006 mm) than German cockroaches (mean ± SE = 0.77 ± 0.007 mm), and the dark dorsolateral stripes that start on the pronotum and extend back along the abdomen are darker and more defined in Asian cockroach. Ultimately, the behaviors of the two species are probably the most useful diagnostic characters for distinguishing between German and Asiatic cockroaches. Asian cockroaches are active outdoors, they are strong fliers, and are attracted to light. German cockroaches tend to dwell in buildings, rarely fly, and are not attracted to light.

Damage Although considered to be a peridomestic cockroach, Asian cockroach is not a building-dwelling species and is normally found outdoors in wooded areas hiding in the leaf litter of wooded areas, citrus groves, and lawns (Brenner et al., 1988; Pfannenstiel et al., 2008). Population densities of 12,000–100,000 per ha have been recorded in Florida. Like other cockroaches, it is nocturnal but differs from some in being a strong flier and being attracted to lights. Thus, its principal contact with humans is around lights and at lighted entries to dwellings. Alternatively, raking tree leaf litter or digging in areas with accumulated organic debris will produce a large number of these cockroaches. In southern Florida, there are large expanses of sugar cane production, and large numbers of cockroaches can sometimes be found hiding in the dense vegetation of sugar cane fields. Sugar cane is a multiyear crop, often harvested for 3 consecutive years before being replanted, which can lead to accumulation of organic matter that favors cockroaches. These cockroaches can invade adjacent crops. As mentioned previously, their damage to plants generally is quite minor, with their greatest impact as a contaminant of lettuce, as these insects will hide between the layers of leaves, only to emerge after harvest. Perhaps offsetting the plant feeding damage potential of Asian cockroach are the predatory tendencies of this omnivore. Pfannenstiel et  al. (2008) reported that Asian cockroaches invaded the canopy of soybean during the evening, where they fed on eggs of corn earworm, Helicoverpa zea (Boddie), and beet armyworm, Spodoptera exigua (Hübner) (both Lepidoptera: Noctuidae). In southern Texas, this cockroach occurred at densities of about 50 per m of a row, making it one of the most abundant egg

Order Blattodea––Cockroaches and Termites Chapter | 4  57

predators. On the other hand, starved Asian cockroaches fed on parasitized brown citrus aphid, Toxoptera citricida (Kirkaldy) (Hemiptera: Aphididae) in citrus (Persad and Hoy, 2004), destroying the parasitoid developing within the aphid mummy, and did not seem to affect citrus psyllid, Diaphorina citri (Hemiptera: Psyllidae) (Qureshi and Stansly, 2009) in abundance.

Management The typical management practices for cockroach control do not apply to Asian cockroach, because residual sprays in and around the perimeter of buildings are not effective for an insect that dwells mostly in mulched areas and leaf litter. Thus, elimination of harborage should target mulch and leaf litter. Snoddy and Appel (2013) investigated preference of Asian cockroach for different mulch materials and documented a preference by males for oak leaf litter and pine straw, and by females for rubber mulches, relative to cypress mulch and topsoil. Nymphs, however, preferred rubber mulches, possibly because it provided smaller, more humid spaces for the nymphs. Thus, the use of cypress as a mulch material, or bare soil around plants in gardens, may reduce the abundance of Asian cockroaches in the home garden environment. Yellow incandescent bulbs used for porch lighting and sodium vapor lamps used for security lighting are unattractive to the Asian cockroaches and reduce the tendency of these insects to alight near lights. Also, pelletized toxic baits scattered outdoors in areas of harborage such as leaf litter can provide effective control (Richman, 2017), as can liquid and granular insecticides used for ants and termites (Snoddy and Appel, 2014) though these products typically are not labeled for use around food plants.

FAMILY BLABERIDAE Surinam Cockroach

Pycnoscelus surinamensis (Linnaeus) and

Indian Cockroach

Pycnoscelus indica (Fabricius) (Blattodea: Blaberidae)

Natural History Distribution. These two are often confused and are closely related species originated in the Indo-Malay region of southern Asia. Pycnoscelus indica is believed to be the ancestral species and is most common in Asia. Pycnoscelus surinamensis has been relocated and become cosmopolitan in the tropical and subtropical regions of the world. In the United States, P. surinamensis is found in southeastern states from North Carolina to Texas, in California, and in Puerto Rico. Its range also extends into Central and South

America, in areas with warm climates. However, P. indica is found in Hawaii. In North America, Surinam cockroach tends to be shipped with potted plants due to their burrowing habits, so it shows up in colder regions and can infest sheltered environments such as greenhouses. Recently, Zangl et  al. (2018) reported the presence of Surinam cockroach in Austria (Central Europe). DNA barcoding documented that this introduction was related to the insects found in the United States. Thus, this species continues to spread around the globe, seemingly without regard to the weather in the new environment. Host Plants. Both species prefer to hide under fallen leaves, mulch, and other debris, and also burrow in the soil. Wet soil is favored. There, they can damage the roots of plants such as sugarcane, tobacco, pineapple, and garden plants, and also potato tubers. They are negatively phototactic, hiding during the day. However, during the night hours these cockroaches emerge to nibble on above-ground plant material from vegetables such as lettuce, broccoli, cucumber, tomato, leaves of sweet potato, and the flowers and fruit of squash plants, including ornamentals and fruit such as roses, lilies, palms, papayas, and figs (Moretti et al., 2011; Capinera, Unpublished). As is the case with many nocturnal plant feeders, these folivores disappear during the daylight hours, and the cause of the missing plant foliage goes undiagnosed. Thus, the extent of plant damage by this insect is largely unknown. Doucette and Smith (1926) concluded that plant feeding was only a secondary food habit, and these cockroaches fed mostly on decaying organic ­matter. Schwabe (1949) reported that in Hawaii P. indicus (as P. ­surinamensis) fed extensively on chicken feces. Natural Enemies. The natural enemies of these cockroaches have not been studied. In tropical environments, ants are very important predators, and determinants of cockroach distribution (Tarli et  al., 2014). However, Surinam cockroach has a positive association with some ants, though as a temporary ant nest commensal, not as a myrmecophile (Deleporte et al., 2002; Moretti et al., 2011). Life Cycle and Description. These species are the only important members of the genus. In Florida, Surinam cockroach is normally found in wooded areas, both dry and wet. Both undisturbed natural plant communities and highly disturbed (suburban) communities are inhabited. Wooded areas provide a favored microenvironment, beneath leaf litter, and these insects not only burrow into loose soil, but can also be found inhabiting rodent burrows. Grandcolas et al. (1996) noted that in Africa P. surinamensis is found only in association with humans; however, Atkinson et  al. (1991) reported that in Florida it is found in natural communities as well as in disturbed areas. Surinam cockroach also benefits from association with others of the same species (so-called “group effect”). Aggregation of members of this species results in synchronized development and faster growth rates (Bell et al., 2007).

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“Surinam cockroach” formerly was believed to consist of two forms of the same species: the bisexual and the parthenogenetic forms. The “Surinam cockroach” of Southeast Asia and Hawaii usually are bisexual, though the parthenogenetic form is also found in Asia. In contrast, the “Surinam cockroach” in the Americas are entirely parthenogenetic; males are nearly nonexistent. Rather than treating them as different forms, Roth (1967) separated them into two species; thus, the parthenogenetic roaches are P. surinamensis whereas the bisexual forms are P. indicus. Consistent with the division of the “Surinam cockroach” species complex into different species is the fact that males from Hawaii cannot fertilize females from Florida, though they can fertilize females from Indonesia. Egg. The females of these cockroaches carry an egg case, or ootheca, until egg hatch occurs. The egg case is 12–15 mm long, light-colored, and crescent-shaped. There normally are about 25–35 eggs per egg case. The first ootheca is normally produced about 1  week after adulthood is attained. Eggs require about 35 days for hatch to occur when held at 18–24°C. Females require 50–80  days between broods. The adult female typically produce three broods annually, but the range is one to five. On average, P. indica produces more eggs than P. surinamensis, though cockroaches from different geographic locations vary in their reproductive capacity (Roth, 1974).

FIG. 4.3  Adult Surinam cockroach. (Photo by L. Buss.)

Adult. The adults are 18–24 mm long. The pronotum is smooth, with minute, widely placed pits, and dark brown to black in color. The forewings are light brown, and largely transparent. In females, the wings nearly cover the abdomen or extend slightly beyond the tip of the abdomen. In males, the wings are shorter, not nearly covering the abdomen (often several abdominal segments are visible), but variable in length. In Hawaii, where P.  indicus is found, females are more abundant than males, but males are readily collected. In contrast, in mainland USA, only females are found, and these are always P. surinamensis.

Damage

FIG. 4.2  Immature Surinam cockroach nymph. (Photo by L. Buss.)

Nymph. Initially, the young are translucent white with orange-brown mandibles, and measure about 4.5 mm long. Within a few hours, they darken to a mahogany brown color. The nymphal instars are quite dark, almost black, and darker than the adults. Nymphs have 8–10 molts over a 130–80 days period.

In the United States, Surinam cockroach is often associated with human habitation, so it is considered to be peridomestic, but it rarely enters buildings. Thus, it is similar to Asian cockroach (Blatella asahinai Mizukubo) in that it is associated with humans but does not actually take up residence in buildings. Both Surinam and Indian cockroach are (in part) plant feeders, though neither has been much studied with respect to plant damage. Again, this is similar to Asian cockroach, a similarly omnivorous diet that includes living plant material. These cockroaches, when abundant, should be considered part of the difficult-to-diagnose nocturnal plant feeders, along with slugs, snails, millipedes, cutworms, and other poorly known pests. Indian cockroach and Surinam cockroach are intermediate hosts of the nematodes Oxyspirura mansoni (Cobbold) and O. parvorum Sweet, which cause tropical eyeworm in chickens and other domestic fowl. Tropical eyeworm is widespread in the southeastern USA, where Surinam cockroach is found, and may also be implicated in transmission to wild birds such as northern bobwhite, Colinus virginianus (Linnaeus) (Galliformes: Odontophoridae).

Order Blattodea––Cockroaches and Termites Chapter | 4  59

Management The typical management practices for cockroach control do not apply to Surinam and Indian cockroaches, as residual sprays in and around the perimeter of buildings are not effective for an insect that dwells mostly in mulched areas and leaf litter. Thus, elimination of harborage should target mulch and leaf litter. Baits and other insecticide formulations scattered around the yard and garden may also be effective for suppression of these species. Doucette and Smith (1926) noted that sweet (sugar-based) baits were attractive to Surinam cockroach. Baits and other products labeled for cockroach control often are not registered for use around vegetables, so the label should be consulted before use.

FAMILY RHINOTERMITIDAE Subterranean Termites

Coptotermes spp. and Reticulitermes spp. (Blattodea: Rhinotermitidae)

Natural History Distribution. Termites occur widely, though in North America they are most abundant in the eastern USA, especially the southeastern region, the tropical islands (Hawaii, Puerto Rico), and also along the West Coast. They are less common in the western Great Plains, desert Southwest, and Rocky Mountain regions. They also are uncommon in most of Canada and Alaska. Termites in North America feed mostly on relatively dry wood, but some feed on living trees and other cellulose-containing material. The most important termites that feed on living plants are found in the tropics, not in North America, but some Reticulitermes spp. and Coptotermes spp. damage living plants in the United States. Host Plants. Termites feed on cellulose-containing material, normally on wood, but some feed on grass, or even on cow dung (which is rich in partially digested plant material). Damage to growing plants by termites is poorly known in the United States, though it is recognized as a significant issue in tropical countries. Formosan subterranean termite, Coptotermes formosanus Shiraki, is the principal problem species attacking living trees. For example, in a survey conducted at one site in New Orleans, Louisiana, Osbrink et  al. (1999) reported that 32.7% of loblolly pine, Pinus taeda (Pinaceae), 12.5% of cedar, Juniperus spp. (Pinales), 8.2% of live oak, Quercus virginiana (Fagaceae), and 4.6% of bald cyprus, Taxodium distichum (Pinales) contained Formosan subterranean termites. Other tree species also were affected, but the sample size was too small to draw firm conclusions. Native termites, Reticulitermes spp., were also found attacking trees at this site, but at a low level of incidence. With both species of termites, external signs of infestation

are rare. Many times it is only following wind damage, which causes splitting of the tree trunks, that termite infestations of large trees are detected. In South Carolina, Chambers et al. (1988) reported 17 species of trees affected by C. formosanus. Chouvenc and Foley (2018) reported tree damage following a survey of slash pine, Pinus elliottii (Pinaceae) in southern Florida, though the termite species was Coptotermes gestroi (Wasman). Also in Florida, citrus groves are damaged by Reticulitermes flavipes (Koller) and R. virginicus (Banks), often in new groves established on cleared pine woodland where remnant termite populations attacked fairly young trees (Stansly et  al., 2001). In South America, several species, mostly in the family Rhinotermitidae, are known to damage fruit, palm, and eucalyptus trees (Constantino, 2002). The most comprehensive review of plant damage in the United States was conducted in Hawaii by Lai et  al. (1983), who reported damage by C. formosanus in Hawaii provides insight into the potential for plant damage elsewhere. Not surprisingly, forest and fruit trees were attacked by C. formosanus, including mango, Mangifera indica (Anacardiaceae); Norfolk Island pine, Araucaria heterophylla (Aurocariaceae); avocado, Persea americana (Lauraceae), mahogany, Swietenia mahogoni (Meliaceae); banyan, Ficus sp. (Moraceae); coconut, Cocos nucifera (Palmae); sapodilla, Manilkara zapota (Sapotaceae); banana, Musa sp. (Musaceae); and lemon, Citrus limon (Rutaceae). However, some ornamental plants also were damaged. Examples of ornamentals damaged include lesser periwinkle, Vinca minor (Apocyanceae); Singapore plumeria, Plumeria obtusa (Apocyanceae); golden rain tree, Cassia fistula (Leguminosae); poinciana, Delonix regia (Leguminosae); hibiscus, Hibiscus spp. (Malvaceae); rose, Rosa sp. (Rosaceae); and lantana, Lantana camara (Verbenaceae). Among annual food crops reported to be damaged in Hawaii were cabbage, radish, beet, castor bean, sugar cane, and corn (Lai et al., 1983). In China, C. formosanus damaged root crops and peanuts, and in Jamaica Heterotermes tenuis (Hagen) damaged potatoes (Sands, 1973). In Africa, yam, cassava, and sugar cane were damaged (Wood et al., 1980; Mora et al., 1996). Mill (1992) provided a long list of crop plants affected by termites in Brazil. Many were the same as those reported in Hawaii, but also included rice, soybean, cotton, peanuts, cassava, sweet corn, and pasture. In Florida, potato tubers, and stems of tomato, eggplant, pepper, and broccoli are sometimes damaged, mostly after the land is cleared of trees and wood debris (Capinera, Unpublished). Natural Enemies. Termites are fed upon by a large number of predators, both invertebrate and vertebrate. The attack occurs most commonly when alate termites make dispersal flights, and when workers forage outside the nest. Birds can be quite attracted to such flights. Most predation

60  Handbook of Vegetable Pests

occurs opportunistically, and it is likely that all orders of entomophagous insects feed on termites. The most important natural enemies of termites are ants, and a few ant species specialize on termite predation, particularly ants in the subfamilies Ponerinae and Mrymicinae. Termites are not defenseless, of course, and Coptotermes are thought to be more resistant to predation than Reticulitermes due to the higher proportion of soldiers, and aggressive responses to disturbance, in Coptotermes. Insect parasitoids seem not to be very important for the biological suppression of termites. A few flies (e.g., Phoridae) and wasps (e.g., Bethylidae) parasitize termites. Several entomopathogenic organisms (viruses, bacteria, fungi, nematodes) are known to affect termites, but mostly in laboratory cultures. The more promising pathogens are fungi and nematodes, but the results of field evaluations have not been consistent. Beauveria bassiana and Metarhizium anisopliae fungi are best studied and are used when incorporated into baiting systems, but their incidence in natural systems is low. Biological control of termites is reviewed by Culliney and Grace (2000). Life Cycle and Description. Termites were formerly considered to be a separate order, Isoptera. However, they are now considered to be an especially social group of cockroaches and considered to be a suborder or infraorder in the order Blattodea. There are several families of termites, of which the most important are Kalotermitidae, Rhinotermitidae, and Termitidae. They also can be classified based on their habitat and feeding preference: drywood, dampwood, and subterranean. Subterranean termites such as Coptotermes and Reticulitermes (both found in the family Rhinotermitidae) live in diffuse nests in the soil and share many similarities in colony development and behavior. They may come to the surface to find ­cellulose-containing material, on which they feed, but they also maintain contact with the soil via tunnels and tubes constructed of soil. In North America, the termites that feed on living plants are nearly always subterranean termites. The powderpost and drywood termites tolerate the dry conditions of dead wood, and are not associated with soil; they also are of no importance as plant pests in North America. Termites in the family Termitidae also may be subterranean, and can be quite damaging to crops elsewhere, but not in North America (Wood, 1996). Termite development is classified as hemimetabolous, which usually means that it consists of eggs, immatures (called nymphs) that resemble adults but lack fully formed wings, and adults. There is no pupal stage in these insects. Development of termites are somewhat modified from “typical” hemimetabolous insects, though this is not unusual; many hemimetabolous insects vary a bit from “typical” development. The terminology used to describe termite development is both variable and confusing (Lainé and Wright, 2003).

Termites also differ in body form, depending on their function in the colony. Termite colonies contain three primary castes: the reproductives (king, queen, alates, alate nymphs, and supplementary reproductives), soldiers, and workers. Nymphs have the potential to develop into any caste, but their eventual form is determined by hormones and pheromones secreted by the reproductives and soldiers. Wings are present only in the reproductive caste. Soldiers have a large, dark, elongated head that is modified for defense, usually by bearing large mandibles. The workers normally are the most numerous members of the colony; they gather food and care for the colony. For most termites, there is only one queen per colony, and she bears a greatly enlarged abdomen. However, each year a large number of winged reproductives are produced. The new reproductives swarm and leave the colony, seeking to colonize distant resources and start their own colony. Annual dispersal flights generally occur following periods of rain or when the rainy season commences, although smaller flights can occur at other times. Depending on the species, flight can occur during the daytime or at night. Before flights, alate (winged) adults congregate in galleries near the soil surface or in the wood above-ground.

FIG. 4.4  Eggs and workers of Reticulitermes hageni. (Photo by L. Buss.)

Egg. Shortly after taking flight, adults mate and begin construction of a new colony. About 1–2  weeks after mating, egg production commences. Females produce numerous small elongate-spherical eggs typically measuring about 0.5–0.7 mm long and about 0.25–0.3 mm wide. In the spring, eggs hatch in from 15  days in the spring whereas in the fall it requires longer, perhaps 60  days. Female Reticulitermes can lay many eggs over the course of several years, estimated to be in the hundreds of thousands. However, they begin slowly, producing only 25–30 eggs during the first year (Howard et  al., 1981). In the well-developed nest, eggs are often found in clumps of 500–1000.

Order Blattodea––Cockroaches and Termites Chapter | 4  61

FIG.  4.5  Workers and soldiers of Reticulitermes hageni. (Photo by J. Castner.)

FIG. 4.7  Queen of Reticulitermes hageni. (Photo by L. Buss.)

Immature. The number of immature forms (instars) is variable, with the first two being quite helpless and fed initially (in the age of the colony) by the adults, but later by the workers. The two initial instars are called larvae and lack any sign of wings. These are followed by the nymphal instars, which may or may not have wing buds. Nymphs that are destined to be workers lack wing buds, whereas those destined to be swarmers (reproductives) have long wing buds, and eventually (after the sixth instar) develop into sexual adults or supplementary reproductives. The first instars of Reticulitermes spp. are 1–2 mm long and require 7–17 days for development. The second instar takes 10–18  days. After the second molt the third instars track to the sexual or neutral paths of development. Termite instars can be differentiated by body size, head capsule width, or the number of antennal segments. In Coptotermes formosanus, for example, the mean head capsule widths are 0.49, 0.65, 0.82, 0.87, 0.95, and 1.10 mm for instars 1–6, respectively (King and Spink, 1974). Mean body lengths during these instars are 1.7, 2.6, 3.2, 3.4, 3.6, and 5.1 mm, respectively.

Adult. Workers are 4–7 mm long, wingless, and lack compound eyes. The number of instars is variable. Worker development requires about a year, and they can live for several years. Workers are the most numerous form in the colony, and they are adapted for procurement and digestion of wood. Workers can differentiate into soldiers from the fourth to the seventh molt. Reticulitermes flavipes soldiers are 6–7 mm long whereas R. virginicus soldiers are 4.5–5 mm long. Soldiers resemble workers but have biting mandibles for defense. They are dependent on the workers for food. Soldiers are formed in two stages from workers; the first is unpigmented and unsclerotized but the second is sclerotized. The alates of Reticulitermes flavipes and R. virginicus are dark brown. Alates of R. flavipes are usually larger (approximately 10–11 mm long including wings) than those of R. virginicus (approximately 8–10 mm long). The wings of alate forms of Reticulitermes species have two thickened veins along the entire leading edge of the wings. Although Formosan termite also has two thickened wing veins, the Reticulitermes species lack the small hairs that are found on the wings of C. formosanus Shiraki. Formosan termite also tends to be orange-brown rather than dark brown, and is 10–15 mm long. The antennae are bead-like or thread-like, and of medium length. Termites are soft bodied, and usually lightcolored. The front wings and hind wings are nearly the same in size and shape, and are held flat over the abdomen when at rest. The abdomen and thorax join broadly; there is not a pronounced constriction. Termites are commonly mistaken for ants, but the elbowed shape of the antennae, the darker coloration (this characteristic is not always reliable), shorter length and different shape of the hind wings, and the marked constriction of the first abdominal segments (the petiole) are characteristics that are used to distinguish ants from termites. After a brief flight, alate adults drop to the ground and shed their wings. Females begin to search for potential

FIG.  4.6  Adult winged Cryptotermes subterranean termite. (Photo by L. Buss.)

62  Handbook of Vegetable Pests

n­ esting sites such as moist crevices with wood, and males follow closely behind. The pair forms a royal chamber in a moist site near food (usually wood) and begin laying eggs, thus starting the life cycle of a new subterranean termite colony. As noted previously, some individuals that hatch from eggs (called larvae) molt into workers. Interestingly, although the reproductives initially can feed, their ability to feed is lost after the appearance of workers, which take over the activity of feeding the reproductives. Some workers molt into the soldier caste or eventually develop into new winged adults. Some workers or nymphs are capable of becoming supplementary reproductives and take over the role of queen or king if and when the king or queen dies or is separated from the colony. It may take several years for a single pair of alate adults to form a mature colony that produces new alate adults.

Damage Termites are known primarily for their damage to wood structures. Food consumed by termites is relatively low in nitrogen content, though containing a high proportion of plant polysaccharides. A combination of termite digestive enzymes and enzymes produced by endosymbiotic gut protozoa and bacteria are responsible for digestion. The net result is efficient digestion not only of soluble carbohydrates—materials that most insect can digest, but also of cellulose, hemicellulose, and lignin—materials that are difficult for most insects to digest. In addition, some termites will damage living plants. The genus Coptotermes is notorious for its habit of

c­ olonizing living trees and hollowing out the heartwood. This can cause the core of the trunk to be “piped” and replaced with nest material and soil, without the tree showing external signs of their presence. However, Coptotermes spp. and Reticulitermes spp. also attack other cellulose-rich plant materials, including the aboveground (stems) and below-ground (tubers) portions of vegetable crops. This often occurs when land containing trees and termites is cleared of timber, and then planted to crops.

Management The typical management practices for termite control are not designed for crop protection. Population assessment in both structural and crop environments can be accomplished in the same manner, however, by burial of wood-based baits below-ground. In most cases, termite control involves the creation of a chemical barrier between termites and a susceptible structure. However, injection of insecticides into trees is sometimes recommended for preservation of living trees. Also, the development of termite baits plus slow-acting toxicants has been shown to provide long-term suppression of some crops (Stansly et  al., 2001) though the need for suppression of termites on agricultural land is infrequent, at least in North America. Applications of liquid and granular insecticides to the soil, which often are made routinely for suppression of soil-dwelling crop-feeding insects, will alleviate termite problems, but if suitable food (wood) remains, termites will recover or reinvade.

Chapter 5

Order Coleoptera—Beetles, White Grubs, and Wireworms

FAMILY CARABIDAE—GROUND BEETLES Seedcorn Beetle

Stenolophus comma (Fabricius) Stenolophus leconti (Chaudoir)

Slender Seedcorn Beetle

Clivina impressifrons (LeConte) (Coleoptera: Carabidae)

Natural History Distribution. The seedcorn and slender seedcorn beetles are commonly found in the United States in the midwest and northeast and occur west through the Great Plains and south to South Carolina. In Canada, they are present in eastern provinces, west to at least Ontario. They are native insects. The early accounts of the Stenolophus spp. are unreliable because the species were often confused; despite early reports suggesting that S. lecontei was the most important of the seedcorn beetles, it now appears that S. comma is the most serious pest. Host Plants. Carabid beetles are known principally as predators of other insects. Animal matter is also the preferred food source of the seedcorn and slender seedcorn beetles. Eggs, larvae, and probably pupae of soil-dwelling insects are readily consumed. For example, Wyman et  al. (1976) documented the reduction of cabbage maggot, Delia radicum (Linnaeus), populations when seedcorn beetles were numerous in crucifer plots. As suggested by the common names, on occasion these beetles will injure germinating seeds, particularly corn seed. However, beetles also feed on weed seeds. Adults of S. comma feed readily on black nightshade, Solanum nigrum; crabgrass, Digitaria sanguinalis; foxtail, Setaria glauca; lambsquarters, Chenopodium album; and purslane, Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00005-4 © 2020 Elsevier Inc. All rights reserved.

Portulaca oleraceae, and to a lesser degree on other seeds (Pausch, 1979). Natural Enemies. The natural enemies of these insects are poorly known. The mite Ovacarus clivinae (Stannard and Vaishampayan) develops inside slender seedcorn beetle, where it is associated with the reproductive system (Stannard Jr. and Vaishampayan, 1971). Similarly, three mites [Eutarsopolipus elzingai Husband and E. brevichelus Husband & Husband (both Acari: Podapolipidae; Crotalomorpha camini Linquist & Krantz (Acari: Crotalopmorphidae))] were identified in midwestern states infesting Stenolophus spp. These mites are subelytral parasites that, although occurring widely, are infrequent in these hosts (Husband & Husband, 2004). Life Cycle and Description. These beetles are quite similar in biology. Adults overwinter and can be found throughout the year. Maximum above-ground activity in South Dakota was observed to occur during early summer, followed by almost total absence in late summer, and then reappearance in the autumn. Eggs are abundant in the spring, larvae and then pupae occur in late spring and early summer, and new adults are produced by mid-summer. Only a single generation is produced annually. Development of the insects from egg to adult requires about 50 days. Egg. The eggs are laid singly in horizontal tunnels dug about 2 cm deep in the soil. The eggs are white and oval in shape, measuring about 1 mm long and 0.5 mm wide. Mean duration of the egg stage is about 5.4 days (range of 3–12 days) when reared at about 22°C. Larva. The larvae are yellowish, with a dark-brown head and prothoracic plate. The larvae display three instars, and all instars can be found during the summer months. They are normally found in burrows in the soil at depths of 2–3 cm. Mean duration (range) is reported 10.5 (4–19), 9.3 63

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(5–16), and 12.8 (9–21) days for instars 1–3, respectively, when reared at about 22°C. Pupa. Pupation occurs within the soil in small cells, each measuring about 13 mm in length and 5 mm in width. The pupa is dark brown or black. Mean duration of the pupal stage is 9.4 days, with a range of 7–13 days, when reared at 22°C.

a­ttacked in this manner may perish. However, if only a small portion of the seed is consumed, the seedling may recover and grow normally. Stenolophus spp. is reported to be more abundant in tilled plots, relative to less disturbed (no-till) plots, which may explain why they are more often plant pests than most other carabids (Davis et al., 2009).

Management The adult populations can be sampled with pitfall and blacklight traps. Rows of corn in which plants are missing should be examined carefully because beetles can often be found in association with the poorly germinating seeds. The populations of seed corn beetles can be controlled by the application of insecticides to the seed, or in liquid or granular form to the soil at planting or shortly after planting (Daniels, 1977). Resistance to insecticides is evident in some populations (Sechriest et al., 1971).

FAMILY CHRYSOMELIDAE, SUBFAMILY ALTICINAE—FLEA BEETLES FIG. 5.1  Adult seedcorn beetle. (Drawing by USDA.)

Adult. The adults are small oblong beetles measuring 5–8 mm long. Adults of the Stenolophus spp. are dark below and yellowish brown or reddish brown above, but with the elytra blackened except for the margin. Slender seedcorn beetle differs in that it is entirely reddish brown and has especially enlarged and flattened front legs suitable for tunneling through soil. Adults are commonly found in moist soil, with a high content of organic matter. In the late autumn, beetles burrow beneath soil, rocks, and logs, where they remain in a small cell until the soil warms to about 5°C in the spring. Adults may fly during the daylight hours in the spring but are more active in the evening during the summer months. Apparently, copulation occurs belowground in small chambers constructed by the adults during the spring. Oviposition commences within 14–15 days of emergence, and continues for 5–10 weeks. The biology of Stenolophus comma was described by Kirk (1975); both S. comma and S. lecontei were considered by Pausch (1979). Slender seedcorn beetle was treated by Phillips (1909) and Pausch and Pausch (1980). These species were included in the keys to adult beetles by Downie and Arnett Jr. (1996).

Damage Damage to germinating seeds caused by seedcorn beetles occurs principally during cool and wet weather in spring. Rarely an ungerminated seed is attacked. The contents of the seed are often consumed, leaving only the hull or seed coat. The result of the attack is often a poor stand, as ­seedlings

Cabbage Flea Beetle

Phyllotreta albionica (LeConte) (Coleoptera: Chrysomelidae)

Natural History Distribution. This native flea beetle is widely distributed in the western United States and Canada. It is found only as far east as Saskatchewan, Colorado, and New Mexico. It is occasionally reported from the eastern United States or Canada, but this appears to be due to misidentification. Host Plants. Cabbage flea beetle feeds principally on cruciferous plants. Vegetable crops attacked by this beetle include broccoli, Brussels sprouts, cabbage, cauliflower, kale, radish, rutabaga and turnip, but radish and turnip are most preferred. It is also reported to attack alfalfa, sugar beet, and tomato, but these events are unusual, or are cases of misidentification. Cabbage flea beetle feeds readily on cruciferous weeds. Natural Enemies. Natural enemies are not well known, but Microctonus epitricis (Viereck) (Hymenoptera: Braconidae) has been reared from cabbage flea beetle. Life Cycle and Description. There is a single generation of this insect in Canada, but its biology has not been studied in detail elsewhere. Adults overwinter in leaf litter along fencerows and in woodlots. They become active in March and April and begin feeding on crucifers. Mating soon ensues, and egg laying commences after a preoviposition period of about 5–10 days. Adults live for about 3 months, continually feeding and depositing more eggs. Egg. The eggs are deposited beneath the host plants in the soil at a depth of about 2.5–5.0 cm. Clusters of 15–20

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eggs are common, and females deposit eggs over a 3-week period. Fecundity is estimated at about 60 eggs per female, but this is likely an underestimate owing to laboratory rearing conditions. Eggs hatch in 15–20 days. Larva. Larvae are whitish and are found feeding on the roots at depths of 5–15 cm. Larval development requires about 4 weeks and can be found during May–July. Pupa. As larvae are about to pupate, they characteristically move closer to the soil surface and create a small pupal chamber in the soil. The nonfeeding prepupal period lasts about 10–12 days and is followed by a pupal period of about 11 days.

FIG. 5.2  Cabbage flea beetle. (Drawing by USDA.)

Adult. The adults of the summer generation emerge from the soil in August and begin feeding. Cabbage flea beetle is a shining blackish species, measuring 1.5–1.9 mm long. This beetle is easily confused with crucifer flea beetle, Phyllotreta cruciferae (Goeze), a more damaging species. However, cabbage flea beetle has a bronze luster, whereas crucifer flea beetle has a blue luster. Also, the fifth antennal segment is elongate in the female and elongate and broad in the male cabbage flea beetle. The legs of cabbage flea beetle are brown except for the femora, which are black. The hind femora are enlarged. The antennae are black basally and reddish black toward the tip. The summer adults feed for about 6 weeks before seeking overwintering shelter. The biology of this flea beetle was given by Chittenden (1927), Burgess (1977, 1982), and Campbell et al. (1989).

Damage Damage is caused principally by the adults in the form of small holes in the leaf surface. These beetles are rarely abundant enough to damage anything but seedling plants. The larvae feed on the roots of crucifers, but they are not usually considered a problem, probably in view of their low numbers.

Management Management of this insect is similar to other crucifer-­ feeding flea beetles. Management has been discussed in detail in the section on crucifer flea beetle.

Corn Flea Beetle

Chaetocnema pulicaria Melsheimer

Toothed Flea Beetle

Chaetocnema denticulata (Illiger) (Coleoptera: Chrysomelidae)

Natural History Distribution. Corn flea beetle and toothed flea beetle are found primarily in the northeastern and midwestern United States, but their range extends as far west as the Rocky Mountains. The northern limit of these native species seems to be Massachusetts, New York, and southern Ontario. Survival is affected by winter weather, and damage resulting from flea beetle feeding is noticed most commonly following mild winters. Host Plants. The adults and larvae of corn flea beetle and toothed flea beetle feed and develop on a wide variety of cultivated and wild grasses and sedges. In addition to corn, other crops attacked include barley, chufa, oats, orchard grass, wheat, and timothy grass. Examples of weeds suitable for these flea beetles are crabgrass, Digitaria spp.; barnyardgrass, Echinochloa crusgalli; witchgrass, Panicum capillare; straw-colored sedge, Cyperus strigosus; and yellow bristlegrass, Setaria glauca. Natural Enemies. The natural enemies of corn flea beetle and toothed flea beetle are poorly known. Several investigators of corn flea beetle biology noted an absence of parasitoids. A wasp was found attacking toothed flea beetle eggs in Virginia and identified as Patasson pullicrura (Girault) (Hymenoptera: Mymaridae) (Poos, 1955). Life Cycle and Description. Corn flea beetle and toothed flea beetle are very similar in appearance and biology. Toothed flea beetle is much less common in corn, and so is less well studied. Except where noted below, their biology is believed to be virtually the same. Two generations of these flea beetles are reported annually on corn in Connecticut and Virginia, with adults from the spring generation occurring in late June and early July, and those from the second brood appearing in mid-August. Poos (1955) suggested that in Virginia, corn flea beetle might complete a generation on wild grasses and sedges before moving to corn, leaving some doubt about the actual number of generations. The time required for completion of a generation (egg to adult) is estimated to average 30 days. Overwintering of corn flea beetle and toothed flea beetle occurs in the adult stage in soil. Beetles are most commonly recovered from the upper 2–3 cm of grass sod planted in the

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vicinity of corn. Bluegrass, Poa sp., and other grasses have been found to be suitable habitats. The beetles become active when the soil surface warms to about 15–20°C. Egg. Poos (1955) speculated that eggs were deposited at the base of corn or wild grasses in the manner of other flea beetles. However, he was unable to locate eggs in the field and obtained them by confining beetles in the laboratory. The eggs of corn flea beetle are ovoid and white in color. They measure about 0.2 mm wide and 0.4 mm long. Development time of the egg stage averages 4–6 days, with a range of 2–10 days, depending principally on temperature. The eggs of toothed flea beetle are slightly larger, measuring about 0.6 mm long, and are pale yellow. Larva. Poos found that larvae were also difficult to locate, but a few could be found in the soil surrounding corn and other grasses. Larval development requires about 16 days (range 10–23 days), not including an additional 2 days (range 1–5 days) as a prepupa. Larvae attain a maximum length of about 4.5 mm before pupation. The pupal stage requires about 5 days (range 3–7 days).

FIG. 5.3  Adult corn flea beetle. (Drawing by USDA.)

Adult. The adult corn flea beetle is shiny black with a slight greenish, bluish or bronze luster. They measure about 1.5–1.8 mm long. Elytra are marked with rows of closely spaced punctures. The adult toothed flea beetle is bronze in color, and measures 2.3–2.5 mm long. The elytra of this beetle are also marked with rows of punctures, but the punctures are not spaced closely. The head also bears punctures, a feature lacking from corn flea beetle. Although these two flea beetle species are not especially difficult to distinguish, there are several other flea beetle species that may occur in corn, usually in small numbers. The biology of corn flea beetle and toothed flea beetle were given by Poos (1936), Elliott and Poos (1940), and Poos (1955).

Damage The adults skeletonize leaves of seedling corn in the spring, sometimes completely defoliating fields. They feed on the lower surface of foliage in narrow linear strips, usually restricting their feeding to the first three leaves. The principal form of injury, however, is through the transmission of Stewart’s bacterial wilt, a disease caused by the bacterium Pantoea (formerly Erwinia) stewartii. Stewart’s wilt is transmitted when beetles feed on corn and defecate in or near the feeding site. The bacterium, which is harbored in the digestive tract of the beetles, thus gains entry to the plant through the wound. Infected beetles remain infected and are capable of transmitting the disease for the duration of their life. The disease is harbored by the beetle during the winter months, but the bacteria also occur in wild grasses, including many species that do not express symptoms of infection (Poos, 1939). The studies conducted in Connecticut demonstrated that the incidence of beetles containing bacteria generally increased from about 40% to 70% during the season, although there was some evidence of a mid-season reduction in disease levels (Heichel et al., 1977). Esker et al. (2002) noted two peaks in beetle abundance in Iowa, one in June–July and the other in August–September. The incidence of disease in beetles is quite variable. In Iowa, the incidence varied from as low as 4% to as high as 86%, with the highest levels of infection tending to occur in August (Esker and Nutter Jr., 2003). In Illinois, the range of infection was reported to vary from 2% to 99% (Cook et al., 2005). Although both species may exhibit high levels of infection, corn flea beetle is much more abundant and therefore is much more important as a vector. Elliott and Poos (1940) provided a long list of insects that were found contaminated with P. stewartii. Although the list is lengthy, incidence of infection was slight except for the aforementioned flea beetles. The symptoms of Stewart’s wilt infection in sweet corn include wilting, pale green linear streaking, stunting, and death. Surviving plants may tassel prematurely and produce deformed ears. About 3 weeks are required before symptoms of infection are evident. Early season infection, or wilt phase of the disease, is most damaging to sweet corn. Field corn varieties are generally resistant to early infection but suffer from late-season infection. Stewart’s wilt is a limiting factor in sweet corn production in the northeastern United States in recent years.

Management Sampling. Hoffmann et al. (1995) studied the distribution of beetles between and within plants. Beetles tend to aggregate; about 50% are found on the uppermost of the fully emerged leaf. A system to aid prediction of the severity of Stewart’s wilt, based on winter temperature, was developed in Pennsylvania (Castor et al., 1975). Mean monthly temperatures during December–February, exceeding 3°C, favor adult overwintering survival. Beetle numbers can also

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be monitored during the growing season. Adams and Los (1986) found that yellow sticky traps hung close to the soil were most efficient in capturing corn flea beetles. Beetles can be counted on plants, and a threshold of six beetles per 100 plants is sometimes used to initiate control (Adams and Los, 1986). In Iowa, Esker et al. (2003) determined that yellow sticky cards positioned vertically at 0.3 m above the soil captured more beetles than other heights tested. Cook et al. (2005) suggested that two corn flea beetles per trap per day be used as a threshold for insecticide treatment to protect against damage by beetles and Stewart’s wilt, though higher numbers also have been suggested. Insecticides. Systemic insecticides are commonly applied in-furrow at planting, and after emergence, to protect the young corn from feeding injury and transmission of Stewart’s wilt (Ayers et al., 1979; Munkvold et al., 1996). For example, Ayers et al. (1979) reported that Stewart’s wilt infection was reduced from about 34% in untreated corn plants to only 2%–4% when effective insecticide was used. The insecticide carbofuran seems to interfere directly with the bacterium, imparting resistance to the plant (Sands et  al., 1979). When corn plants were grown from seeds treated with the systemic insecticide imidacloprid, the treatments reduced the number of feeding scars, the number of leaves with disease symptoms, and the number of plants infected with P. stewartii as compared to untreated plants (Munkvold et al., 1996; Kuhar et al., 2002), though Kuhar et al. (2002) observed that genetic resistance was more effective than treatment with insecticide. Cultural Practices. Sweet corn varieties differ in their susceptibility to Stewart’s wilt. Some varieties are quite resistant, but if high overwintering populations of corn flea beetles are forecast, insecticide treatment is also advised (Ayers et al., 1979). Early maturing varieties tend to be more susceptible to injury. The availability of transgenic Bt corn may also affect corn flea beetle abundance. Bhatti et  al. (2005) observed fewer beetles in Bt corn plots than non-Bt plots, but attributed this to an indirect effect, postulating that the flea beetles were attracted to the non-Bt plots by the damage caused by western corn rootworm, Diabrotica virgifera.

Crucifer Flea Beetle

Phyllotreta cruciferae (Goeze) Coleoptera: Chrysomelidae

Natural History Distribution. Crucifer flea beetle is also found in Europe, Africa, and Asia, as well as in North America. Crucifer flea beetle was first found in North America in 1921, in British Columbia. It moved steadily eastward and was recovered from Canada’s Prairie Provinces in the 1940s, and Ontario and Quebec by 1954. A second introduction on the east coast was evident as this insect was found in Pennsylvania in 1943 and was well established in Delaware by 1951

(Westdal and Romanow, 1972). It is now widely distributed in southern Canada and most of the northern United States. Crucifer flea beetle is common in the prairie and other open environments and is rare in forested areas (Burgess, 1982). The expansion of acreage planted to rape seed (canola) both in Europe and North America has facilitated the spread of this insect and enhanced its status as a crop pest. Host Plants. Crucifer flea beetle feeds principally on plants in the family Cruciferae, though other plant families containing mustard oils (glucosinolates) (also found in families Capparidaceae, Tropaeolaceae, Limnanthaceae) have been reported to be attacked (Feeny et al., 1970). Broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, horseradish, kohlrabi, radish, rutabaga, and turnip are the vegetable crops commonly damaged by this flea beetle. Preference among crucifer crops is sometimes reported, but the preferences are rarely consistent owing to changes in leaf age and leaf type (Palaniswamy and Lamb, 1992). Although glucosinolates are attractive, high concentrations or differing compositions of glucosinolates may affect plant acceptance (Soroka and Grenkow, 2013). Crucifer flea beetle also damages rape, and sweet alyssum, as well as several cruciferous weeds such as tansymustard, Descurainia sp.; wild mustard, Brassica kaber; stinkweed, Thlasi arvense; pepperweed, Lepidium densiflorum; yellow rocket, Barbarea vulgaris; and hoary cress, Cardaria draba. Reports of this insect feeding on noncruciferous crop plants such as beets apparently stem from misidentification; these insects are easily confused with related species. Natural Enemies. There are few effective natural enemies of crucifer flea beetle in North America. A parasitoid, Microctonus vittatae Muesebeck (Hymenoptera: Braconidae), attacks adult P. cruciferae, placing about two-thirds of its eggs in the host’s head. Although several eggs may be deposited in each beetle, only a single wasp survives, emerging 16–19 days after parasitism (Wylie and Loan, 1984). The level of parasitism by M. vittatae may be 30%–50% (Wylie, 1982), but crucifer flea beetle is less preferred than striped flea beetle, Phyllophaga striolata (Fabricius), for oviposition (Wylie, 1984). Parasitized-flea beetles emerge earlier from overwintering sites than unparasitized beetles (Wylie, 1982). Parasitism by nematodes, particularly by the allantonematid Howardula sp., is generally low, and the nematodes are not very pathogenic. In Europe, several other nematodes attack crucifer flea beetle (Morris, 1987). General predators such as lacewings (Neuroptera: Chrysopidae), soft-wing flower beetles (Coleoptera: Melyridae), and big-eyed bugs (Hemiptera: Lygaeidae), and omnivores such as field crickets (Orthoptera: Gryllidae), are occasionally observed feeding on adults, but their impact is undetermined (Gerber and Osgood, 1975; Burgess, 1980; Burgess and Hinks, 1987). Life Cycle and Description. There is generally only one or possibly two generations annually, although development from egg to adult can be completed in just 7 weeks. Overwintering occurs as an adult in soil and leaf litter. Fence

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rows, windbreaks, and other forms of shelter are thought to favor overwintering sites, although this does not seem to be active dispersal to such sites. Sometimes large aggregations of overwintering beetles are found; densities as high as two million beetles per hectare have been observed overwintering in a grove of trees (Burgess and Spurr, 1984; Turnock et  al., 1987). Adults diapause under short day length (8 h) conditions, and readily survive 5 months of refrigeration. In Ontario, it was reported that peak emergence of overwintering adults occurred by early May (Kinoshita et  al., 1979), with summer generation adults most abundant in late June and then again in late July. The two summer generations overlap considerably, and could easily be a protracted single generation. These authors also reported only a single summer generation during a cool summer. In the northeastern United States, Andersen et al. (2005) reported two peaks in abundance, with the second due to a partial second generation.

Adult. The adult measures about 2.2 mm long and is metallic blue-black except for the tarsi and antennae, which may be partly amber. The elytra, and to a lesser degree the head and thorax, bear small punctures. The hind femora are enlarged. The adults disperse principally by jumping and are usually trapped within 20 cm of the soil surface (Vincent and Stewart, 1983). They will fly throughout the season, however, and in the earlier part of the year, they are more likely to be found flying at greater heights (1–2 m) (Lamb, 1983). The biology of crucifer flea beetle was given by Westdal and Romanow (1972), Burgess (1977), and Kinoshita et al. (1979). Rearing procedures were provided by Kinoshita et al. (1979). Evidence for a male aggregation pheromone is presented by Peng and Weiss (1992) and Peng et al. (1999); trap capture is enhanced by including host plant volatiles (Soroka et al., 2005).

Egg. The eggs are laid in the spring, singly or in groups of 3–4 in the soil, usually near the base of food plants. The temperature threshold for oviposition is 16.7°C. The egg measures 0.38–0.46 mm long and 0.18–0.25 mm wide, and is yellow in color. Eggs hatch after about 11–13 days.

Damage

Larva. There are three instars. Larvae are white except for a brown head and anal plate. Head capsule width measurements for the instars are 0.13, 0.17, and 0.26 mm, respectively. Body lengths are approximately 0.9, 4.5, and 6.7 mm, respectively. Development time is about 5, 3, and 4 days, respectively, at 25°C. Larvae feed on the root hairs of plants for 25–30 days at 20°C, and then form a small earthen pupal cell in the soil. The prepupal period, during which feeding ceases and the larval body shortens and thickens, lasts 3–6 days. Pupa. The pupa is white, measuring about 2.4 mm long. Pupation lasts 7–9 days, followed by the emergence of a whitish adult that darkens completely in about 2 days.

The adult injury is in the form of small holes in the foliage. They do not eat completely through the leaf but leave the lower epidermis intact. However, the remaining tissue soon dies, dries, and falls from the plant, producing a hole. When beetles are abundant, all leaf tissue may be riddled with holes, resulting in drying of adjacent tissue and death of emerging seedlings. Severe injury typically occurs in spring when the weather is hot and dry. Defoliation of young seedlings and small broccoli transplants in Manitoba at levels of  200 eggs. Females may produce eggs over a period of about 2 months. Moisture is an important criterion for egg deposition and hatching. Females seek moist areas for egg deposition, and lack of moisture retards hatching. Temperature also influences hatching, with embryonic development being completed in 6–8 days during warm weather, but 11–12 days during cool weather. Larva. The larvae are about 1 mm long and white when they first hatch from the egg. As they mature, the head and

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thoracic plate and to a lesser degree the anal plate, become brown, while the remainder of the body remains pale. The larva eventually attains a length of about 4–5 mm. The terminal abdominal segment bears an anal proleg. Larvae normally complete their development in about 20–25 days, but the larval development period may range from 13 to 45 days, depending on the weather. Pupa. The larvae pupate in the soil after preparing a small pupal chamber from soil particles. Pupation occurs near the food plant, close to the area of feeding. The pupa measures about 1.5–2.0 mm long and initially it is white, but eventually darkens. Pupal development requires about 6–10 days (range 3–22 days).

FIG. 5.10  Adult potato flea beetle. (Drawing by USDA.)

Adult. The adult potato flea beetle is fairly light in color after emerging. However, by the time it has dug to the soil surface, it is fully darkened and cannot readily be distinguished from individuals that have long been emerged, including those that have overwintered. Adults can be found feeding until the plants have been killed by frost. Beetles measure 1.5–2.0 mm in length and are black. The antennae and legs are orange, except that a portion of the fore and middle femora are black, and the hind femora are completely black. The hind femora are enlarged. The pronotum is marked with fine punctures, the features of which can be used to distinguish this insect from the closely related tuber flea beetle, Epitrix tuberis Gentner. The fine punctures in the center of the pronotum of potato flea beetle are separated by a distance exceeding the diameter of the punctures. In contrast, the punctures at the center of the pronotum of tuber flea beetle are relatively coarse and are separated by a distance less than the diameter of the punctures (Gentner, 1944). The pronotum of tuber flea beetle also has a deep transverse depression near the hind margin. Both the pronotum and elytra bear numerous fine hairs, but their density is less than the similar

eggplant flea beetle (see eggplant flea beetle for comparative description). Western potato flea beetle has been poorly studied, but its biology in Washington seems similar to that of potato flea beetle (Hanson, 1933). It seems to be two-brooded in Oregon. It is similar in appearance to potato flea beetle, but is bronze rather than black, and lacks the transverse depression on the prothorax. A good account of potato flea beetle was given by Johannsen (1913), based on the work conducted in Maine. However, a more comprehensive treatment of biology, including rearing methods, was given by Hoerner and Gillette (1928) from Colorado. This and most other life-history research from the western United States conducted before a systematic revision of the Epitrix flea beetles resulted in recognition of tuber flea beetle as a separate species (Gentner, 1944). Thus, some studies may contain mixed populations, and despite Gentner’s attempt to identify the true species involved in the previous research on “potato flea beetle,” the biology of potato flea beetle remains somewhat uncertain. The work of Hill and Tate (1942) in Nebraska is particularly suspect because tuber flea beetle appears to be present in this area. Beirne (1971) and Campbell et al. (1989) provided a useful perspective on potato flea beetle in Canada. Keys and other diagnostic aids for the Epitrix flea beetles of North America are available in Deczynski (2014), and Bienkowski and Orlova-Bienkowskaja (2016, 2017). Germain et  al. (2013) demonstrated the ability of RFLP-based diagnostic methods to distinguish among flea beetle species, so in the future, determinations should be more accurate.

Damage The damage by adult potato flea beetles is typical flea beetle injury. Small pits are eaten in either the upper or lower surface of the foliage. Although the beetles may not eat completely through the foliage, the remaining epidermis soon dries and dies. Dead leaf tissue eventually falls away, leaving a hole. Leaves on heavily infested plants may be riddled with holes, although the veins usually remain. Small plants may be killed. Once potato plants become established, however, they can withstand considerable flea beetle feeding. Senanayake et al. (1993a,b) estimated that densities of 100 beetles per plant would not depress potato yield of healthy plants. If potatoes were previously stressed by Colorado potato beetle feeding, then densities as low as 5–25 flea beetles per plant would depress yield. In eastern Canada, flea beetle populations regularly reduce potato yield by 15% unless corrective actions are taken. Damage by adults is greater under hot and dry conditions. The larvae are root feeders but occasionally cause considerable damage by feeding on potato tubers. By feeding on the tuber surface, larvae produce tracks or scars that materially damage the commercial value of the potato tubers.

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Occasionally they burrow directly into the tuber to a depth of up to 3 cm. Eventually, the pits are filled with corky material that blackens. Although tuber feeding is noted wherever potato flea beetle occurs, it is not as serious a problem as it is with tuber flea beetle. In the case of potato flea beetle, adult damage is the predominant form of damage. Potato flea beetle has been implicated in the transmission of several plant diseases. Tuber feeding also enhanced infection of tubers by Rhizoctonia and potato scab fungi (Hanson, 1933). Potato flea beetle also has been shown to be capable of transmitting ring rot bacteria (Christie et al., 1993), and probably many other diseases. Damage to plants by western potato flea beetle is similar to potato flea beetle damage, although much less frequent. Also, western potato flea beetle larvae are less prone to feed on the tubers of potatoes, feeding instead primarily on the fibrous roots.

Management Sampling. Senanayake and Holliday (1988) compared various sampling procedures for potato flea beetle monitoring. Use of a sweep net was efficient early in the season but declined as plants matured. Visual samples were not highly satisfactory because beetles were often feeding on the underside of foliage, and difficult to observe. Whole plant bag sampling was determined unwieldy for routine sampling despite its high degree of precision. Stewart and Thompson (1989) determined the spatial distribution of potato flea beetles on potato and recommended that 10–20 plants be sampled when beetle densities were four per plant or higher, but that sample size be increased to 50–60 plants when beetle densities were only one per plant. Because the beetles are small, mobile, and often feed on the underside of foliage, plant damage may be a better indicator of insect density than beetle counts. The relationship of feeding puncture abundance to yield has been studied, but owing to variability in plant response, the damage threshold remains uncertain (Howard et al., 1994). Insecticides. Foliar applications are made for the suppression of adults, usually 1–2 weeks after adults first appear. Commonly, several applications are necessary to protect young plants from overwintering beetles, and the first or spring generation adults, due to protracted emergence. Granular formulations are applied at planting or after if there is reason to expect damage by larvae. Systemic insecticides are applied at planting, using various formulations; although some systemic treatments provide excellent seedling protection against both adults and larvae, phytotoxicity is a potential hazard (Chalfant et al., 1979b). Insecticide resistance has been detectable since the 1950s (Kring, 1958), but populations remain manageable (Ritcey et al., 1982). In many areas, excellent control of flea beetles occurs as part of Colorado potato beetle chemical suppression. As more selective management techniques are developed for Colorado potato beetle, flea beetle problems may

be increased. Some organic methods of insect suppression (e.g., rotenone or kaolin clay) provide some benefits, but conventional insecticides may be more beneficial in terms of production. Cuthbertson (2015) provides a historical perspective on control of potato-infesting Epitrix spp. Cultural Practices. Glandular trichomes deter feeding by potato flea beetle (Tingey and Sinden, 1982), but this character is not yet incorporated into horticulturally acceptable varieties. Row covers can prevent the attack of plants by adult flea beetles, and subsequent damage by larvae, if the crop is grown on land that is not previously infested by potato flea beetle. If the beetles were abundant in the previous season, however, the adults may be overwintering in the soil and can emerge under the row cover. Thus, there is considerable benefit from crop rotation with a nonsolanaceous crop. Destruction of solanaceous weeds is beneficial. Because adults do not fly readily, planting of crops at some distance from previous crops or weed hosts is considered beneficial. Wolfenbarger (1940), working in New York, studied the distribution of potato flea beetle injury in potato fields with respect to weedy, uncultivated areas. Damage to tubers was greatest within 25–30 m of uncultivated areas containing solanaceous weeds. Further, the number of beetles overwintering in soil of uncultivated areas was estimated at about 470,000 per hectare, whereas it was only 55,000 per hectare in potato fields.

Redheaded Flea Beetle

Systena frontalis (Fabricius)

Smartweed Flea Beetle

Systena hudsonias (Forster) (Coleoptera: Chrysomelidae)

Natural History Distribution. Redheaded flea beetle is distributed widely in the eastern United States and Canada. Although it has been collected in Montana, it is rarely found to the west of Manitoba, Kansas, and eastern Texas. Smartweed flea beetle occupies a similar range. It is distributed widely in the eastern states and provinces but is found only as far west as South Dakota, Colorado, and New Mexico. Host Plants. The adults of both redheaded and smartweed flea beetle have been observed to feed on various vegetables, including bean, beet, cabbage, corn, eggplant, lettuce, okra, parsley, potato, and sweet potato. Other crops such as alfalfa, apple, clover, cranberry, currant, gooseberry, grape, horseradish, raspberry, soybean, sugar beet, sunflower, strawberry, and numerous woody ornamental shrubs are also attacked. As might be expected of insects with such a wide host range, numerous weed hosts have been reported, including smartweed, Polygonum

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­pensylvanicum; ­lambsquarters, Chenopodium album; giant ragweed, Ambrosia trifida; plantain, Plantago major; beggartick, Bidens frondosa; redroot pigweed, Amaranthus retroflexus; Canada lettuce, Lactuca canadensis; Canada thistle, Cirsium arvense; and giant foxtail, Setaria faberii. Of the aforementioned weeds, the first four are thought to be preferred by redheaded flea beetle. Redheaded flea beetle larvae have been observed developing on corn, and both species attack sugar beet. It has also been reported feeding on lowbush blueberry in New Brunswick, Canada, by Maltais and Ouellette (2000), who also provide a long list of host records. Life Cycle and Description. Apparently, there is only a single generation per year. Adults are present from midsummer to winter. Eggs are present at all times except spring. The biology of smartweed flea beetle is almost unknown. Following is a description of redheaded flea beetle. Egg. The eggs are elliptical, and slightly more rounded at one end. They measure 0.70–0.85 mm long. Eggs are deposited in the soil beneath suitable food plants, usually at a depth of 1.5–5.0 cm. This is the overwintering stage, and eggs must receive a period of chilling before they will hatch. Larva. Larvae are whitish with a brownish head capsule and bear numerous spines. There is a prominent anal tubercle bearing a tuft of setae. Larvae develop from May to July, feeding on plant roots. Larvae display three instars and reach a length of about 8 mm during development, which requires about 30 days. Pupa. Pupation occurs in the soil.

among the common crop-feeding flea beetles only Systena and Disonycha spp. exceed 3.5 mm. The adults are active from June through September or October in the North, and through November in the South, depending on weather. There is a report of adults overwintering in Indiana, but this has not been confirmed. Eggs are deposited beginning in July, but remain in diapause until the following spring. The adults of smartweed flea beetle are distinguished from redheaded flea beetle only by the lack of an orange-red head. In all other respects, they are virtually identical (Smith, 1970). Further study may demonstrate that smartweed flea beetle is merely a color variant of redheaded flea beetle. The biology of redheaded flea beetle is not well studied. Hawley (1922a,b), Smith (1970), and Jacques and Peters (1971) have provided the most complete information on life history.

Damage The adults feed on the epidermis of foliage, usually the upper surface. They create elongate holes, often leaving the lower epidermis intact as a transparent membrane. They may aggregate on certain plants, and often prefer weeds to crops. Jacques and Peters (1971) noted that the adults were abundant on corn only in the absence of preferred weeds. Corn silks were observed to be preferred over leaf tissue, and beetles were observed to leave corn fields after silks mature. Hawley (1922a,b) noted that bean plants usually recovered from defoliation; the exception was during dry weather when permanent damage or death could occur. The larvae feed on and burrow within plant roots. Jacques and Peters (1971) found redheaded flea beetle larvae interspersed with corn rootworm larvae among corn roots in Iowa, but flea beetle larvae were much less numerous, and less damaging than northern corn rootworm. Riley (1983) documented severe damage to germinating soybean seeds and seedlings by redheaded flea beetle in Mississippi. The larvae scored the seed surface and burrowed through the seed, stem, and roots.

Management

FIG. 5.11  Redheaded flea beetle. (Photo by L. Buss.)

Adult. Adults measure 3.5–5.5 mm long. They are dark reddish brown, with an orange-red head. Head color, the basis for the common name, is diagnostic, although related species such as S. elongata (Fabricius) may have a reddish-brown head. The hind femora are enlarged. The large size of Systena spp. is a good diagnostic character and

Weed management is a key to the effective management of both redheaded flea beetle and smartweed flea beetle, as these insects consistently are reported as pests only where favored weeds are abundant. Chemical treatments applied for corn rootworm larvae are adequate for redheaded flea beetle larvae, though seldom needed. Adult damage is easily prevented with foliar applications of insecticide if the crop is carefully monitored.

Spinach Flea Beetle

Disonycha xanthomelas (Dalman)

Three-Spotted Flea Beetle

Disonycha triangularis (Say)

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Yellow-Necked Flea Beetle

Disonycha mellicollis (Say) (Coleoptera: Chrysomelidae)

Pupa. Larvae drop to the soil to form a small pupation cell. Pupae resemble adults but are grayish in color. Pupation requires 10–14 days.

Natural History Distribution. Spinach flea beetle and yellow-necked flea beetle occur throughout the United States and Canada, east of the Rocky Mountains. Three-spotted flea beetle occupies the same area as the aforementioned species, but are also found in British Columbia, Idaho, and Utah. These insects are native to North America. Host Plants. These flea beetles are known principally as pests of the family Chenopodiaceae—beet, spinach, and Swiss chard. Therefore, it is not surprising that they damage sugar beet, but they are also known to damage cabbage, canola, horseradish, lettuce, and radish on rare occasion. Weeds are their principal host, principally chickweed, Stellaria media; purslane, Portulaca spp.; lambsquarters, Chenopodium album; and pigweed, Amaranthus spp. Natural Enemies. Little is known concerning the natural enemies of these beetles, although Chittenden (1899) reported that Medina barbata (Coquillett) (Diptera: Tachinidae) was a parasitoid of the adult spinach flea beetle, and Loan (1967a,b) found three-spotted flea beetle to be attacked by Microctonus disonychae (Loan) (Hymenoptera: Braconidae). Life Cycle and Description. These insects are poorly known, but spinach flea beetle has two generations annually in Maryland, with adults present throughout the year. The life cycle is reported to require 30–60 days. Apparently, these beetles overwinter in the adult stage and have been observed under loose bark of trees and other sheltered locations during the winter months. From the report of Beirne (1971) that adults are present in May and June and again in September and October, we might surmise that only one generation occurs in Canada. The life cycle of the other Disonycha flea beetles seems to be similar. Egg. The eggs of spinach flea beetle are laid in clusters of about 4–30 eggs, attached on end to leaves, stems, and sometimes on soil. They measure 1.25–1.50 mm long and 0.40–0.57 mm wide, and are orange in color. The eggs of three-spotted flea beetle seem to be undescribed, but they are likely similar to spinach flea beetle. Yellow-necked flea beetle is known to deposit clusters of eggs in a manner similar to spinach flea beetle, but they are somewhat unusual in that they are red in color. Eggs hatch in 4–10 days, the larvae escaping by chewing a slit in the side of the egg. Larva. The larvae are normally grayish, though sometimes purplish when feeding on beet foliage. The head is darker, and the body is well equipped with short and stout spines. Initially measuring about 1.8 mm long, the larvae reach 8–9 mm at maturity. Larvae require 10–30 days to complete their development.

FIG. 5.12  Three-spotted flea beetle. (Photo by J. Capinera.)

Adult. Adults of Disonycha are fairly large for flea beetles, measuring 5–6 mm long. Among the common crop-­ infesting flea beetles, only Systena spp. approach Disonycha in size. Their elytra are shiny black, sometimes tinged with blue or green. The prothorax is yellow or red. Three-spotted flea beetle can be distinguished from the other species by the three black spots on the thorax. Yellow-necked flea beetle can be distinguished from spinach flea beetle by the color of the femora: in the former, the femora are entirely yellow, whereas in the latter they are partly blue or green. Females produce an estimated 100–300 eggs. The biology of spinach flea beetle was provided by Chittenden (1899), that of three-spotted flea beetle by Maxson (1948), and of yellow-necked flea beetle by Chittenden (1912b).

Damage The larvae of Disonycha flea beetles differ from the more common genera in that they feed on foliage rather than roots. They tend initially to be gregarious, but this habit dissipates as they mature. They usually feed on the underside of the leaves, initially feeding only partially through the leaf tissue, but eventually creating holes. Adults similarly feed on the leaf tissue, skeletonizing the foliage. Generally, these flea beetles are not considered serious pests.

Management Because weeds are often important in the biology of these insects, an important element of insect control is weed ­suppression. Should the insects require suppression, foliar insecticides provide quick relief. The beetles do not seem to overwinter in the soil, so covering crops with netting or row cover material will prevent damage in the spring.

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Striped Flea Beetle

Phyllotreta striolata (Fabricius)

Western Striped Flea Beetle

Phyllotreta ramosa (Crotch) (Coleoptera: Chrysomelidae)

Natural History Distribution. Striped flea beetle, Phyllotreta striolata (Fabricius), is common in Europe and Asia, and has gained access to North America before 1700 (Bain and LeSage, 1998). It is now widely distributed in Canada and the United States; however, it is not abundant in the Rocky Mountain region and west coast. It is well known from Canada’s Prairie Provinces, where crucifer oil-seed crops are grown extensively, but is not as abundant, or damaging, as crucifer flea beetle, Phyllotreta cruciferae (Goeze). In the United States, striped flea beetle is most common in the northeast and midwest. Western striped flea beetle, Phyllotreta ramosa (Crotch), apparently is native to North America. Its distribution is restricted to the western United States, principally Washington, Oregon, and California. A report of this insect from British Columbia is suspect. Host Plants. Striped flea beetle is a crucifer-feeding species, and is often called the striped cabbage flea beetle. As is the case with other Cruciferae-feeding insects, it will also feed on mustard oil-containing plants from the families Capparidaceae and Tropaeolaceae. It commonly attacks such vegetables as Brussels sprouts, cabbage, cauliflower, collards, kale, mustard, radish, rutabaga, turnip, and watercress. It is reported to have attacked noncruciferous garden vegetables, but these are erroneous, with the insects actually feeding on cruciferous weeds found among the vegetables. Weeds serving as suitable hosts include black mustard, Brassica nigra; flixweed, Descurainia sophia; pepperweed, Lepidium densiflorum; shepherdspurse, Capsella bursapastoris; yellow rocket, Barbarea vulgaris; and other common cruciferous weeds. Information on host preference was provided by Feeny et  al. (1970) and Tahvanainen (1983). Western striped flea beetle has similar host preferences. Natural Enemies. A parasitoid, Microtonus vittatae Muesebeck (Hymenoptera: Braconidae) attacks the adults of several crucifer-feeding flea beetles, including striped flea beetle. The biology of this parasite was given by Wylie (1982, 1984) and Wylie and Loan (1984). General predators such as the shield bug, Podisus maculiventris (Hemiptera: Pentatomidae) also sometimes feed on beetles (Culliney, 1986). The soil-dwelling stages of striped flea beetle are susceptible to infection by entomopathogenic nematodes, leading to reduced adult populations and reduced adult feeding injury (Yan et al., 2013). Life Cycle and Description. In Canada, striped flea beetle has only one generation per year, though a ­complete

life cycle requires only about 1  month. Overwintering ­occurs in the adult stage in the soil and leaf litter (Burgess and Spurr, 1984; Turnock et  al., 1987). Adults become ­active in early spring, which is March in the southern states, April in the midwest and southern Canada, and as late as June in northern Canada. In Saskatchewan, the abundance of overwintering adults is high through mid- to late-June, with summer generation beetles becoming abundant in late July. Similar population trends have been observed in New York. Beetles are more active fliers early in the season and fly at greater heights (1–2 m) than later in the season (Lamb, 1983). They become active about 2 weeks earlier than crucifer flea beetle, with which they commonly coexist. A second generation is reported from New York and North Carolina, but this is uncertain. The life cycle of western striped flea beetle is unknown, but likely is quite similar to that of striped flea beetle. Egg. Soon after their emergence, beetles begin to deposit eggs in the soil adjacent to host plants. In Manitoba, egglaying females were collected from April to early August. Eggs hatch in about 5 days. Larva. There are three instars. The larvae measure about 1.25 mm long at hatching but eventually attain a length of about 4.9 mm. They are white or yellowish white, with a brownish head and anal plate. The larvae bear an anal proleg. Pupa. Pupation occurs in the soil within a small cell formed from soil particles. Pupae resemble the adults in form but lack fully developed wings.

FIG. 5.13  Adult striped blea beetle. (Drawing by USDA.)

Adult. Adults are about 2.0–2.4 mm long. The hind femora are enlarged. The head and pronotum are black, and the elytra are brownish black, each with a slight metallic luster. The elytra usually bear an irregular yellow band running nearly the length of each elytron. However, the band

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is sometimes discontinuous, resulting in two yellow spots on each elytron. The proportion of the population with the aforementioned discontinuous banding pattern is higher in southern portions of the beetle’s range. The adults emerge and are very numerous in July and August in northern states. Striped flea beetle and western striped flea beetle are very similar in appearance. Males can be easily differentiated, however, by the size of the fifth antennal segment. In western striped flea beetle, antennal segments four to six are nearly the same size. In striped flea beetle, the fifth segment is wider and longer than adjacent segments, nearly twice the length of the sixth segment. The other common striped flea beetle that may get confused with these species is Zimmermann’s flea beetle. However, male P. zimmermanni have their fifth antennal segment even more enlarged, about three times the length of the sixth segment. Information on striped flea beetle biology was given by Dugas (1938), Burgess (1977), and Wylie (1979). Males of striped flea beetle are reported to produce an aggregation pheromone (Beran et al., 2011). Apparently, adults respond to the pheromone only in the presence of host plant volatiles, particularly allyl isothiocyanate. Rearing techniques were provided by Burgess and Wiens (1976). Information on western striped flea beetle can be found in Chittenden (1927) and Smith (1985).

Damage Striped flea beetle damages both the above-ground and below-ground portions of crucifers. Adults are damaging to seedlings in the spring, but even mature plants can be severely damaged when beetles are particularly abundant. Dugas (1938), for example, reported that the foliage from an entire field of mature turnips in Louisiana was so damaged that it appeared burned. Although the beetles produce only small holes in the foliage, when excessive feeding occurs the adjacent tissue dies, leading to a bronzed or burned appearance. Even modest levels of feeding reduce the value of foliage crops such as mustard if the plants are nearing maturity. Beetles sometimes feed on the young florets of broccoli, greatly reducing yield. Larvae feed on both rootlets and the principal portions of the plant root. Larvae often feed along the surface of the main root, leaving shallow trenches on the root surface. On plants where the root is harvested, such as turnip, larval feeding destroys the crop’s marketability. Even in crops where the root is not harvested, such as cabbage, flea beetle larvae can be very damaging, because their root pruning activities stunts the plants and may cause seedling death.

Sweetpotato Flea Beetle

Chaetocnema confinis Crotch (Coleoptera: Chrysomelidae)

Natural History Distribution. In the United States, this insect is found practically wherever sweet potato is cultivated, including California. However, it is considered a pest in the east of the Rocky Mountains only and is not known to be damaging in Canada. Although likely originating in the New World, this species is now ubiquitous in tropical regions around the world. Host Plants. Among vegetable crops, only sweet potato and corn are attacked, and corn only rarely. However, several other crops are sometimes consumed, including clover, oats, raspberry, rye, sugar beet, timothy, and wheat. Weeds of the family Convolvulaceae, particularly bindweed, Convolvulus spp., and morning glory, Ipomoea spp., are the preferred hosts. Natural Enemies. No natural enemies are known from this poorly studied insect. Life Cycle and Description. The life cycle of this insect is poorly known. It appears that there is only one generation annually in the north, while several generations may occur per year in the south. The complete life cycle is estimated to require 4–8 weeks for its completion. Adults are found throughout the season, from April through October, but they are damaging to sweet potato foliage principally early in the season. Egg. The egg is deposited in the soil. It is white and elliptical, measuring about 0.2 mm long. Larva. The larvae are white, and likely resemble larvae of corn flea beetle, Chaetocnema pulicara Melsheimer, or other members of the genus Chaetocnema. Larval development requires about 3 weeks. The larvae attain a length of about 4.8 mm at maturity. Pupa. Pupation occurs in the soil. Pupae are white, resembling the adult in form.

Management These insects are very similar to crucifer flea beetle and often coexist simultaneously. The management considerations discussed under crucifer flea beetle are also applicable to the striped and western striped flea beetles.

FIG. 5.14  Adult sweetpotato flea beetle. (Drawing by USDA.)

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Adult. The adult is small, measuring only about 1.5– 1.8 mm long. Its color is black but has a bronze cast. The elytra are marked with rows of punctures. The legs are reddish yellow, and the hind femora are enlarged. The adult is the overwintering stage and is found under debris and in wood lots near cultivated areas. The adult becomes active early in the year, often in May, and seeks suitable host plants for feeding. Beetles are reported to abandon sweet potato later in the season, as soon as bindweed is available. There is lack of good source of biological information on this insect. Elements of biology can be found in Smith (1910a,b) and Sorensen and Baker (1983).

Damage Damage by adults is not the typical round-hole injury found in most flea beetles, but rather like that of other Chaetocnema spp. Specifically, they feed in long, narrow strips on the foliage. Often the feeding strips are parallel to the major veins, but as damage levels increase, the feeding becomes more general. It is only the young plants that are injured severely. The larvae are not usually found feeding on sweet potato; the adults apparently prefer to oviposit near bindweed. When feeding on sweet potato, the damage is principally due to consumption of fibrous roots, although sometimes larvae bore into the storage roots (tubers). Occasionally they etch the skin of the tuber, forming winding channels on the surface. The amount of damage is positively correlated with soil temperatures, and to a lesser degree, it is negatively correlated with rainfall (Jasrotia et al., 2008). Less damage occurs when sweet potato is planted and harvested late in the season, relative to early in the season.

Management In fields that have a history of flea beetle infestation, preand postplanting applications of granular insecticides are recommended (Chalfant et al., 1979a). If systemic materials are not applied, then foliar applications may be necessary to prevent injury to the vines. Some sweet potato varieties are resistant to root injury (Cuthbert Jr. and Davis Jr., 1970; Abney and Kennedy, 2011). Resistant and nonresistant cultivars can be interplanted, with some lessening of injury among interplanted susceptible stock (Schalk et al., 1992). Reduction in larval injury was reported by application of the entomopathogenic nematode, Steinernema carpocapsae (Nematoda: Steinermatidae) (Schalk et al., 1993).

Tobacco Flea Beetle

Epitrix hirtipennis (Melsheimer)

Southern Tobacco Flea Beetle

Epitrix fasciata Blatchley (Coleoptera: Chrysomelidae)

Natural History Distribution. Tobacco flea beetle, Epitrix hirtipennis (Melsheimer), is distributed widely in the United States. It is found most commonly in the southeast, but also as far north as Maryland and Michigan, and west to southern Colorado and California. It is infrequent in the northwest and Great Plains, but apparently is found in Hawaii and Mexico. In Canada, it is known from Ontario and Quebec. Since the 1980s it has also become a serious pest in Europe and Japan (Bienkowski and Orlova-Bienkowskaja, 2016). Southern tobacco flea beetle, Epitrix fasciata Blatchley, has limited distribution in the United States and is known only along the Gulf Coast, from Florida to Texas. It occurs widely in Central America and the Caribbean, and as far south as Argentina. Host Plants. Tobacco flea beetle adults feed readily upon tobacco and other plants in the family Solanaceae but sometimes attack other plants as well. Vegetable crops that are most frequently attacked are eggplant, potato, and tomato, but cabbage, cowpea, pepper, snap beans, and turnip are consumed occasionally. Weeds commonly serving as hosts are nightshade, Solanum spp.; jimsonweed, Datura stramonium; ground-cherry, Physalis heterophylla; pokeweed, Phytolacca americana; burdock, Arctium minus; cocklebur, Xanthium spp.; and many others. The larvae develop successfully on many solanaceous plants but do not survive on nonsolanaceous plants. In studies conducted in Virginia, tobacco, potato, and jimsonweed were particularly good hosts for larvae (Glass, 1943). The little information available on southern tobacco flea beetle suggests that the host range is nearly identical to that of tobacco flea beetle (White and Barber, 1974). Natural Enemies. Tobacco flea beetle is preyed upon to a limited extent by general predators such as big-eyed bug, Geocoris punctipes (Say) (Hemiptera: Lygaeidae) (Dominick, 1943). The principal parasitoid seems to be Microctonus epitricis (Vierick) (Hymenoptera: Braconidae), which causes parasitism rates of up to 25% (Dominick and Wene, 1941). This parasitoid also attacks southern tobacco flea beetle. A nematode, Howardula dominicki Elsey (Nematoda: Allantonematidae), parasitizes up to 70% of larvae and 50% of adults of tobacco flea beetle. Host larvae are killed, and adult females are made sterile by the nematodes. Male beetles are important in nematode dissemination. The biology of this nematode was provided by Elsey and Pitts (1976), and Elsey (1977b,c). Life Cycle and Description. The number of annual generations of tobacco flea beetle varies, with three reported from Kentucky, 3–4 in Virginia, and 4–5 from Florida. In Kentucky, Jewett (1926) observed first-generation adults, resulting from reproduction by overwintering beetles, in mid-June. This was followed by a second generation in late July, and a third in September. Not all insects undergo the third generation, however, some commence o­ verwintering

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after only two generations. In Florida, Chamberlin et  al. (1924) reported four well-defined generations, but there may be additional generations. Overwintering occurs in the adult stage, under plant debris, and often in weedy or wooded areas adjacent to crop fields. In the south, the beetles may remain active throughout the winter. Egg. Tobacco flea beetle eggs are elongate and slightly pointed at one end. Initially, the egg is white but gradually changes to lemon yellow as it matures. The egg measures about 0.41 mm (range 0.36–0.48 mm) long and 0.18 mm (range 0.16–0.26 mm) wide. Eggs are deposited in the soil near the base of the host plant, frequently in clusters of 5–6 eggs. Moist areas are preferred for oviposition. Eggs hatch in 6–8 days. Overwintering females deposit, on an average, 2.2 eggs per day, and total production is estimated at about 200 eggs per female. Later generations seem to live for a shorter period of time, and the number of eggs produced per female is about 100. Larva. The larvae have three instars and grow in size from  radish > collards > cabbage. Similarly, Balusu and Fadamiro (2011) reported preference of turnip and napa cabbage relative to cabbage and collards. These authors also reported that the volatiles from nappa cabbage were most attractive, followed by turnip, relative to the other crops. Yellowmargined leaf beetle has also been collected from pepperweed, Lepidium virginicum; dock, Rumex sp.; and various clovers and vetch, but it is not clear whether the insects were feeding or just seeking shelter on these plants. Fecundity and longevity on several host plants were reported by Ameen and Story (1997a). They found that turnip was the most favorable host, with an average of 490 eggs produced per female, followed by radish (440), mustard (425), cabbage (271), and collards (199). Adult longevity varied from 68 days when fed turnip and collards to 105 days on radish. Surprisingly, there was no significant relationship between longevity and fecundity. Natural Enemies. There are no known parasitoids of M. ochroloma in the United States. In Florida, predators of this insect include spined soldier beetle, Podisus maculiventris (Say) (Hemiptera: Pentatomidae); a green lacewing, Chrysoperla rufilabris (Burmeister) (Neuroptera: Chrysopidae); and convergent lady beetle, Hippodamia convergens (Say) (Coleoptera: Coccinellidae). Podisus maculventris has been shown to be more effective than C. rufilabris at predation of yellowmargined leaf beetle (Balusu et al., 2017), and both eggs and larvae are consumed. Montemayor and Cave (2012) suggested that the release of 10 first instars of P. maculventris per six turnip plants would suppress the yellowmargined leaf beetle population in Florida. Chrysoperla rufilabris prefers aphids, and although it eats M. ochroloma eggs and larvae; it performs poorly during the winter in Florida, when yellowmargined leaf beetles remain active (Niño and Cave, 2015). Niño Beltrán (2013) noted that C. rufilabris ate more first instars than eggs. Other predators of M. ochroloma have been identified, though poorly studied. These include a brown lacewing, Micromus posticus (Walker) (Neuroptera: Hemerobiidae); a damsel bug, Nabis americoferus Carayon (Hemiptera: Nabidae); anchor stinkbug, Stiretrus anchorago (Fabricius) (Hemiptera: Pentatomidae); a lady beetle, Coccinella septempunctata Linnaeus (Coleoptera: Coccinellidae); spined assassin bug, Sinea diadema Fabricius (Hemiptera: Reduviidae); and milkweed assassin bug, Zelus longipes (Linnaeus) (Hemiptera: Reduviidae) (Balusu et al., 2017). Naturally occurring entomopathogens affecting yellowmargined leaf beetle in the United States are unstudied or nonexistant, as none have been reported. In Brazil, however, Beauveria bassiana has been observed to affect this species under field conditions. Also, epizootics can be induced by application of entomopathogens (see section on management).

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Life Cycle and Description. The life cycle is not well known, and no evidence of diapause has been reported. There appears to be a single generation annually, but the insect can complete its life cycle in less than a month under favorable conditions, so it is possible that more than one generation occurs during mild Gulf Coast winters. In Florida, adults remain active throughout the winter months. Elsewhere, the adults overwinter but become inactive. Damage is noted in the spring and early summer, when both larvae and adults can be found feeding on crucifers. Summer aestivation from mid-June to September or October has been suggested.

but eventually, become solitary. There are normally four instars, lasting a total of 11 days at 15°C, but 7 days at 30°C. Pupa. The larva usually spins a loose net-like pupal case on the underside of foliage and pupates within. However, sometimes pupation occurs beneath leaf litter or other debris. Duration of the pupal stage is 11 days at 15°C, but 4 days at 30°C. Newly formed adults remain within the cocoon for about 2 days before escaping.

FIG. 5.49  Adult of yellowmargined leaf beetle. (Photo by L. Buss.) FIG. 5.47  Eggs of yellowmargined leaf beetle. (Photo by L. Buss.)

Egg. The adults can begin copulation about 6 days after emergence, followed by oviposition in another 3–6 days. The eggs are bright orange, elongate, and deposited upright (on end) singly or in small groups, often on the underside of the foliage, where they are arranged loosely and irregularly. Eggs measure 1.2 mm long and 0.5 mm wide. Duration of the egg stage is 5 days at 30°C, but 16 days at 15°C. Females can deposit up to about 1500 eggs over their lifetime, but often it is much less.

Adult. The adult is about 5 mm long and 2.5 mm wide, oval in shape, and predominantly dark bronze or black. The peripheral edges of the elytra, however, are marked with a margin of yellowish or brownish, which is the basis for the common name of this insect. Each of the elytra is marked with four rows of deep punctures. Males are smaller than females. Adult longevity may exceed 100 days. They may mate repeatedly. Biological information on yellowmargined leaf beetle was given by Chamberlin and Tippins (1948), Woodruff (1974), Oliver and Chapin (1983), Ameen and Story (1997a,b), and Manrique et  al. (2012), but the most complete summary was given by Balusu et al. (2017).

Damage The adults and larvae feed on foliage, making small holes and feeding on the leaf margin. Damage to cruciferous crops is reported mostly in the spring. In the United States, it is a pest in small or home garden plantings, and increasingly in commercial organic farming. In South America, it has been observed to damage crops grown conventionally on a commercial scale, but only before the advent of modern insecticides.

Management FIG. 5.48  Larva of yellowmargined leaf beetle. (Photo by L. Buss.)

Larva. The larva is yellowish brown to black and covered with a fine layer of hairs. The head capsule is dark brown or black. Larvae are gregarious during early instars,

Often this is a minor pest that does not require control except on an isolated plant or planting, but under some conditions, it can become abundant. Monitoring is usually accomplished by visual examination of individual plants, emphasizing the underside of leaves. Sanitation is an important cultural

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practice. Growers should not leave unharvested plants in the field during the winter months, as this favors successful overwintering. Similarly, sanitation should extend to the presence of wild crucifers, which are common winter weeds. Trap cropping has been suggested as a cultural management technique. Specifically, planting turnip along the perimeter of a field can attract adults from less preferred crucifer crops such as cabbage, broccoli, or cauliflower can reduce the need for insecticide application on the main crop (Balusu et al., 2017). Mulching with straw is not a suitable management technique, instead exacerbating damage (Manrique et  al., 2010). Foliar applications of synthetic chemical insecticides are often used when beetle suppression is warranted, although products derived from plants and microbes are often substitutes, especially for organic producers (Balusu et al., 2017). Microbial suppression is largely limited to entomopathogenic fungi such as Isaria fumosorosea, Beauveria bassiana, and Metarhizium anisopliae, and the bacteria Chrombacterium subtsugae, and Bacillus thuringiensis (Balusu et al., 2017). Other nontraditonal products such as the actinomycete-derived spinosad and pyrethrum-based insecticides are effective, as are numerous other synthetic insecticides (Balusu and Fadamiro, 2013).

FAMILY COCCINELLIDAE—LADY BEETLES Mexican Bean Beetle

Epilachna varivestis Mulsant (Coleoptera: Coccinellidae)

Natural History Distribution. Mexican bean beetle is native to Mexico and Central America, and it is likely found in Arizona and southern New Mexico since the introduction of cultivated beans by indigenous peoples several hundred years ago. By the late 1800s, it was damaging beans throughout the southwest, particularly Colorado. A major increase in damage followed the accidental transport of Mexican bean beetle to northern Alabama about 1918, apparently in shipments of alfalfa hay from Colorado and New Mexico. Once gaining access to eastern states, this beetle has spread rapidly to the northeast. By 1922, Mexican bean beetle had invaded Georgia, North and South Carolina, Virginia, Tennessee, and Kentucky. It reached Ohio and Pennsylvania in 1925, Ontario in 1927, New Jersey in 1928, and Connecticut in 1929. It is now found all over the continental United States. In Canada, Mexican bean beetle is found in eastern provinces, from Ontario to New Brunswick, and also is reported from British Columbia, but it is a common pest only in Ontario. It was discovered in Japan in 1997. The central Great Plains, from North Dakota to Texas, formerly provided a natural barrier in spreading of the ­beetles. Although this barrier was bridged through human

intervention, there remains an area that is relatively inhospitable to bean beetle, so they are infrequent in this region. This insect is also infrequent in Pacific Coast states. Thus, there are two fairly discrete populations—a western population in the Rocky Mountain region including the western edge of the Great Plains, and an eastern population that inhabits most of the eastern United States west to Kansas. Host Plants. Mexican bean beetle develops only on legumes. Other plants are occasionally reported injured, but these invariably are growing adjacent to defoliated legumes and will not support reproduction of beetles. Among vegetable crops eaten are cowpea, lima bean, and snap bean, particularly the latter two bean types. Related crops such as faba bean, lentil, and mung bean seem to be immune in the United States, but mung bean, Vigna angularis (L.), is affected in Japan (Abe et al., 2000). Field crops that may be attacked include alfalfa, sweet clover, various dry beans, and soybean. Formerly, field crops other than dry beans (Phaseolus vulgaris) were relatively unsuitable and rarely injured. However, starting in the 1970s the eastern and then midwestern states began experiencing considerable damage to soybean by Mexican bean beetle. The natural host appears to be tick trefoil, Desmodium spp.; however, in the United States, Mexican bean beetle is almost always found associated with cultivated legumes. Lupine, Lupinus spp., were found to support adults in California, but no reproduction occurred. Natural Enemies. Numerous predators, parasitoids, and microbial disease agents of Mexican bean beetle have been identified, but few native natural enemies are considered to be important (Howard and Landis, 1936). Among predators, the soldier bug Stiretrus anchorago (Fabricius) and the spined soldier bug, Podisus maculiventris (Say) (Hemiptera: Pentatomidae), are often cited as the most effective. Lady beetle species such as convergent lady beetle, Hippodamia convergens Guerin-Meneville; transverse lady beetle, Coccinella transversoguttata Brown; and Coleomegilla maculata (De Geer) (all Coleoptera: Coccinellidae) sometimes prey on the eggs or young larvae of bean beetle, and on occasion have been considered important predators. Other predators include minute pirate bugs (Hemiptera: Anthocoridae), damsel bugs (Hemiptera: Nabidae), assassin bugs (Hemiptera: Reduviidae), and lacewings (Neuroptera: Chrysopidae). Parasitoids have not been very effective at suppressing bean beetle. Species native to the United States have not adapted to Mexican bean beetle as a host, whereas species imported from Central and South America have failed to establish permanently. Special attention was given to a species from Mexico, Aplomyiopsis epilachnae (Aldrich) (Diptera: Tachinidae). It was released widely in the eastern states but apparently is not adapted to cold winters (Landis and Howard, 1940). In recent times a parasitoid from India and Japan, Pediobius foveolatus (Crawford) (Hymenoptera: Eulophidae), has

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been ­cultured and ­released in eastern states. The annual release is necessary because the parasitoid is unable to overwinter in the United States (Schaefer et al., 1983). In Maryland, nurse crops of snap beans have been planted early to attract bean beetles, with parasitoids released into these small plantings. As overwintering or reproducing beetles appear elsewhere, parasitoids naturally disperse from the nurse crops to attack the expanding bean beetle population. The nurse crop approach allows development of a large population of parasitoids with minimal effort (Stevens et  al., 1975a). Small gardens, such as those in urban and suburban communities, are also suitable for Pediobius release (Barrows and Hooker, 1981). Fungal (microsporidian) pathogens, especially Nosema epilachnae and N. varivestri’s occur in bean beetles (Brooks et  al., 1985). These pathogens are deleterious to Mexican bean beetle; Nosema epilachnae, in particular, reduces longevity and fecundity in bean beetles (Brooks, 1986). However, these pathogens also infect the parasitoid Pediobius foveolatus. Infection of the parasitoid occurs when the immature stage develops in its host, or when the adult ingests the pathogen (Own and Brooks, 1986). Weather. Although natural enemies may affect bean beetle abundance, the weather is also thought to play an important role in population dynamics. Hot and dry weather is thought to be detrimental to survival of all stages, but especially the egg stage. Temperatures above 35–37°C can be lethal. Mellors et  al. (1984) are among the authors to give information on the effects of various temperatures for several time intervals, with important earlier studies presented by Miller (1930), and Sweetman and Fernald (1930). Marcovitch and Stanley (1930) developed a climatic index based on temperatures and moisture that predicts potential damage by Mexican bean beetles. More recently, Barrigossi et  al. (2001) studied Mexican bean beetle survival in Nebraska, reporting high mortality due to “desiccation.” The egg and early larval stages were most affected. Thus, higher insect populations and greater plant damage are associated with high precipitation years. Life Cycle and Description. Mexican bean beetle usually exhibits 1–3 generations annually. In the western United States, there is normally one complete g­ eneration, with a small number of individuals reproducing and developing a small second generation. However, during cool summers the members of the second generation are unable to complete their development and perish. In the southeast, where three generations are more common, a few beetles deposit eggs that produce a small fourth generation. Throughout the nation, adults are the overwintering stage. They often overwinter in wooded areas if such shelter is available. Lacking wooded areas, they overwinter in and around bean fields. The emergence of the adults occurs late in the spring, and weather affects the timing of adult activity. A prolonged period of warmth usually precedes emergence from ­overwintering.

Dry weather delays emergence. Overwintered adults typically are most abundant in June, followed by first, second, and third generation (when present) beetles in July–August, August–September, and October, respectively. A life cycle may be completed in 30–40 days during the summer months but may require 60 days during cooler weather. In recent years much of the research has concentrated on Mexican bean beetle as a soybean pest, but life history parameters, while similar, are not the same as on more suitable hosts. Larvae reared on soybean tend to have higher mortality and longer development times than on snap bean (Bernhardt and Shepard, 1978; Hammond, 1985).

FIG. 5.50  Eggs of Mexican bean beetle. (Photo by L. Buss.)

Egg. The eggs are deposited on end in clusters of 40–60 eggs, usually on the underside of leaves. They are elliptical and measure about 1.3 mm long and 0.6 mm wide. Eggs generally are yellow, but turn orange-yellow before hatching. They hatch in 5–14 days, with a mean incubation period of 5.7 days. All females from the first-generation deposit eggs, but in South Carolina only 94% of the second and 60% of the third generation beetles reportedly produced eggs as more and more beetles entered reproductive diapause.

FIG. 5.51  Larvae of Mexican bean beetle. (Photo by L. Buss.)

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Larva. Upon hatching, larvae are yellow and armed with a dense covering of branched spines arrayed in six longitudinal rows. The tips of the spines, when examined closely, usually can be observed to be black. There are four instars. The mean duration of instars in a South Carolina study has been reported to be 3.9, 3.6, 3.6, and 3.6 days, respectively (Eddy and McAlister Jr., 1927). In Colorado, its development required 4.8, 4.1, 4.9, and 5.3 days, respectively (Kabissa and Fronk, 1986). The development time has been studied extensively, and values vary somewhat with location and weather, but most of them are similar to the aforementioned studies. After attaining its full size, about 8 mm long, the larva attaches its anal end to a substrate, usually the leaf on which it fed, and pupates. Larvae do not disperse far, usually only a few meters from where they hatched (Barrigossi et al., 2001). Pupa. During the process of pupation the larval covering, which contains the spines, is pushed back toward the point of attachment to the substrate. Thus, the pupa appears to bear spines, but this is simply remnants of its earlier life, and not firmly attached. Rather, the yellow-orange pupa is quite free from projections. In some cases, particularly late in the season, the pupa is not completely yellow but bears brown or black lines. Duration of the pupal stage averages 8.1 days in South Carolina and 9.6 days in Colorado.

d­ epending on availability of food. In behavior, the beetle is rather sluggish and responds to significant disturbance by dropping from the plant and by excreting small drops of blood from the articulations of their legs. This excretion reportedly is a form of defensive chemistry that reduces predation by other insects. The preoviposition period of adults averages 11.5 days. Females commonly produce about 500 eggs, sometimes depositing over 1200 eggs. Mean fecundity is lowest in overwintering females, and increases in later generations. Males are distinguished from females by the presence of a notch at the tip of the abdomen in males. Adults overwinter under leaves and other plant debris, and under logs and stones. Aggregations of several hundred overwintering beetles are not uncommon. In some cases, overwintering may occur adjacent to larval host plants, but adults are strong fliers and often disperse several kilometers to suitable overwintering quarters, often in wooded areas. The biology of Mexican bean beetle was provided by many authors, but among the most complete are List (1921), Howard and English (1924), Eddy and McAlister Jr. (1927), Friend and Turner (1931), and Auclair (1959). Rearing procedures for both Mexican bean beetle and the parasitoid Pediobius foveolatus were given by Stevens et al. (1975b). A review of biology and management was published by Nottingham et al. (2016).

Damage

FIG. 5.52  Adult of Mexican bean beetle. (Photo by L. Buss.)

Adult. The adults are brightly colored beetles that resemble many beneficial lady beetle species, differing from the beneficial species principally in their phytophagous feeding habit. The beetle is hemispherical in shape and bears 16 black spots. The spots are arranged in three rows, with six spots in each of the first two rows and four in the third row; thus, there are eight on each elytron. The background color is usually orange or copper, but ranges from yellow when freshly emerged, to reddish-brown when old. The reddish color is especially evident among overwintered individuals. The beetles normally measure about 6–8 mm long and 4–6 mm wide, but size varies considerably

Larvae and adults feed principally on leaf tissue, but under high-density conditions and when faced with starvation they also feed on blossoms, pods, and stems. Bean beetles feed on the lower surface of the foliage, removing small strips of tissue and usually leaving the upper epidermis and veins intact. The upper epidermis soon dies and becomes transparent, leaving characteristic injury consisting of a number of small transparent spots that is reminiscent of a stained glass window. Entire leaves are quickly reduced to skeletal lacelike remains that have little photosynthetic value and usually dry and die quickly. Larvae are particularly damaging. As the larvae are not very mobile, they usually concentrate their feeding on a single leaf, inflicting complete defoliation before relocating to another leaf. Bean plants are quite tolerant of defoliation. Bean plants can often withstand 50% leaf loss, especially if it occurs early in the season, leaving time for the plants to recover, or if the affected foliage is older and physiologically inferior. Plants are most sensitive to damage during the pod establishment and filling stages (Waddill et al., 1984). On average, defoliation in excess of 10%–20% usually results in decreased yield. Each larva may consume over 30 cm2 of foliage, with about 70% of the consumption occurring during the final larval stage (McAvoy and Smith, 1979). In studies of dry bean response to bean beetle defoliation, Michels Jr. and Burkhardt (1981) estimated that loss would occur with as few as 1.0–1.5 larvae per plant. In contrast, Capinera et al.

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(1987) determined that field beans could tolerate up to 19% defoliation without yield reduction, and that the number of beetles necessary to inflict this level of injury varied from 3 to 20 depending on the length of the adult feeding period. Mexican bean beetle is also capable of plant disease transmission. When Jansen and Staples (1970a) allowed larval and adult bean beetles to feed on plants infected with cowpea mosaic virus, they became capable of transmitting the virus to healthy plants for a 2-day period. Wang et al. (1994) reported slightly longer virus retention times, about 4 days, and suggested that beetles lost their ability to transmit virus in direct proportion to the amount of foliage consumed. Thus, Mexican bean beetles are not particularly effective vectors, but because adults fly freely they may be important in the redistribution of such diseases.

Management Sampling. The eggs and larvae are highly aggregated in distribution. Sampling is usually accomplished by visual examination of plants. Larval, pupal, and adult stages can be collected with sweep nets, however. Barrigossi et al. (2003) recommended counting egg clusters because this gives the maximum amount of time for growers to make plans for management. These same authors documented considerable early-life mortality, however, making it challenging to compute damage potential. They made adjustments in their economic threshold values to allow for mortality, but weather affects mortality rates so predictability of mortality and economic thresholds remain problematic. They also calculated sequential sampling plans for two rates of insecticide use. Insecticides. Modern insecticides have relegated Mexican bean beetle to low status in commercial bean production. They remain a serious problem, however, in home gardens and elsewhere when insecticides are not used. Also, at least occasionally and sometimes regularly, they threaten commercial production if insecticides are not used routinely. With all insecticides, thorough coverage of foliage, particularly the lower epidermis of leaves, is necessary. Systemic insecticides applied at planting often provide good early-season protection, but when beetle densities are very high crops often benefit from later-season applications, in addition to seed or planting-time treatment (Webb et al., 1970; Elden, 1982). Cultural Practices. Cultural practices are of limited value. Beetles fly long distances to overwinter, so crop rotation and destruction of overwintering sites generally are not practical. It is a useful practice, however, to destroy bean plants as soon as they have been harvested, as this may disrupt the development of many immature insects and inhibit the development of additional generations. Early-planted crops may be useful to attract adult beetles, where they can be destroyed by disking, insecticide application, or release of parasitoids. However, though this trap crop approach works to lure beetles from a relatively unpreferred crop such as soybean to a preferred crop such as lima bean, it is not effective in the protection of preferred crops (Rust, 1977).

The use of reflective plastic mulch to repel bean beetles has been investigated. Adults and larvae are deterred by bright light. Beans planted on metalized and white plastic mulches have less damage and greater yields than beans grown on black plastic or bare soil (Nottingham et al., 2016). Row covers, which are usually made from synthetic polyester, can limit access by beetles to plants. The principal limitation to the application of this technology is cost, both for the cover and the labor is associated with placement and removal of the covers. If covers are removed and replaced on a frequent basis, which is normal for many bean crops due to the need to pick the beans regularly, row covers can be burdensome. Biological Control. The principal means of biological suppression of Mexican bean beetle is the release of Pediobius wasps. (This species is discussed in the section on natural enemies.) Despite extensive research demonstrating the feasibility of such releases, commercial bean producers rarely consider such an approach, tending to rely instead on chemical insecticides. Insecticides are somewhat compatible with parasitoid releases because some insecticides dissipate to nontoxic levels in 1–3 days and parasitoid pupae are relatively immune to field applications of insecticides (Flanders et al., 1984). Microbial insecticides also have potential to be used to manage Mexican bean beetle populations. The utility of strains of the bacterium Bacillus thuringiensis was demonstrated by Tamez-Guerra et al. (1999) and Peña et al. (2006). Behle et al. (2006) reported that both blastospores and conidia of Paeciliomyces fumosoroseus were infective to bean beetle. Host-Plant Resistance. Considerable effort has been directed toward identification of species and cultivars resistant to bean beetle oviposition or feeding. In Ohio, Wolfenbarger and Sleesman (1961) showed that Phaseolus spp., principally snap and to a lesser extent lima beans, were susceptible to injury. Cowpea, faba, and other beans generally were not attacked. Campbell and Brett (1966), working in North Carolina, were able to identify some commercial varieties of both snap and lima beans that displayed resistance. Resistance was reflected in reduced oviposition on resistant varieties and decreased the size and lower fecundity of adults reared on resistant plants. Insect development time was not affected. Also, some so-called vegetable-type soybean cultivars, suitable for such uses as sprouts and tofu, have been evaluated for bean beetle resistance and found to vary considerably in susceptibility to damage (Kraemer et al., 1994). Raina et al. (1978) reported that greenhouse screening could be used to identify resistance.

Squash Beetle

Epilachna borealis (Fabricius) (Coleoptera: Coccinellidae)

Natural History Distribution. Squash beetle is reported to occur throughout much of the eastern United States from Massachusetts

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to Kansas in the north and from Florida to central Texas in the south. However, its impact as a pest is generally limited to the Atlantic Coast, from Connecticut to Georgia. It appears to be a native insect. A related and similar-appearing species, E. tredecimnotata (Latreille), feeds on cucurbits throughout Mexico and Central America and has been found in western Texas, and southern New Mexico and Arizona. Host Plants. Squash beetle larvae feed only on cucurbits, and successful larval development has been reported on various squashes, cucumber, watermelon, cantaloupe, and gourds. In addition, wild cucurbits can serve as hosts. Underhill (1923) indicated that prickly cucumber, Echinocystis lobata; and one-seeded bur cucumber, Sicyos angulata, were readily attacked in Virginia. Adults are less restrictive in their diet; in addition to cucurbits, they have been observed to feed on the blossoms and pods of lima beans and cowpeas, on lima bean foliage, and on fresh corn silks. Natural Enemies. Because this insect is a relatively minor pest, it has not been thoroughly studied, and therefore its natural enemies are not well-known. However, general predators such as stink bugs (Hemiptera: Pentatomidae) and assassin bugs (Hemiptera: Reduviidae) are known to attack larvae. Underhill (1923) reported some evidence of attack by tachinids (Diptera: Tachinidae), and he further noted that 25%–33% of eggs were sometimes consumed by predatory ladybeetles (Coleoptera: Coccinelliae) and lacewings (Neuroptera: Chrysopidae). Smith (1893) noted that squash beetle larvae frequently attack eggs, so cannibalism may be an important mortality factor. Life Cycle and Description. Squash beetle can complete its life cycle in 25–48 days, but the average is 32 days. Two generations are produced per year in Virginia, but only one in Connecticut (Britton, 1919). Egg. The yellow eggs are elongate, measuring about 1.8 mm long and 0.7 mm wide. They are laid on end in clusters averaging 45 eggs per mass (range 12–60 eggs). Females deposit eggs at 4–5-day intervals and generally produce about 300–400 eggs during their life span. Eggs hatch 7–8 days after oviposition.

FIG. 5.53  Larva of squash beetle. (Drawing by USDA.)

Larva. The larvae are yellow and have six rows of large spines running the length of the body; the spines are often darker or tipped with black. Underhill (1923) indicated that

the branching characteristics of the lateral row of spines could be used to distinguish the four instars. The number of branches on each spine was given as 0, 4–7, 10–12, and 16–18 for instars 1–4, respectively. The duration of the larval stage is 16–18 days, with average instar duration of 3.4, 3.2, 4.0, and 6.0 days for instars 1–4, respectively. Pupa. Pupation occurs on the underside of cucurbit leaves, or on adjacent weeds, often in a shaded location. The pupa is about 9 mm long, yellow or orange in color, with branched brown spines attached to the old larval covering and clustered near the tip of the abdomen. Duration of the pupal stage is about 7 days.

FIG. 5.54  Adult of squash beetle. (Drawing by USDA.)

Adult. The adult is 8–10 mm long, with the females averaging larger than males. Beetles are yellow to dull red, with 12 large black spots on the elytra and four small black spots on the thorax. The spots on the elytra are arranged in three transverse rows. Adults begin production of eggs about 8 days after emergence. Beetles normally abandon cucurbit fields in August or September, seeking shelter under plant debris or structures in preparation for overwintering. Cracks in the rough bark of trees are especially preferred as a resting site for overwintering though they may also seek shelter under leaves and pine straw. Beetles seem not to penetrate >12–15 m into forested areas, preferring trees near the edge. In Virginia, they move back into the cucurbit fields in May or June. After overwintering, they may live up to several weeks without food while awaiting the availability of cucurbits. The beetles usually begin egg deposition within 2 weeks of feeding, so eggs from the overwintering beetles are abundant in June. The adults from this first generation begin to lay eggs in July, producing the second generation in August. By late summer, the overwintering beetles have died, but both first and second-generation adults are available to overwinter. Underhill (1923) gave a complete description of squash beetle biology, and the report of Brannon (1937) is informative.

Damage The larvae and adults feed on the underside of cucurbit foliage, though adults may also feed on the upper surface.

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Larvae make a circular cut in the leaf before consuming the lower layer of epidermis within the circle. The circular cut or trenching behavior is believed to inhibit the production of sticky phloem sap by the plant, which inhibits feeding (McCloud et al., 1995). The plant tissue is not removed in a conventional manner but crushed between the jaws, and the remnants left in ridges. Adults sometimes feed on the rind of the fruit. Squash beetle feeding behavior is similar to Mexican bean beetle, but these two species are unusual among the family Coccinellidae, which is normally regarded as a very beneficial group of insect predators.

Management Insecticides. Squash beetle is relatively easy to kill with foliar insecticides, though usually, this species does not warrant control. Cultural Practices. Squash beetles are not strong fliers, so relocation of cucurbit fields away from overwintering sites is recommended. Prompt destruction of cucurbit vines in the late summer may kill many of the late-­developing larvae and pupae, reducing the number of beetles that will overwinter successfully. Row covers should serve adequately.

FAMILIES CURCULIONIDAE AND BRENTIDAE—WEEVILS AND PRIMITIVE WEEVILS Cabbage Curculio

Ceutorhynchus rapae Gyllenhal (Coleoptera: Curculionidae)

(Hymenoptera: Eulophidae) has been reported to parasitize the larvae (Chittenden, 1900). Life Cycle and Description. There is one generation per year, with the adult stage overwintering. In Maryland and Missouri, adults emerge from diapause in April. Mating and oviposition occur soon after emergence. Adults feed briefly on foliage and stems before oviposition, but cause little damage. Egg. Females deposit eggs in stem tissue within cavities created by feeding. Some swelling of the plant tissue near the eggs is noticeable. Oviposition is complete by early May. Eggs are elliptical and gray. Egg length is 0.65–0.85 mm; and its width is 0.35–0.45 mm. Eggs hatch in 5–8 days. Larva. The larvae feed in stem tissue. They are whitish, with a yellowish or brownish head. The body tapers markedly toward both the anterior and posterior ends. The body is fairly wrinkled in appearance, with no evidence of legs apparent. Larvae attain a length of 5–6 mm at maturity. Larval development requires about 21 days, and most of them have abandoned their feeding sites by the end of May. Pupa. Larvae drop to the soil to pupate, secreting an adhesive material to form a small cell from soil particles. The larva remains in the cell for about 14 days before pupation occurs. The period of pupation is an additional 21 days. The pupal cell is about 5 mm in length, only slightly larger than the white pupa. Pupation occurs at a depth of  B. juncea (Chinese or brown mustard) = B. carinata (Ethiopian mustard or Abyssinian cabbage) > Sinapsis alba (white mustard). Natural Enemies. Numerous parasitoids are known to attack cabbage seedpod weevil, including Bracon sp. (Hymenoptera: Braconidae); Eupelmella vesicularis (Retzius) (Hymenoptera: Eupelmidae); Eurytoma sp. (Hymenoptera: Eurytomidae); Necremnus duplicatus Gahan and Tetrastichus sp. (both Hymenoptera: Eulophidae); Asaphes californicus Girault, Habrocytus sp., Trichomalus fasciatus (Thomson), and Zatropis sp. (all Hymenoptera: Pteromalidae); and Megaspilus sp. (Hymenoptera: Ceraphronidae). The most important species are Trichomalus fasciatus and Necremnus duplicatus, with the former species probably introduced from Europe accidentally at the same time as the weevil (Hanson et al., 1948). Life tables for cabbage seedpod weevil in Europe have been developed (Haye et  al., 2010) and demonstrated high generational mortality (99.6%), with about half of the mortality occurring during the winter and half during the immature stages (mostly the larval period). Larval ectoparasitoids were the major larval mortality factor. Cárcamo and Brandt (2017) provided an updated list of parasitoids found in North America and Europe. Parasitism is not as effective in North America as it is in Europe (Dosdall and Cárcamo, 2011). Some parasitoids have been introduced to North America, but the early records of parasitoids in North America are based on misidentifications (Gibson et al., 2006). Interestingly, the presence of a co-occurring insect, brassica pod midge, Dasineura brassicae (Winnertz) (Diptera: Cecidomyiidae) seems to affect the survival of cabbage

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s­eedpod weevil. The midge oviposits though weevil feeding and ovipositional punctures, or weevil larval exit holes. The presence of the midge is correlated with damage to weevil eggs, though it is not known if the midge oviposition or larval feeding is responsible for damage to the weevils (Haye et al., 2010). Life Cycle and Description. There is a single generation per year. Adults overwinter in soil or under debris, becoming active in the spring as the air temperatures reach about 15°C. Adults typically feed on nectar and pollen for about a month before beginning egg production. Females denied the opportunity to feed at flowers, which exhibited poor development of their ovaries (Ni et al., 1990). Females are particularly attracted to volatile emissions of flowers (Evans and Allen-Williams, 1992). The egg development is favored by temperatures of 15°C and 20°C, relative to 10°C and 25°C (Ni et  al., 1990). Flight is limited to periods of relatively low wind speed ( 200 days if provided with adequate food. An interesting aspect of adult biology is that they are flightless. Therefore, dispersal is dependent on walking by adults, or transport of infested tubers or other plant material containing larvae or other unapparent stage; the transport of plant material is undoubtedly the principal means of long-distance dispersal. The biology of this species was reviewed by Sherman and Tamashiro (1954), and Raman and Alleyne (1991). Pierce (1918) provided a detailed description and pictures the different stages.

Damage This is the most severe weevil pest of sweet potato in the South Pacific and Caribbean areas. Successful cultivation of sweet potato on some Caribbean islands hinges largely on the successful management of this insect, which is known locally as “scarabee.” It damages tubers both in the field and storage. Insect damage induces terpenoid production in sweet potatoes, making them bitter and unpalatable. Larvae infesting the tuber often burrow deep into the tuber, leaving little evidence of their presence on the surface. However, the center of the tuber may be thoroughly tunneled and packed with fecal material. Eventually, the tuber becomes blackened, and often infected with plant pathogens. Vines that are infested become wrinkled, cracked, or collapsed.

Management Insecticides. Insecticides are often used to protect plants from injury. Planting stock sometimes is dipped in insecticide before treatment; this is particularly important if the planting stock is already infested. Insecticide is also incorporated into the planting bed and applied to foliage. Systemic insecticides are preferred because so much of the life cycle occurs inside the plant. Cultural Practices. Several cultural practices materially affect West Indian sweetpotato weevil abundance (Sherman and Tamashiro, 1954). Foremost is sanitation. Propagation of sweet potato is often by apical cuttings or “slips,” which can be infested. Plant debris is often infested and may serve as a breeding site for weevils between crops; such material should be destroyed. Ipomoea weeds should be removed from areas near crop fields as they serve both a source of initial infestation and provide harborage between crops. Crop rotation is particularly effective, because the adults are flightless. Storage of tubers should be distant from fields used to culture sweet potato. Host-Plant Resistance. Considerable work has been done to assess naturally occurring host plant resistance. Varieties with thin stems are less injured by weevils. Unfortunately, they produce less satisfactory yields and

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their tubers tend to grow close to the surface, where they are more easily accessed by weevils. However, though numerous varieties or isolates contain some elements of resistance, considerable work remains to be done before commercially acceptable varieties with reliable resistance are available. It is likely that host-plant resistance will prove to be only a component of the management program, and cannot be relied upon to provide complete protection from attack (Raman and Alleyne, 1991). Other Methods. Irradiation can be used to implement the sterile insect technique with West Indian sweetpotato weevil. Release of sterile, laboratory-reared insects neutralizes the fertility of wild insects, disrupting their reproductive abilities. This has been shown to be technically feasible with this insect in Japan (Follet, 2006; Kumano et  al., 2008b).

Whitefringed Beetle

Naupactus spp. (Coleoptera: Curculionidae)

Natural History Distribution. The status of whitefringed beetle has long been in dispute because of disagreement over one or more species of “whitefringed beetle.” Even the genus designation is disputed. Lanteri and Marvaldi (1995) synonymized Graphognathus (the old genus of whitefringed beetles) with Naupactus, recognizing five species. They are native to southern South America. The species now occurring in the United States are N. leucoloma Boheman, N. peregrinus (Buchanan), and N. minor (Buchanan); the first two species are of greatest economic significance in the United States. They are usually considered to be a pest species complex, and even in the scientific literature, rarely are attempts made to distinguish these very similar insects except by insect taxonomists. Two other species of Naupactus are found in South America. Whitefringed beetle invaded the southeastern United States from South America about 1936, and was first recovered from western Florida. Alabama and Mississippi are the states most infested with this insect, but it is now distributed from Virginia to Florida in the east, and to Missouri and Texas in the west. This beetle has also been found in New Mexico and California, but its distribution is limited there. The potential range of this species is believed to include most of the southern United States, from Virginia and Kentucky to southern Colorado and nearly all of California. The Naupactus spp. differ in their environmental preferences, however, so although N. leucoma is predicted to thrive in both the southeast and western coastal regions of the United States, N. peregrinus is expected to be limited to the southeastern region (Lanteri et al., 2013). Whitefringed beetle is considered a pest in Argentina, Brazil, Chile, and

Uruguay. Australia, New Zealand, and South Africa have also been invaded. Host Plants. Whitefringed beetle (particularly N. leucoma) is considered to have a very wide host range, with the number of plants eaten estimated at > 250 species. Plants in the family Fabaceae often are most affected. Grasses, however, are considered to be relatively poor hosts. Vegetable crops attacked include bean, carrot, corn, cowpea, cucumber, lima bean, mustard, okra, potato, pea, squash, sweet potato, and watermelon. Whitefringed beetle perhaps is better known as a field crop pest, attacking alfalfa, cotton, peanut, soybean, tobacco, and velvetbean. Numerous broadleaf weeds are fed upon by both adults and larvae, whereas shrubs and trees are fed upon principally by adults. Young pine trees planted on converted croplands containing residual larval populations can be seriously damaged. The effects of host plant were well demonstrated by Ottens and Todd (1979), who documented the effects of 10 plant species on fecundity and growth of N. peregrinus. For example, mean fecundity (eggs per beetle) on peanut, squash, soybean, coffeeweed, cowpea, okra, crotalaria, cotton, corn, and sorghum was 1013, 765, 715, 560, 470, 325, 266, 133, 24, and 0, respectively. Fecundity is often highly correlated with overall suitability of a food source. Longevity and development time were less affected than fecundity, with few differences among diets. Natural Enemies. No insect parasitoids are known from the United States or South America. Nematodes, particularly steinernematids, have been observed as significant mortality factors in some locations, but apparently, they are limited to heavier soil types. Several pathogens, including the fungus Metarhizium anisopliae, a fungus (microsporidian), Nosema sp., and undetermined bacteria have been observed on numerous occasions to infect larvae. Also, generalist predators such as ground beetles (Coleoptera: Carabidae), stink bugs (Hemiptera: Pentatomidae), ants (Hymenoptera: Pentatomidae), and birds prey on adults. Life Cycle and Description. In North America, there is generally one generation per year. In Alabama, eggs occur in mid- to late-summer, early instars tend to be found in late summer, intermediate instars in autumn and winter, and late instars in the spring months (Zehnder, 1997). In the colder regions of South America, a complete life cycle requires 2 years, so prolonged development may also occur among some populations in the United States, and this has been reported even among some southern populations. In all locations, the larval stage generally overwinters, but overwintering eggs have also been observed. Reproduction is by parthenogenetic females in North America and most of South America, but in Argentina and Paraguay some populations are bisexual (Lanteri and Marvaldi, 1995). Egg. Females deposit eggs in clusters, usually numbering from 10 to 15 per egg mass. Eggs measure about

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0.9 mm long and 0.6 mm wide and are elliptical. The eggs initially are white, but after 4–5 days they turn yellow. The eggs are covered with a sticky gelatinous material that hardens, causing the eggs to adhere to the substrate. They are deposited in various locations, including on the foliage and stems of plants, and below-ground, but most commonly they are deposited at the surface of the soil adjacent to the host plant. Egg production varies from only a few (15–60 per female) when larvae feed on less suitable host plants such as grasses, to numerous (1500 or more per female) when favorable host plants such as peanut or velvetbean are available. Eggs deposited during the warmer seasons hatch in 10–30 days, with an average of about 15–17 days during the summer months. During the winter months eggs may require 100 days to hatch; viability is greatly reduced during the winter. The eggs reportedly require moisture for the hatch to occur. Larva. The larva is legless, and yellowish white, except for the head, which is brown. The larva lives within the soil and feeds principally on roots, though it frequently burrows into below-ground parts of the plant, such as tubers. The summer, autumn, and winter months normally are spent in the larval stage, followed by pupation in the spring, but some apparently pass a second year in the larval stage. Pupa. At maturity, the larva secretes adhesive which hardens and forms a pupal chamber. Larvae normally pupate in May–July. Pupation normally occurs within the top 5–15 cm of soil, but sometimes occurs at greater depths. The pupa measures about 12 mm long and is initially whitish but darkens as maturity approaches. Duration of the pupal stage is about 13 days, ranging from 8 to 15 days. Emergence is stimulated by moist conditions—neither dry nor wet soil is favorable.

sides of the head also bear light-colored bands. The body is densely covered with short hairs. They range in size from about 6 to 12 mm. The elytra are fused together and the wings reduced; thus, the beetles are unable to fly. Only females are known. Adult emergence is typically protracted, with beetles emerging from the soil in May–August. Emergence is particularly apparent after rainfall. The biology of whitefringed beetle was provided by Henderson and Padget (1949), Young et al. (1950), Warner (1975), and Zehnder (1997). Descriptions and keys to the species were found in Lanteri and Marvaldi (1995). Observations from South America were given by Berry (1947).

Damage These insects are able to develop on a wide range of host plants and are relatively unchecked by natural enemies. Thus, they build up to very high levels. All the larvae are long lived; they may destroy replanted fields unless they are killed. They seem to thrive on field crops such as alfalfa, peanuts, and velvetbean, so vegetables following these crops are especially prone to injury. Lawns, woods, swamps, and old-stand vegetation are not considered to be suitable habitats. Rather, well-drained and disturbed environments are most suitable. The larvae feed on the roots, sometimes completely severing small roots. However, the principal damage results from burrowing into fleshy tissue such as the main tap root, tubers, and below-ground stems. Often, the damage is limited to surface scars or channels. Large sweet potato tubers are especially damaged (Zehnder, 1997). On fibrous roots, such as those found on trees, feeding may be limited to removal of the cortical tissues. Early symptoms of feeding injury include discoloration and wilting, but this may be followed by plant death. As is the case with some other soil-dwelling beetles, the larval stage can damage subsurface drip irrigation tape (Nicholas, 2010). The adults may feed on the foliage, particularly of broadleaf plants. They make irregular feeding sites, or notches, along the margins of leaves. Although they may occasionally defoliate a plant, the adult injury is not usually considered serious.

Management

FIG. 5.74  Adult of whitefringed beetle. (Photo by J. Capinera.)

Adult. The adult is brownish gray, with a lighter band along the outer margins of the elytra. The thorax and the

Sampling. Damage potential is sometimes determined by sampling soil and sieving for larvae. Soil beneath favored plants, particularly weeds, is often selected for sampling. The frequency of leaf notching is directly related to the abundance of adults and serves as a convenient index of damage potential. Insecticides. Persistent soil insecticides were formerly used extensively for larval suppression, but their use has been discontinued owing to environmental concerns. Less

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persistent insecticides are often applied at spring planting and kill many overwintering larvae. However, larvae that survive may produce another generation that causes damage in the autumn after insecticide dissipation. Adult control is sometimes practiced. Foliar insecticides targeted against adults have been shown to reduce grub damage to sweet potato roots (Zehnder et al., 1997). Planting-time seed treatment with systemic insecticides followed by foliar treatments later in the summer seem to provide good protection against damage to sweet potatoes. Cultural Practices. Although whitefringed beetle has a wide host range, rotation to grains and grasses is recommended because the fibrous root systems of grasses are not easily damaged, and larval growth is suboptimal. Also, planting of legume crops is discouraged, except during the winter months, because they are favored hosts.

FAMILY ELATERIDAE—CLICK BEETLES AND WIREWORMS Corn Wireworm

Melanotus communis (Gyllenhal)

Oregon Wireworm

Melanotus longulus oregonensis (LeConte) (Coleoptera: Elateridae)

Natural History Distribution. The genus Melanotus is widespread in North America, but the western half of the continent has relatively few species. This is likely related to their preference for moist habitats. Corn wireworm is found throughout the eastern United States and occurs as far west as Nebraska and Texas. In Canada, it is known from Ontario and Quebec. Oregon wireworm occurs in all states west of the Rocky Mountains and in British Columbia. Although these are the most common Melanotus spp. affecting vegetables, others including M. depressus (Melsheimer), M. verberans (LeConte), and M. cribulosus (LeConte) are sometimes reported to be damaging, particularly to corn. Apparently all these are native species. Host Plants. Wireworms are omnivorous, often feeding on other soil insects as well as the roots of grasses, broad-leaf weeds, and crops. Melanotus species are reported to damage vegetables such as cabbage, corn, escarole, lettuce, pepper, potato, sweet potato, and other crops such as field corn, sorghum, soybean, sugarcane, and wheat. Adults of wireworms (click beetles) tend to favor grassland and pastures for oviposition. Due to their multiyear development times, crops may experience wireworm injury for several years after land conversion from pasture to vegetable crop production. Natural Enemies. The natural enemies of wireworms are not well known, and generally, seem to be u­ nimportant.

In Florida, M. communis is parasitized by the wasp Pristocera armifera (Say) (Hymenoptera: Bethylidae), but Hall (1982) reported that only about 4% of wireworm larvae were affected. Insect predators such as ground beetles (Carabidae) and rove beetles (Staphylinidae), as well as stiletto flies (Therevidae) are known to feed on wireworms. Birds, particularly crows, also feed on them (Thomas, 1940). The same microbial pathogens that affect most other soil-dwelling insects affect wireworms, namely Beauveria bassiana and Metarhizium anisopliae. Life Cycle and Description. The biology of Melanotus communis is poorly studied relative to other wireworms, which is surprising considering the frequency of association with corn. In Iowa, they are reported to have a 5-year life cycle, with egg deposition occurring in June, egg hatching in July, and larvae requiring four additional summers to complete their development. In southern Florida, however, the life cycle apparently is reduced to 2–3 years (D. Seal, personal communication). Pupation occurs in the autumn. Although Fenton (1926) suggested that adults emerge from the soil to overwinter under the bark of trees, Hyslop (1915) documented overwintering by adults in the soil within pupal cells. Adults reportedly feed on pollen (Fenton, 1926). Studies in Florida showed that most flight activities occurred in May and June. Adults fly in the evening hours. Females contain 50–100 eggs within their ovaries, indicating a fairly high fecundity (Cherry and Hall, 1986). A strong association of larvae with cool and moist soil is evident, though warmer soil may be tolerated if it is moist (Shepard, 1973b). Adults are reddish brown to dark brown, and measure 10–13 mm long. Larval morphology is fairly typical of wireworms. They are yellow to yellow brown, shiny, and elongate. The head, thoracic plate, and anal plate are darker. The most distinctive feature about Melanotus larvae is the lack of a notch at the tip of the abdomen in combination with a flattened abdominal tip. A key to distinguish Melanotus from the other common vegetable-attacking genera of wireworms can be found in Appendix A. Melanotus longulus is perhaps better studied, though it is less important except west of the Rocky Mountains. As is the case with many other wireworm species, development is protracted. Stone and Howland (1944) reported a mean development time (egg to adult) of 1218 days. Most individuals completed their development in 2 or 3 years, but some required 4 or 5 years. The life cycle typically follows the following pattern: overwintering as adults; adult emergence from the soil in April–May; egg deposition in May; egg hatching in June; larvae are most abundant in the summer, but because the larval period is so protracted it may extend for 2 or more years; pupation occurs about August; overwintering adults are found beginning in August or September. Egg. The egg is grayish white, but becomes darker as the embryo develops. It is elipso-cylindrical, with the ends

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broadly rounded. It measures about 0.5 mm in length and 0.37 mm in width. The eggs are mostly deposited near the surface of the soil (upper 3 cm). In California, under field conditions, mean egg incubation was about 32 days, though ranging from 26 to 38 days. Females typically deposit about 150–175 eggs during their life span.

FIG. 5.75  Larva of corn wireworm. (Photo by J. Capinera.)

Larva. Larval development occurs in the soil. The larva is yellow brown and elongate, with a flattened head capsule. Larval development is protracted, usually requires 3–4 years. Mean larval development time was 1158 days, though quite variable. For individuals with a 2 year life cycle, larval development averaged 433  days, whereas for the 3 year life cycle it was 803 days, for 4 years it was 1176 days, and for 5 years it was 1547 days. Prior to pupation, the larva enters a prepupal period that is characterized by inactivity and a thickened body. Duration of the prepupal period averages 8.5 days. Pupa. Pupation occurs in the soil. The pupa is similar to the adult, though the appendages are immovable. The whitish pupa darkens as it matures.

FIG. 5.76  Adult of corn wireworm. (Drawing by USDA.)

Adult. Males are 8–12 mm in length and measure 2.2– 3.2 mm in width. They are rather elongate, and brown or reddish brown in color. Females are similar though larger, measuring 10–13 mm in length, and 2.8–3.8 mm in width. The appendages are lighter in color. Adults emerge in the spring (April–May) and typically survive only about a month. Males tend to emerge earlier than females. The insects are more active late in the day or on cloudy days. Mating likewise is more frequent under these conditions. Newly emerged females typically require about 11 days before oviposition commences. Biology of M. longulus was given by Stone and Howland (1944). Keys for the identification of the genera of adult Elateridae can be found in Arnett (1968), and of larvae in Wilkinson (1963) and Becker and Dogger (1991) and Appendix A. Keys for the identification of adult Melanotus were provided by Quate and Thompson (1967). A key for Melanotus larvae of mid-western corn fields was developed by Riley and Keaster (1979). The ecology and management of wireworms were reviewed by Thomas (1940). Vernon and van Herk (2013) provided a useful treatment of wireworms and wireworm management in potatoes; this chapter contains modern perspectives and is broadly applicable to other vegetable crops. Barsics et  al. (2013) review management of Agriotes spp. Traugott et al. (2015) provided a broader but similar overview of wireworm pests.

Damage The larvae of Melanotus feed below-ground on seeds, roots, tubers, and other plant tissue. They kill young plants and deface the surface of potato and sweet potato tubers. Normally they dig small pits on the surface rather than large cavities. Melanotus spp. prefer high moisture, and the heavier, wet portions of fields are most likely to experience damage. Damage is exacerbated by dry soil conditions, however. Larvae tend to move toward the soil surface in the spring as the soil warms, where they feed until the soil attains about 21°C; they then move deeper in the soil where it is cooler. Larvae again return to the upper layer of the soil in the autumn when the soil surface is cool, and feed until it becomes cold, then they return to a position deep in the soil until the soil warms in the spring (Fisher et al., 1975). Wireworms were reduced in pest status from being regularly serious pests to only sporadic pests by the introduction of chlorinated hydrocarbon (organochlorine) insecticides in the 1940s. These insecticides were characterized not only in being toxic but in being persistent. Thus, many agricultural fields were inhospitable to wireworms, and their overall abundance and damage decreased. However, following the deregistration of these persistent insecticides in the 1970s and 1980s, damage by wireworms increased. Many of the replacement insecticides (many organophosphates, carbamates, pyrethroids, and neonicotinoids) were not as toxic to wireworms, and much less persistent. Thus, their effect

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was limited mostly to wireworms present early in the season, and relatively ineffective on insects present late in the season. By the 1990s, there was a worldwide resurgence in wireworms as pests. Only recently have some new insecticides (e.g., a phenyl pyrazole, fipronil) provided longterm control of wireworms, thereby reducing the necessity for multiple applications of insecticides within a season. However, some of the insecticides now used for soil insects, including wireworms, do no not act in the traditional manner. Specifically, some of the chemicals act more as repellents than as toxicants, or induce a moribund condition but do not actually kill the wireworms, which then become active again at a later date. Thus, damage may be reduced, but the wireworm population persists and may cause damage at a later date, sometimes months later (Vernon and van Herk, 2013).

Management Sampling. Considerable effort has been made to develop wireworm sampling techniques. Growers are advised to assess the population densities of wireworm larvae before planting in the spring. Soil cores can be taken, or pits can be dug, then the insects present can be separated from the soil. Larvae normally are separated from the soil by sifting; a screen with 6–7 meshes per centimeter is desirable. These absolute sampling approaches allow relatively accurate determinations of insect abundance, at least at higher insect densities, but are time and labor intensive. Relative sampling techniques are easier to implement. Population estimates at low larval densities are perhaps best accomplished using baiting techniques, as wireworms move through the soil to reach the bait. Baiting is accomplished by burying whole wheat, corn, sorghum, or other attractive food sources in the soil at a depth of 10–15 cm and then counting the number of wireworm larvae attracted (Apablaza et  al., 1977; Jansson and Lecrone, 1989). As few as 12 bait stations per hectare are often recommended. Soaking the bait for 1 day before distribution increases its attractiveness. Baiting does not work well if the soil is  1 m during a season, whereas in fallow ground they may move several meters.

Egg. The eggs are usually deposited in the proximity of favored adult food plants. Grasses are the favored oviposition site, and moist soil is preferred over dry. The female digs in the soil to a depth of 5–10 cm to oviposit. She normally deposits about 3–4 eggs at a location, and repeats the digging and oviposition processes over a period of weeks until 40–60 eggs are produced. The maximum number of eggs produced is about 130. The eggs are unusual as they vary slightly in shape, ranging from spheres initially measuring 1.5 mm in diameter to ellipsoids measuring 1.5 mm long and 1.0 mm wide. The eggs are white to yellowish white. They enlarge as the embryo matures, as is common among scarab beetle eggs, until they are about twice the original size. Enlargement is due to water absorption, and lack of water inhibits egg development or causes the death of eggs. During dry periods females selectively oviposit in low, poorly drained areas and irrigated fields. Duration of the egg stage is about 21 days at 20°C, but only 8 days at 30°C. No egg hatch occurs below 15°C or above 34°C, and 30°C seems to be about optimal.

Pupa. The pupa develops within a cell in the soil. The pupa resembles the adult, though the elytra are shrunken and distorted. It measures about 14 mm long. Initially, the pupa is white but gradually assumes a metallic green color. Pupae develop at temperatures of 13–35°C. Duration of the pupal stage is about 17 and 7 days at 20° and 25°C, respectively.

Larva. There are three instars during the larval stage. Upon hatching the young grub measures about 1.5 mm long,

FIG. 5.93  Adult of Japanese beetle. (Photo by D. Hall.)

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Adult. The adult remains in its pupal cell for 2–14 days before digging to the surface and emerging. The beetle is oval, and measures 8–11 mm long and 5–7 mm wide. On average, the female is slightly larger than the male. The beetle is principally metallic green, including the legs, but the elytra are coppery brown. The elytra do not quite cover the tip of the abdomen, exposing a row of white spots on each side of the abdomen and a pair of spots on the dorsal surface of the last abdominal segment. The dorsal surface of the body is largely smooth and lacks pubescence, but the ventral surface bears short gray hairs. The head and thorax bear many punctures, and the grooves of the elytra also are punctate. Adults normally live 40–50 days, but under extreme weather conditions may live much shorter or longer. Beetles often disperse to sunny locations to feed. They also seem to be attracted to damaged or weakened plants. Adults usually mate while on foliage and may remain in copula for several hours and mate repeatedly. Males often aggregate near recently emerged females and attempt to mate. They sometimes grapple with one another as they attempt to mate, forming balls of beetles that consist of 25–200 males surrounding a female! Males locate virgin females by her production of sex pheromone (Tumlinson et al., 1977). The most complete treatments of Japanese beetle were provided by Fleming (1968, 1972, 1976), but there are numerous lengthy reports on biology or management, and a recent comprehensive treatment is provided by Potter and Held (2002). Sim (1934) gave an account of characters to distinguish Japanese beetle larvae from similar grubs.

Damage This is a very important insect in the northeastern United States, but less so on vegetables than some other plants. Their overwhelming abundance and the tendency of adults to aggregate are significant elements in their damage potential. Adults feed on the upper surface of foliage, eating the tissue between the veins and leaving a lace-like skeleton. Such leaves invariably perish. They also feed on the stems of succulent asparagus, and on the young silk of corn. They attack fruit and have the curious habit of feeding readily on the fruit of peach but avoiding its foliage. Adults consume foliage at the rate of 30–40 sq. mm per hour. Grubs feed on plant roots; initially, the rootlets are attacked, but larger roots are consumed as the larva matures. Although the roots of grasses are preferred, roots of many other plants, including numerous vegetables, are consumed.

Management Sampling. Adult population monitoring is usually accomplished with traps that use chemical odors as a lure. Both food-based and sex pheromone-based traps, or a combination trap, can be used successfully. The components incorporated into multilure traps usually include phenethyl propionate, eugenol, geraniol, and sex pheromone. Metcalf

and Metcalf (1992) gave a good synopsis of Japanese beetle chemical attractants. Klostermeyer (1985), working in Tennessee, reported higher catches with traps baited with sex plus food lures than food lures alone. Volatiles released by foliage damaged by insects are also highly attractive to adults Loughrin et al. (1995). Trap design is not as important as is the composition of the lure. However, trap design and placement are important considerations, and sex ­attractant-containing traps placed close to the soil (30 cm height) are more effective than those positioned at 90 cm elevation (Alm et  al., 1994). Interestingly, traps containing only food-based lures are more effective when placed at more elevated locations (Ladd Jr. and Klein, 1982). This is not too surprising when the ecology of Japanese beetle is considered: virgin females emerge from the soil, and trees are preferred-host plants. Populations are aggregated, and Allsopp et al. (1992b) estimated that 11–66 traps might be necessary to estimate population density accurately, depending on the density of the beetles. Smitley (1996) reported a positive correlation between adult catches in traps and larval abundance, thereby justifying the use of traps for population monitoring. Insecticides. Persistent insecticides formerly were applied to the soil to prevent injury to roots by grubs. Insecticide resistance developed, especially on golf courses and other areas where esthetic values are important (Niemczyk, 1975). The registrations on many persistent insecticides have since been canceled, and less persistent insecticides are used for root protection in crops or turf. Nevertheless, they remain a principal tool for grub management in the field, incorporated into potting soil, and as a dip for ornamental plant root balls when plant material is to be shipped. Insecticides are also applied to foliage for adult control. Foliar protectants usually must be applied repeatedly, especially if materials with short residual activity are used. Biological Control. Milky spore disease formulations consisting of Paenibacillus popilliae and P. lentimorbus are commercially available. This product usually is recommended for control of grubs on turf, where most larvae develop, but this materially benefits vegetable production by reducing the numbers of adults present as defoliators. In recent years there has been considerable interest in the use of entomopathogenic nematodes as biological insecticides for larval control. Steinernema and Heterorhabditis spp. (Nematoda: Steinernematidae and Heterorhabditidae) can be applied to turfgrass successfully, especially if applications are followed by watering. High levels of suppression can be attained, though there are significant differences in nematode species and strain effectiveness (Klein and Georgis, 1992; Selvan et al., 1994; Potter and Held, 2002). Some work indicates that nematodes may be more effective than insecticides (Cowles and Villani, 1994). Research on nematodes has been largely restricted to the turf, though the aforementioned findings are likely applicable to crops also.

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Traps. Several studies have been conducted to assess the potential of trapping to eliminate or suppress beetles, or to protect plants. In general, though large numbers of beetles have been captured in such efforts, the results are disappointing. For example, large numbers of traps containing food-based lures were placed on Nantucket Island, Massachusetts, for a 3-year period. Although densities were reduced by about 50%, the beetles persisted (Hamilton et al., 1971). In Kentucky, placement of 2–7 traps adjacent to host plants not only failed to protect susceptible plants, but they also seemed to increase damage levels. Cultural Practices. Nonhost plants act as an impediment to the movement of Japanese beetle. Sorghum, for example, which is not eaten by Japanese beetle, reduces the rate of dispersion of adults from soybean patch to soybean patch (Bohlen and Barrett, 1990). In contrast, strip cropping of corn and soybean has no effect on adult distribution (Tonhasca Jr. and Stinner, 1991), probably because both plants are suitable hosts. Screening and row covers can often be used to prevent adults from gaining access to vegetable plants. Unfortunately, corn is probably the most highly preferred vegetable, and this plant is too large to cover conveniently. Injury to corn can be prevented or reduced by modifying the time of planting. Early planting, in particular, is beneficial in the culture of corn because the corn ears are pollinated before the adults become abundant and consume the corn silks; once the ear is pollinated the silks have no value. Where the growing season permits, planting of lateseason varieties is also beneficial because the silks are not produced until after most adults have perished. There are numerous reports that certain plants if eaten are toxic to Japanese beetles. Plants reported to be toxic are geranium, Pelargonium sp.; bottlebrush buckeye, Aesculus parviflora; and castorbean, Ricinus communis. There is evidence that zonal geranium (Pelargonium × hortorum) flowers are toxic, but castor bean does not seem to affect Japanese beetle. Castor bean is known to cause mammalian toxicity, which may account for this erroneous information concerning Japanese beetle. Unfortunately, though geranium and bottlebrush buckeye are, in fact, somewhat poisonous to Japanese beetle, they are rarely eaten by this insect, so impart little suppressive value. Soil preparation can affect Japanese beetle grub populations, though some reports seemingly are contradictory. Because beetles favor acidic soil, application of lime to acidic soil to attain a neutral pH is reported to lower grub populations (Polivka, 1960). On the other hand, the application of aluminum sulfate, which results in acidification, is also reported to reduce grub abundance (Potter et al., 1996). Hydrated lime, though not insecticidal, is a fairly good repellent (and may account for the seemingly contradictory recommendations). Lime must be applied frequently, as either dust or aqueous suspension, to be effective (Fleming et al., 1934). Tillage can destroy grubs, and repeated tillage or rototilling is more disruptive than single cultivation.

Oriental Beetle

Anomala orientalis Waterhouse (Coleoptera: Scarabaeidae)

Natural History Distribution. Oriental beetle is an immigrant species. It is likely originated in the Philippines or Japan but was found in Hawaii in 1908 and Connecticut in 1920. It has since spread to other northeastern states and southward, with its current distribution extending from Massachusetts and New York in the north to North Carolina in the south, and also in Ohio near Lake Erie. Since its initial introduction, its rate of spread has been surprisingly slow. Host Plants. Oriental beetle causes relatively little injury to vegetables, though it may feed on bean, beet, onion, and rhubarb. Also, sugarcane is severely damaged in Hawaii, as well as strawberry and nursery stock in the northeastern states. Oriental beetle is principally a pest of turfgrass, where the larval stage attacks roots. Other plants occasionally injured, principally by the adults, include such flowers as dahlia, hollyhock, phlox, and rose. Most damage to economically important plants results from feeding by the larval stage. Natural Enemies. Several natural enemies were imported into Hawaii and released, including two species which were successfully established—Campsomeris marginella modesta Smith (Hymenoptera: Scoliidae) and Tiphia segregata Crawford (Hymenoptera: Tiphiidae). The former species is considered to be quite important as a mortality agent of larvae. Although at least five species of scoliids were introduced to the northeastern states, no parasitoids were successfully established. However, a survey of microbial pathogens in Connecticut (Hanula and Andreadis, 1988) demonstrated that gregarines infected up to 100% of the population and Paenibacillus popillae was found in up to 25% of the grubs sampled. Although the effect of these pathogens is not known, they likely help to suppress population densities of oriental beetle. Birds are sometimes observed to flock to grub-infested turfgrass, where they feed extensively on larvae. Weather also seems to limit the abundance of oriental beetle, and populations diminish during abnormally dry summers. Life Cycle and Description. Although a life cycle may be completed in only 3 months in Hawaii, there normally is but a single generation annually in the temperate climate of the northeastern states. In New England, some individuals even require a second year to complete their development. Reding and Klein (2007) reported that in Ohio 60% of the oriental beetles required 2 years to complete development. Overwintering normally occurs in the last larval instar. In Connecticut, larvae feed during April–June, and pupate in June. Pupae transform to adults in late June–August and emerge from the soil within a few days. Following mating and oviposition, eggs begin to hatch in July and August. Larvae usually attain the third instar by late September.

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Egg. The eggs are oval in shape. They typically measure about 1.2 mm wide and 1.5 mm long when first deposited, but swell slightly as they absorb water, and increase to 1.6 mm wide and 2.0 mm long. The period of incubation is about 17 days at 24°C, averaging 25–28 days under field conditions.

FIG.  5.94  Terminal abdominal segments of oriental beetle larva. (Drawing by USDA.)

Larva. Upon hatching the larva digs nearly to the surface to feed on roots and dead organic matter. Larvae display three instars. The first and second instars each require about 30–40 days to complete their development. The third instar overwinters, moving from a feeding position near the soil surface to greater depth in the soil, often 25–40 cm. Larvae are about 4 mm long following hatching and attain a length of about 4, 15, and 20–25 mm by the end of instars 1–3, respectively. Head capsule width is about 1.2, 1.9, and 2.9 mm, respectively, for the corresponding instars. The larva is similar in form to other scarab grubs—thick-bodied and C-shaped. Its body is whitish, though the head is brown. The body bears numerous short hairs, with the arrangement of hairs on the ventral surface of the terminal abdominal segment important for diagnostic purposes. Pupa. Pupation occurs in early June when larvae dig to a depth of 3–20 cm and create a small cell. The mature larva remains in the cell for 5–8 days before molting to a pupa. The duration of the pupal stage is about 15 days. The pupa is about 10 mm long and 5 mm wide, and light brown in color. The pupa generally resembles the adult though the wings are short and twisted to the ventral surface.

FIG. 5.95  Adults of oriental beetles. (Drawings by USDA.)

Adult. The adults are elongate-oval, and more tapered at the anterior end. The elytra bear pronounced ridges. The male beetles measure about 9 mm long and 4 mm wide; the females measure about 10.3 and 4.5 mm, respectively. The beetles are highly variable in coloration, ranging from yellowish brown to black. However, the head is invariably black, and the pronotum usually bears two moderately sized irregular black areas that coalesce into a single large spot in darker forms. The elytra often bear irregular black areas that form a V-shaped pattern, but as noted previously, coloration is not consistent in this species. Adults are active during morning and afternoon as well as at night. They may be attracted to flowers, where they feed slightly, or to lights. Females produce a sex pheromone that attracts males (Zhang et al., 1994). Both sexes mate repeatedly, and females commence oviposition within 1–5 days of mating. Oviposition normally continues for about 7 days (range 4–20 days). Females burrow into the soil, usually to a depth of 10–20 cm, to deposit eggs. Detailed description and biology were provided by Friend (1929). The larva of oriental beetle was included in the key by Ritcher (1966), and the adults in the keys of Downie and Arnett Jr. (1996).

Damage Damage is caused principally by the larval stage, which feeds on the roots of plants. Adults are generally considered to be of no consequence, though occasionally they feed on foliage of vegetable crops. Normally, only turfgrass experiences severe injury, but other plants are sometimes damaged. In the northeastern states, where oriental beetle co-occurs with Japanese beetle, Popillia japonica Newman, and Asiatic garden beetle, Maladera castanea (Arrow), the oriental beetle is sometimes the most important cause of turfgrass injury (Adams, 1949; Facundo et al., 1994).

Management Population density is usually assessed by examining areas of dead or dying sod for the presence of larvae. However, populations also may be monitored with sex pheromone-baited traps (Facundo et al., 1994; Alm et al., 1999). The pheromone can also be used to disrupt mating (Koppenhöffer et al., 2005) by competing with females for the attention of males (RodriguezSaona et al., 2010) though when evaluated in blueberrry plots a large number of pheromone releasers (at least 50 per hectare) or hand applied formulations are used (Rodriguez-Saona et al., 2009, 2010), which can be cost constraints. Applications of insecticide or Paenibacillus popillae to turfgrass are the principal methods of oriental beetle suppression. The susceptibility of oriental beetle grubs to P. popillae seems to vary over time, perhaps accounting for some of the variation in abundance (Dunbar and Beard, 1975). Compared to some other turf-damaging species, oriental beetle is relatively susceptible to most insecticides (Villani et al., 1988), though there may be site-related differences in field efficacy (Baker, 1986).

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Oriental beetle seems to be relatively nonsusceptible to some of the commonly available entomopathogenic nematodes (Steinernematidae, Heterorhabditidae) (Alm et  al., 1992), but Steinernema scarabaei is an exception, providing good suppression and persistence for scarab beetles (Koppenhöffer and Fuzy, 2003; Koppenhöffer et al., 2006), including oriental beetle. For example, Koppenhöffer and Fuzy (2009) reported 77%–100% suppression in turf plots within 1 month of application and 86%–100% suppression when assessed 3–4 years later.

Rose Chafer

Macrodactylus subspinosus (Fabricius)

Western Rose Chafer

Macrodactylus uniformis Horn (Coleoptera: Scarabaeidae)

Natural History Distribution. Rose chafer occurs principally in eastern North America, from Maine, Quebec, and Ontario south to Virginia, Georgia, and Tennessee. However, to a lesser degree, its range also extends west to Nebraska, and Oklahoma, and is replaced in the southwestern states by western rose chafer. These species are native to North America. Host Plants. Known mostly as a pest of rose, peony and grapes, rose chafer adults feed on a wide variety of plants, including vegetables such as asparagus, bean, beet, cabbage, corn, pepper, rhubarb, sweet potato, tomato, and perhaps others. Apple, blackberry, cherry, grape, peach, pear, plum and strawberry are among the fruits damaged. Flowers injured include dahlia, daisy, foxglove, geranium, hollyhock, iris, hydrangea, peony, poppy, and rose, though foxglove apparently is poisonous to beetles. Trees such as elm, magnolia, oak, sassafras, sumac, and others are also attacked; even conifers are not immune to attack. Larvae attack the roots of grasses and other plants. The dietary of western rose chafer is poorly documented but apparently is similar to rose chafer. Natural Enemies. The natural enemies are not well documented. General predators such as birds and toads are known to feed on rose chafer beetles, and ground beetles (Coleoptera: Carabidae) are thought to consume larvae, but there appear to be no records of parasitoids. It is difficult to imagine native species without a large complement of natural enemies, so the absence of records must simply reflect a lack of attention by entomologists in recent times. Life Cycle and Description. In New Jersey the winter is passed as a partly or fully grown larva, pupation occurs in April, adults emerge in late May and June, eggs are deposited in June–July, and larvae develop from July until cold weather drives them deep into the soil, usually in October–November.

Egg. The eggs reportedly are deposited in light, sandy soil or tilled soil, but not in dense sod or heavy, wet soil, because larvae are usually found only in the former environments. However, in preference tests, adults oviposited preferentially in wet soil and were not affected by soil texture (Allsopp et al., 1992a,b), so oviposition behavior is not fully known. The eggs are nearly spherical or oval, yellowish white, and measure about 0.8 mm in diameter. The adult deposits eggs singly along a burrow, which she digs belowground, sometimes to a depth of 10 cm, but also very near the soil surface. Fecundity is reported to be 24–36 eggs, all of which are deposited in a single burrow. Duration of the egg stage is about 21 days. Larva. Upon hatching, larvae may feed on organic matter in the soil but generally feed on roots. Larvae are yellowish white, with a tinge of blue toward the tip of the abdomen, well equipped with hairs, and attain a length of about 20 mm by autumn. The head is pale red. The body is C-shaped like most scarab beetles, though they are not as heavy-bodied as most white grubs. Larvae descend below the frost line in the winter, returning to the surface in the spring to feed for 2–4 weeks before pupating. Pupa. Pupation occurs in a small cell prepared by the larva near the soil surface. The pupa is yellowish white, about 15 mm long, and generally resembles the adult, though the wings are undeveloped and the legs are drawn up close to the underside of the body. Duration of the pupal stage is 2–4 weeks.

FIG. 5.96  Rose chafer adult. (Photo by P. Choate.)

Adult. The adults of both species are quite similar in appearance, though the western rose chafer averages slightly larger in size and has longer pubescence on the elytra. The rose chafer is fairly long and slender, measuring about 8–12 mm, though the very long legs give the impression of a much larger insect. The dorsal surface of the body is reddish brown but mostly covered with yellow hairs, imparting

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a tan color; it is either reddish brown or blackish below. The head and legs are reddish or brown, and the spines and claws reddish brown or blackish. The antennae are expanded terminally. The legs are well armed with spines and large terminal claws. The adults tend to be abundant for about a month, usually June, during which most damage occurs. The adults are active during daylight. Beetles mate soon after emerging from the soil, feed for about 7–10 days before beginning oviposition, and oviposition is completed in a few days. Thus, adult longevity is no > 3 weeks. The biology of rose chafer was given by Riley (1890), Smith (1891), Chittenden (1916b), and Lamson Jr. (1922).

Damage Most damage is caused by the adults, which while preferring to feed on blossoms, also consume fruit and foliage. Leaves are often skeletonized. Light or white-colored blossoms are preferred. Damage occurs mostly in areas with sandy soil, with beetles infrequent elsewhere. Damage to fruit causes the greatest degree of alarm among gardeners, though flower blossoms also may be shredded. Rose chafer is not generally known as a pest in commercial crops, though commercially produced grapes sometimes suffer. The beetles also are poisonous to chickens, with mortality due to a toxin, not to mechanical injury by the beetle’s long and sharp claws as was thought previously (Lamson Jr., 1922). Beetle larvae feed on the eggs of Melanoplus spp. grasshoppers, but the importance of this beneficial behavior is unknown.

Management Several chemicals have proven to be attractive to adults, and have the potential for development as lures. Among the attractive chemicals are caproic acid, hexanoic acid, valeric acid, octyl butyrate, nonyl butyrate, and others. Williams et al. (2000) claimed the development of a greatly improved five-component chemical lure. Heath et al. (2002) observed that plant volatiles released by beetle feeding increased beetle attraction. Lures are most effective when combined with a white trap (Williams and Miller, 1982; Williams et  al., 1990). Clean cultivation of crops is recommended not only because tillage can destroy stages in the soil, particularly pupae, but also because tillage deprives larvae of grasses and other preferred food. Insecticides applied to the foliage are usually recommended when adults are abundant.

Spring Rose Beetle

Strigoderma arboricola (Fabricius) (Coleoptera: Scarabaeidae)

Natural History Distribution. Spring rose beetle is a native species found in the northeastern United States from New York to

North Carolina, west to Colorado and Minnesota. In Canada it is known from southern Ontario. Host Plants. The adults feed on the blossoms of many plants. Wild and cultivated rose are favored food plants, which is the basis for its common name. Blossoms, and sometimes fruits, of other economically important plants such as blackberry, cotton, clover, coreopsis, hairy vetch, hollyhock, honeysuckle, iris, lilies, perennial pea, timothy, and peony are also consumed. Vegetable crops attacked by adults include bean, cantaloupe, corn, cowpea, and cucumber. Among weeds fed upon by adults are dewberry, Rubus spp.; elderberry, Sambucus canadensis; hoary verbena, Verbena stricta; horsemint, Mondarda punctata; plantain, Plantago spp.; prickly pear cactus, Opuntia humifusa; water lily, Nymphaea sp.; water willow, Justicia americana; wild parsnip, Pastinaca sativa, and others. Larval feeding habits are not well-known, but spring rose beetle larvae are reported to damage roots of corn, cotton, peanut, potato, soybean, strawberry, sweet potato, and various grasses and grain crops. Natural Enemies. Little is known about the natural enemies of this insect. An entomopathogenic nematode, Steinernema glaseri (Nematoda: Steinernematidae), is known to affect the larvae of this and other scarab beetles (Poinar Jr., 1978). Life Cycle and Description. There is a single generation per year, with overwintering occurring in the last larval instar. In Virginia, adults are abundant starting in May or June, while eggs are abundant in June. In Minnesota and Ontario, adults were reported to be common in June and July, while eggs were found in July and early August. Egg. The eggs are deposited singly in the soil at a depth of 5–10 cm. Grayson (1946) reported a fecundity of 30–40 eggs per female, but this seems to be an underestimate. The eggs are white and oval. Size varies because the eggs absorb water as they mature; the length increases from about 1.59 to 1.86 mm, the width from 1.07 to 1.63 mm. Duration of the egg stage is reported to average about 13.5 days (range 8–21 days). Larva. The larva is whitish, with a light brown head. There are three instars. Duration of the instars is about 16 days for each of the first two stages and 270 for the third, or overwintering, instar. The terminal portion of the last instar, or prepupal period, is relatively short, about 11 days. Larvae occur in the soil at a depth of 12–28 cm. Under warm conditions, larval development proceeds faster. Hoffmann (1936) reported larval development of about 160 days in the laboratory. Pupa. Pupation occurs in the soil, normally during April or May. Larvae form an earthen cell measuring about 30 mm long and 10 mm wide and pupate within. The pupa

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measures 9–12 mm long and gradually changes its color from light to dark brown as it matures. Duration of the pupal stage is usually 12–24 days.

FIG. 5.97  Adult spring rose beetle. (Drawing by J. Capinera.)

Adult. The beetles are moderate in size, measuring 8.5–12.5 mm long. They are somewhat variable in color, but the head and thorax are usually blackish green, and the elytra brownish yellow. The elytra are marked with distinct ridges. The underside of the beetles bears long hairs and is brown or brownish gray. Adults appear to persist for about 30 days. Mating is relatively brief in duration, requiring only 2–15 min. Mating is often observed on flowers, a favorite food for adults. The preoviposition period is 7–16 days, followed by oviposition in the soil. The biology of spring rose beetle was given by Hayes (1921), Hoffmann (1936), and Grayson (1946). The larvae were described by Ritcher (1966).

Damage This insect is known principally as a pest of roses, as the adults feed greedily on the blossoms. However, the larvae feed on the below-ground parts of several plants, and they are not infrequently associated with sweet potato and peanuts.

Management Historically, larvae are usually more abundant in low and heavy soil than in well-drained, sandy soil. Soils high in organic matter are also more prone to injury. Application of liquid or granular insecticide to the soil at planting-time or early in the growth of the crop has generally prevented injury. However, in Colorado injury has occurred in sandy soils, and it has proven difficult to attain protection with planting-time treatments because the insecticide dissipates before eggs hatch (Peairs, pers. comm.).

Sugarcane Beetle

Euetheola humilis (Burmeister) (Coleoptera: Scarabaeidae)

Natural History Distribution. Sugarcane beetle is found throughout the southeastern United States, though it is not a pest

e­verywhere. It can be damaging as far north as southern Ohio, and Maryland, and is found west to Texas and south to Florida. It is also found south into Central and South America. Sugarcane beetle is often known as Euetheola humilis rugiceps (LeConte). Earlier it was known as Ligyrus rugiceps LeConte. Thus, the literature can be confusing. Host Plants. The crops seriously affected have changed through the years, and E. humilis is clearly polyphagous. Initially (late 1800s) it was known as a pest of sugarcane, and perhaps rice (though this latter host is questionable). For much of the 1900s, it was known mostly as a pest of corn and called rough-headed corn stalk-beetle, Eutheola rugiceps (LeConte). Now it is more commonly known as a pest of sweet potato, as well as corn, though it has also been reported to affect tobacco, bermudagrass, zoysiagrass, tall fescue, rose, and strawberry. Apparently, its normal hosts are species of grass in the genus Paspalum (Poaceae), especially P. laeve (field paspalum) and P. plenipilum. Oddly, in recent year it has also become something of an urban pest, damaging expansion joint material, caulking, and running tracks. Young larvae seem unable to feed on seeds, roots, and plantlets, instead of developing on “vegetable mold,” the partly decomposed culms of common rush, Juncus effusus (Juncaceae) (Phillips and Fox, 1924). Once the larvae are about half-grown, however, they can feed on corn kernels that were planted as seed and are moist. Phillips and Fox (1924) concluded that decayed and disintegrated vegetable matter is the normal food of larvae. Accumulation of vegetable matter typically occurs in low areas and the edges of marshes, which coincides with the occurrence of Paspalum spp., favored food plants of the adults. Thus, the most common habitat of this insect is open grasslands on low or poorly drained soil. Likewise, it is the heavier, wetter areas of cropland that support larger numbers of the beetle. Natural Enemies. This aspect of sugarcane beetle biology is poorly studied. Vertebrate predators such as crows, meadowlarks, bluebirds, and skunks have been reported to feed on the larvae, but their importance has not been documented. Generalist predators such as robber flies (Diptera: Assilidae), ground beetles (Coleoptera: Carabidae), and ants (Hymenoptera: Formicidae) affect grubs (larvae). Insect parasitoids such as the tachinid fly Megapariopsis opaca (Coquillet) (Diptera: Tachinidae), the mite Rhizoglyphus phylloxerae Riley (Acari: Acaridae), and entomopathogenic fungi (including Beauveria bassiana) and nematodes have been observed to affect this insect, but neither specific determinations nor biological impact of these species is always available. Life Cycle and Description. The insects have a life cycle duration of about 85 days during the summer months. Apparently, they have one generation per year and overwinter as adults. Upon emerging from their overwintering state, they feed on grasses, particularly Paspalum spp.

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Oviposition can occur from June to August, and adults are present from April to October. Adults are nocturnal, and commonly found to be attracted to lights at night, but during the daytime they can be occasionally found crawling on the soil surface. Most of their life is spent below-ground, but because the adults overwinter, there are fall and spring flights of this insect.

antennal structures are apparent, only the wings are abnormally small. The development time for this stage was reported as 10 days at warm temperatures and about 17 days at cooler temperatures; 10–19 days is normal. Phillips and Fox (1924) illustrate the sexual characters used to distinguish the male and female pupae. There is no evidence that the pupal stage can overwinter.

Egg. The smooth, white eggs are deposited singly in the soil, several cms below the surface. Although several eggs may be found in the same vicinity at times, these insects do not deposit an egg mass or cluster, and more commonly are found scattered. Initially, the oval eggs measure about 1.6 mm long, 1.3 mm wide, and 1.5 mg in weight, but they enlarge over time, and after 10 days they are about 2.4 mm long, 2.0 mm wide, and 4.5 mg in weight. The eggs must be kept moist for hatching to occur, though overly wet conditions are not favorable. Baerg (1942) reported that females could produce about 100 eggs over their lifespan, although acknowledging that this is probably an underestimate; Phillips and Fox (1924) estimated oviposition at 1–2 eggs per day. Under field condition, an incubation period of about 10–15 days was normal during the summer months in Arkansas. In contrast, Phillips and Fox (1924) suggested that 14–21 days were required for egg hatch in Virginia. In Virginia, oviposition occurred in June after mating was observed in May and June.

Adult. The adult stage is glossy reddish black when first emerging, turning black within 4 or 5 days (though longer under cooler conditions). It measures 13–16 mm in length. Later it becomes dull black. The pronotum is quadrate, but the anterolateral angles each tape to a point. The posterolateral angles of the pronotum are broadly rounded. Overall, the pronotum is wider than long. Together the elytra are about as wide as long, resulting in a rather stout-looking beetle. The elytra are marked with double rows of punctures that give the elytra a striate appearance. The forelegs are stout and broad, well adapted for digging. The tibiae bear four tooth-like projections, three of which are long. The tibiae of the second and third legs bear a stout spine apically. This beetle is similar to other scarabs, especially carrot beetle, Bothynus gibbosus (De Geer) (Scarabaeidae). Phillips and Fox (1924) and Baerg (1942) picture this beetle, including characters that can be used to distinguish the sexes. Bieleisen and Brandenburg (2014) discuss separation of sugarcane beetle from similar species and suggest that head characteristics are most useful in the field. Sugarcane beetle is predominantly nocturnal, but occasionally is seen above-ground during the day.

Larva. The larva (white grub) has the typical robust body form associated with scarab beetles. There are three larval instars. Distinguishing characteristics include the top of the head being highly punctate, and the ventral surface of the last abdominal segment bears an irregular, median, double row of bristles. Although this species resembles Phyllophaga and Anomala spp., the latter species lack the punctate head capsule, and the double row of raster bristles is reported to be regular in occurrence and more conspicuous. As with other white grubs, the body color is generally whitish except for the posterior section, which is gray or brown. The head is orange-red, the legs yellow and the spiracles orange. When first hatched, the larva measures about 3 mm long. At maturity, the larva measures about 32 mm long and 6 mm wide. Baerg (1942) reported mean development times 14.4, 14.3, and 26.4 days for instars 1–3, respectively, when held at 31–35°C. The latter figure includes about 7 days in the prepupal stage, a relatively quiescent period that ends with the appearance of the pupa. Development time was longer when held at cooler temperatures. Overall, a larval development period of 45–65 days was commonly observed. There is no evidence that the larval stage can overwinter. Pupa. The whitish pupa soon turns brown and measures about 15 mm long. As is usually the case with beetles, the form of the pupa resembles the adult; the legs, mouth and

Damage As noted previously, the larval stage seems not to damage crops; only the adult stage is damaging. The adults cause damage to grasses and corn by forcing their way beneath the tufts or coming up under them from below and boring into the culms. Beetles feeding on sugar cane damage the plants early in the season, normally before the apical meristem emerges from the soil, but up to 1.2 m tall, by burrowing into the soil and attacking the stalk beneath the surface of the soil, and chewing a ragged hole in the plant. When feeding on corn, most of the damage caused by sugarcane beetles occurs early in the life of corn plants, when they are 7–15 cm tall. The beetle burrows through the soil and chews a cavity in the base of the plant belowground. Often the first sign of attack is plant wilting, and usually, the plants do not mature and do not produce an ear. However, if the plants are not yet sprouted, the beetle also may attack, feeding on the kernels and preventing aboveground growth. In Arkansas, the period of damage was reported to be April–June. When attacking sweet potato tubers, the beetles produce large holes and scars on the surface of the tuber. This happens mostly in the fall, as this is when the tubers mature.

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Turfgrass is injured mostly by the adults as they attack the stems at the soil surface, killing the plants. As noted previously, larvae feed on decaying plant material in the soil. However, they may incidentally damage roots of turf grasses, weakening the stand when grub densities are high.

Management Sampling. Visual examination of dying plants can be used to diagnose infestation by sugarcane beetle. Also, the beetles are highly attracted to light, so insect light traps are used to monitor the overall abundance of the beetles. Light traps used in late March and in April can aid in early detection of problems. Insecticides. Most corn and sweet potato growers who are concerned about this insect depend on persistent insecticides to prevent damage or to cure problems, although once the infestation exists curing the problem is very difficult. Insecticides can be applied as seed treatments, in-furrow liquid insecticide application, application of granular insecticides into the furrow or with incorporation into the soil, or granule application on the top of the soil. Cultural Practices. Several cultural practices may help prevent damage from sugarcane beetle. Elimination of dormant crop fields and pastures helps alleviate damage because grubs thrive on Paspalum spp. grasses. Planting is low and wet fields should be avoided, as egg and grub survival is higher under moist conditions. Frequent cultivation is beneficial because tillage can destroy insects near the soil surface, and because cultivation can help dry out the soil, reducing survival of the moisture-susceptible stages, particularly eggs. Finally, early planting helps reduce injury to sweet potato roots because tubers can be harvested before large numbers of adults are present.

White Grubs

Phyllophaga and others (Coleoptera: Scarabaeidae)

Natural History Distribution. The whitish, soil-dwelling larvae of scarab beetle species are called white grubs. Most are in the genus, Phyllophaga, but sometimes included are some members of the genera Anomala, Cyclocephala, and Polyphylla. The adults are called June beetles, May beetles, chafers, or cockchafers. They are found throughout the United States and southern Canada but are most abundant in the east of the Rocky Mountains. Nearly all species are native. Among the most damaging species are common June beetle, Phyllophaga anxia (LeConte); common cockchafer, P. fervida (Fabricius); northern June beetle, P. fusca (Froelich); fever June beetle, P. futilis (LeConte); southern June beetle, P. inversa (Horn); wheat white grub, P. lanceolata (Say); and tenlined June beetle, Polyphylla decemlineata (Say).

Host Plants. Both larvae and adults can be damaging, but they usually have markedly different feeding preferences. Larvae are found in the soil and normally feed on the roots of grasses. Adults, however, usually feed on the leaves and flowers of deciduous trees and shrubs, particularly ash, elm, hackberry, hickory, locust, oak, poplar, walnut, and willow, but sometimes pine. Hammond (1948) gives a useful list of adult host plants for P. anxia (LeConte). Root crops such as beet, potato, and turnip are among the crops most susceptible to injury by larvae, but corn and strawberry are sometimes damaged, and the seedlings of virtually all vegetables are susceptible if grub density is high. Vegetables crops are not preferred oviposition sites, so damage usually occurs only when crops are planted into the land that recently has been grass sod, pasture, or sometimes grass crops such as timothy, small grains, or sorghum. Tree seedlings, especially conifers, are damaged in both northern and southern states when planted into grass sod or pasture. Alfalfa and clover seem to be avoided even though both crops often contain some grass. Adults disperse widely, and trees adjacent to grassland, especially when few in number, are sometimes injured by defoliation. Natural Enemies. A large number of predators, parasitoids, and pathogens of white grubs is known. Undoubtedly, many soil-inhabiting predators, especially beetles (Coleoptera), kill white grubs, but there is little documentation of this. Larvae of robber flies (Diptera: Asilidae) and beeflies (Diptera: Bombyliidae) are frequently found in association with grubs. Among the parasitoids are such tachinids as Cryptomeigenia theutis (Walker), Eutrixa exilis (Coquillett), Eutrixoides jonesii Walton, and Microphthalma spp. (all Diptera: Tachinidae); the light flies, Pyrgota spp. (Diptera: Pyrgotidae); and the wasps Pelecinus polyturator (Drury) (Hymenoptera: Pelecinidae) and Tiphia spp. (Hymenoptera: Tiphiidae). Of the parasitoids, only Tiphia spp. are documented to be of considerable importance, sometimes accounting for up to 50% mortality among grubs. Mites (Acari: several families) are commonly associated with grubs and beetles; eggs and pupae are generally free of mites. However, few of these mite associates thought to be are parasitic or detrimental to white grubs (Poprawski and Yule, 1992). All the common groups of insect pathogens have been found associated with white grubs, especially Metarhizium anisopliae and Beauveria bassiana fungi. There is little evidence that these pathogens, and many of the other associates of white grubs, have a significant impact of grub density. However, some entomopathogens seem to be important, at least for some species of hosts, and of nematodes. See the section on oriental beetle for examples, especially for the importance of gregarines and of the nematode Steinernema scarabaei. A review of many natural enemies of white grubs was published by Lim et al. (1981b), but there has been important research since that date. Variability in host susceptibility,

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pathogen efficacy, and environmental interactions with host and pathogen are evident. Life Cycle and Description. The life cycle of the various species ranges from 1 to 4 years, though in the Phyllophaga species it is 2–4 years, with the 3-year life cycle most frequent. Duration of the life cycle is often related to latitude. In the southern United States 2-year life cycles are common, whereas in the North Central states 3-year cycles predominate, and in northern states and Canada 3–4year cycles occur. The general 3-year life cycle starts with oviposition in the spring or early summer, followed by feeding and growth until autumn when cold weather induces a period of inactivity, and overwintering by second instar larvae. During the second year, feeding is resumed in the spring, the third instar is attained, and feeding continues until cold weather when another period of inactivity occurs. In the spring of the third year, larval growth is completed and pupation occurs during the summer. The adult remains in the soil until the following spring. Thus, the 3-year life cycle occurs during a 4-year calendar period. In the typical 2-year life cycle, oviposition occurs in the summer, the eggs hatch in late summer, and young larvae commence feeding, quickly attaining the third larval instar and overwinter. They feed again throughout the second summer and overwinter as mature third instars. The following spring they feed briefly, pupate, and adults emerge, mate, and oviposit. Thus, in the 2-year life cycle larvae develop very quickly and the adult does not undergo a period of arrested development, as occurs in the 3-year cycle. The 2-year period of development occurs over three calendar years. Following is a description of common June beetle, P. anxia, perhaps the most abundant and damaging Phyllophaga in northern areas, and known from nearly all of Canada and the United States. It serves well to illustrate the biology of white grubs and June beetles. In Quebec, common June beetle displays the typical 3-year life cycle. The adults begin to fly in May and oviposit in June. First instar grubs are abundant in July and molt into second instars in August. The second instars overwinter and molt into third instars during June of the second year. Third instars overwinter at the end of the second year, developing into prepupae during June of the third year, and pupating in July. They are fully formed by September but remain in the soil until the following spring (Lim et al., 1981a). As noted earlier, the 3-year life cycle is spread over four calendar years. Additional information on the life cycle of the P. anxia life cycle in southern Canada is found in Hammond (1948). Egg. About 10 days after mating the female deposits pearly white and elongate-oval eggs about 2.4 mm long and 1.5 mm wide. The eggs absorb water, becoming enlarged

in size, and eventually measure about 3 mm long and 2 mm wide. Mean fecundity is about 55 eggs per female. Eggs hatch after about 20–30 days.

FIG.  5.98  Terminal abdominal segments of white grub. (Drawing by USDA.)

FIG. 5.99  White grub larvae. (Photo by L. Buss.)

Larva. Young grubs often feed on decaying vegetation before turning to root feeding, but they are not associated with manure. The grubs are pearly white, with a dark head and legs. They bear a thin covering of stout hairs, and when dug from the soil, they assume a curved-body posture or C-shape. Although the grubs feed at a depth of perhaps 5–15 cm during the summer months, they descend to about 30–100 cm during the winter months. Depth of burrowing is markedly affected by soil and moisture conditions and the age of the grub. Survival of grubs is higher in light soils and in moderate moisture conditions; heavy rain is detrimental both to oviposition and survival of young grubs. Grubs pass through three instars over about a 24-month period that is spread over part of three calendar years (Lim et al., 1980a). Pupa. The larva prepares a cell in the soil for pupation. This cell also serves as the overwintering site for the adult, which does not emerge from the soil until the following spring.

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p­ rotracted emergence by adults and the defoliation of trees is less apparent. Adults prefer young foliage, feeding from the edge toward the center of the leaves. They feed at night and hide during the day, so the cause of damage is often overlooked.

Management

FIG. 5.100  White grub beetle (June or May beetle). (Photo by P. Choate.)

Adult. The body of adult P. anxia is oblong-ovate, dark brown in color, and measures 17–21 mm long. The elytra are marked with numerous fine and shallow punctures. The underside of the thorax bears a dense coat of long hairs. The antennae are 10-segmented, elbowed, and expanded at the tip. The basal segments of the legs are generally stout, the tibiae are broadened and armed with teeth or stout spurs, but the tarsal segments are elongated and narrow. An excellent treatment of Phyllophaga biology, and a taxonomic treatment of adults was presented by Luginbill Sr. and Painter (1953). Other reliable sources of general biology, damage, and management information were found in Davis (1922), Luginbill (1938), and Ritcher (1940). The occurrence of white grub species in the North Central states was given by Pike et  al. (1977) and in the southeastern states by Forschler and Gardner (1990) and Kard and Hain (1990). The publication of Ritcher (1966) was an especially comprehensive treatment of grubs. Rearing techniques were reviewed by Lim et al. (1980a).

Damage White grubs are only occasional pests in vegetable production. Grubs have limited mobility; their distribution is largely a function of oviposition preference by females. Thus, though the roots and tubers of vegetables may be consumed, this is often due to conversion of grassland bearing partially grown grubs to vegetable production areas, and a decrease in the availability of their preferred host plants, or grasses. Typically, larvae clip the roots of plants with fibrous root systems, especially during the second year of larval life, resulting in the wilting and death of young plants. However, they may chew holes into larger roots and tubers, damage which may not be apparent until harvest. Damage to trees by adults is more frequent in northern states. In such areas the transition from cold, wet weather to warm, dry weather is abrupt, causing mass emergence of adults and defoliation of nearby trees. In contrast, in the south, the gradual increase in temperatures causes a

Sampling. Estimates of grub density are often made by examining 0.3 m cubes of sod and soil. Owing to the highly aggregated distribution of grubs and eggs (Guppy and Harcourt, 1970), however, extensive sampling is required to obtain a high degree of precision, often 50–100 samples (Guppy and Harcourt, 1973). Light traps are used to monitor populations of June beetles because they are nocturnal; such traps collect mostly males. Sex pheromone produced by the females, and attractive to males, is documented (Ochieng et al., 2002). Flights normally occur between sunset and midnight, and then again at sunrise, and are initiated by a combination of temperature and photoperiod stimuli (Guppy, 1982). Examples of the results of light-traps studies can be found in Kard and Hain (1990), Forschler and Gardner (1991), and Dahl and Mahr (1991). Insecticides. White grubs and June beetles are controlled with insecticides by application to soil or tree foliage, respectively. Soil applications predominate except perhaps for protection of foliage in fruit crops. Persistent insecticides are used in soil environments, applied preplanting (Rolston and Barlow, 1980) or at planting (Rivers et al., 1977). Insecticide resistance has developed in some regions (Lim et al., 1980b). Cultural Practices. Because of the attraction of June beetles to grasses, particularly short or mowed grass, there is a considerable risk when crops are planted into land that supported grass during the previous year, or when crops are grown adjacent to grass pastures or sod. If clover is planted immediately following grass, the risk to subsequent crops is reduced, partly because grubs rarely injure clover, but also because June beetles prefer not to oviposit in clover fields. It is also important that crops be free of grass and other weeds during the flight of June beetles, or considerable oviposition may occur. However, if grubs are already in the soil, the presence of grass weeds can be advantageous because grubs will feed preferentially on the grass (Rivers et al., 1977). Plowing and disking are sometimes recommended for destruction of grubs because the soft bodies of larvae are easily damaged by tillage. Also, tillage exposes grubs to birds, which can often be seen following tractors and consuming large numbers of exposed insects. Host-Plant Resistance. There is not much useful information on the varietal susceptibility of vegetable crops to white grubs. An exception is sweet potato, where not only has resistance been demonstrated (Rolston et al., 1981), but it has been shown that interplanting a resistant cultivar with a susceptible cultivar confers a reduction in damage to the susceptible variety (Schalk et al., 1991, 1992).

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Biological Control. There have been few attempts to implement biological suppression of white grubs, other than the use of domestic animals such as poultry and swine to consume larvae. The spore-forming bacteria Bacillus popilliae and Bacillus lentimorbus can infect and kill white grubs, but this expensive treatment is usually reserved for use on turf, where the bacteria can recycle through generation after generation of grubs. Entomopathogenic nematodes (Nematoda: Steinernematidae and Heterorhabditidae) have been evaluated for white grub suppression. Grubs are susceptible to infection and high levels of suppression have been attained by both injection and surface application, but the results are not consistent (Kard et al., 1988). The considerable variation in grub susceptibility to biological control agents is attributed to inherent differences in susceptibility, differences in environmental factors from place to plcace and time to time, and to changes in host susceptibility and biological control agent efficacy due to natural selection.

FAMILY TENEBRIONIDAE—DARKING BEETLES AND FALSE WIREWORMS False Wireworms

Blapstinus, Coniontis, Eleodes, and Ulus spp. (Coleoptera: Tenebrionidae)

Natural History Distribution. Several native species of Tenebrionidae damage vegetable crops in the western United States and Canada. The affected area is west of the Mississippi River, and invariably is arid. Although numerous species may be involved, the common species and area where damage has been reported are: Blapstinus elongatus Casey (California), B. fuliginosus Casey (California), B. pimalis Casey (Arizona, California), B. rufipes Casey (California), B. substriatus Champion (Montana), Coniontis globulina Casey (California), C. muscula Blaisdell (California), C. subpubescens LeConte (California), Eleodes hispilabris Say (Alberta, Washington), E. omissa (Say) (California), E.  tricostatus (Say) (western United States and Canada), and Ulus crassus (LeConte) (California). The list of damaging species is likely longer, but species-level identifications are not usually made. In fact, these insects commonly are confused with wireworms (Coleoptera: Elateridae), a similar and much more common group of pests. Host Plants. False wireworms are associated with grassland environments, and both larvae (called false wireworms) and adults (called darkling beetles) feed on seeds and seedlings of grasses and grain crops. Most false wireworm species, including many which are not mentioned above, are best known as wheat pests, principally because this is the crop most often grown in the arid areas inhabited by these insects. However, when irrigation is applied and vegetable crops cultured, false wireworms may also cause

injury to vegetables. Among the vegetables reported injured are asparagus, cabbage, cantaloupe, corn, lettuce, lima bean, mustard, onion, pepper, potato, radish, snap bean, tomato, and watermelon. Natural Enemies. Many parasitoids and diseases are known from false wireworms and darkling beetles, but the significance of these natural enemies is not always apparent. Flesh flies (Diptera: Sarcophagidae), particularly Blaexoxipha eleodes (Aldrich), are known from several Eleodes spp. Eleodes spp. are also parasitized by Eleodiphaga and Sitophaga spp. (Diptera: Tachinidae) and Microctonus eleodis (Viereck) (Hymenoptera: Braconidae). The fungi Beauveria bassianna and Metarhizium anisopliae occasionally are associated with false wireworm larvae (Allsopp, 1980). Rodents are important predators of darkling beetles in grass and shrub ecosystems, but not all rodent species consume them regularly. Work in Utah indicated that though northern grasshopper mouse, Onychomys leucogaster arcticeps, and omnivorous deer mouse, Peromyscus maniculatus nebrascensis, were effective predators, Uinta ground squirrel, Spermophilus armatus, and Great Basin pocket mouse, Perognathus parvus clarus, developed an aversion to the beetles after several feedings (Parmenter and McMahon, 1988). Birds feed readily on larvae and pupae, and often follow tractors as the soil is tilled, and capture the exposed insects. Larvae also can be forced to the soil surface when it becomes saturated by heavy rain, and birds are frequently observed to take advantage of this temporary abundance of food. Consumption of the adult stage is more infrequent, but some species are adapted to consume these insects. Among the species which feed on darkling beetles most frequently are bronzed grackle, Quiscalus quiscula seneus; western crow, Corvus brachyrhynchos hesperis; western robin, Planesticus migratorius propinquus; and sage hen, Centrocercus urophasianus (Hyslop, 1912c). Life Cycle and Description. The aforementioned false wireworm species have, at most, a single generation per year. Most species seem to require 2–3 years to complete their development. False wireworms generally overwinter as larvae, but some species pass the winter as adults. Some apparently overwinter in both stages. The life cycle of Eleodes hispilabris, one of the most common species that affect vegetables, is fairly typical of false wireworms. Adults pass the winter under dense masses of leaf debris, in cracks in the soil, and in rodent burrows. In the spring the adults feed on young weeds. In June they commence egg laying, with eggs deposited just beneath the soil surface. The eggs hatch in July, with the larvae about one-half grown at the onset of winter. The larvae commence feeding again in the spring and pupate in the soil in about August. They have about 11 instars, and the duration of the larval stage is about 270 days. The adults emerge in the autumn, feed briefly, and then overwinter, completing the 2-year developmental cycle.

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Damage

FIG.  5.101  Adult of a darkling beetle (false wireworm). (Drawing by USDA.)

The species differ somewhat in appearance, but the following description of Eleodes letcheri Blaisdell is typical. The egg is oval, measuring about 1.1 mm long and 0.6 mm wide. It is white, and free of sculpturing. Larvae are elongate-cylindrical, and somewhat flattened ventrally. Larvae are yellowish, but the head, mandibles, tarsi, and anal segment are brown to black. The antennae are short, enlarged or clubbed at the tip and yellow in color. The enlarged antennae of false wireworms serve to differentiate them from wireworms, which lack expanded antennae. The thoracic legs are stout. The larvae move freely and rapidly on the soil surface, a characteristic that helps distinguish them from wireworms, which are not agile on the soil surface. The pupa generally resembles the adult, except that the elytra are shrunken. Initially white in color, the pupal stage soon turns black. The adult is a shining black beetle with punctate elytra. The legs are long and stout. The maximum width of the abdomen is considerably greater than the juncture of the thorax and abdomen, and the tip of the abdomen tapers to a point. The adults are easily distinguished from predatory ground beetles (Coleoptera: Carabidae), with which they could be confused because the apical segments of the antennae are enlarged in tenebrionids but not in carabids. Most adults measure about 2.5 cm long. They display a defensive posture that involves extending the hind legs and tilting the body forward so that the tip of the abdomen is greatly elevated. Release of chemical exudates usually accompanies the head-standing behavior. In some cases, the exudate is forcibly expelled, but more commonly it is simply released at the tip of the abdomen as a drop, or spreads over the posterior part of the body. The exudate usually contains toluquinone, ethyl-p-quinone, and often other chemical components, that help deter predation. A worldwide review of false wireworm biology was published by Allsopp (1980). Observations on several western species were provided by Wakeland (1926). Rearing was outlined by Matteson (1966) and Wright Jr. (1972).

These insect normally are pests of dryland agriculture, and particularly of fields that have recently been converted from grassland. However, as noted by Daniels (1977), irrigated crops can be invaded. Both larval and adult stages may injure vegetables, but such injury is infrequent. Larvae tend to reside in the soil near the surface and feed on roots, seeds, and the stems of seedlings. Farmers often first detect a false wireworm infestation when they observe an irregular or patchy stand of seedlings in a field; widespread or uniform damage is rare. The adults are more mobile and active above-ground, they often feed on aerial plant parts. For example, Campbell (1924) observed the beetles feeding on the stems of tomatoes, peppers, and lima beans— both young seedlings sprouting from seed and transplanted plants were injured. Such injury is usually inflicted at the soil surface, and sometimes several beetles aggregate and feed on a single plant, quickly girdling it. Older plants are quite resistant to attack. False wireworms are omnivorous. In addition to feeding on seeds and seedlings, they also may feed on tubers, leaf tissue, plant detritus, and insect cadavers. Toba (1985a) studied damage to potato in Washington and reported that the surface scarring inflicted on potatoes was so minor that it would not lower the grade, and reduce the value, of potatoes.

Management Sampling. The larval and adult populations are often assessed by direct count of the number of individuals per unit area. This count is often accompanied by soil sieving, because larvae are usually in the soil. The adults may also be censused by using pitfall traps. In irrigated fields, damage often occurs at the field margins as adults invade from nearby weedy or uncultivated areas. Therefore, field edges and adjacent areas warrant monitoring. Insecticides. Insecticides may be applied to seeds or around the base of young plants. Phytotoxicity is a consideration with seed treatment. Residual insecticides are desirable, but not always available when protecting against injury by false wireworm. Baits containing insecticides are often effective for adult suppression, and bait application can often be limited to field margins. Daniels (1977) reported variable responses to insecticides in Texas, but complete suppression with some materials. Cultural Practices. Soil characteristics are often linked to false wireworm problems, with damage most often occurring in dry, light, sandy, or sandy loam soil (Wade, 1921). However, research in South Dakota showed that though some species (e.g., E. hispilabris) were most frequent in such soil types, other species (e.g., E. tricostatus) occur across most soil types, including heavier clay soils (Calkins and Kirk, 1975).

Chapter 6

Order Dermaptera—Earwigs

European Earwig

Forficula auricularia Linnaeus (Dermaptera: Forficulidae)

Natural History Distribution. The European earwig was first observed in North America at Seattle, Washington in 1907. It spread quickly and was reported from Oregon in 1909, British Columbia in 1919, and California in 1923. It reached Rhode Island in 1911, New York in 1912, and most other provinces and northern states in the 1930s and 1940s. Now it occurs south to North Carolina, Arizona, and southern California, but owing to its preference for temperate climates it is unlikely to become abundant in the southeastern states. It is not very tolerant of arid environments, but survives where irrigation is practiced. The European earwig is native to Europe, western Asia, and northern Africa, but has also been introduced to Australia and New Zealand. There is some mitochondrial and biological evidence that the European earwig is a species complex consisting of at least two species. The species seem to occupy different habitats, and these data suggest at least two separate introductions of these earwigs to North America (Guillet et al., 2000). Other earwigs are abundant in North America, but few are as numerous as the European earwig, and none are nearly as damaging. The ringlegged earwig, Euborellia annulipes (Lucas) (discussed separately), and the African earwig, Euborellia cincticollis (Gerstaecker), are probably the only other species of concern to crop producers. The ringlegged earwig is widespread, and occasionally of concern, but the African earwig is limited to the southwestern states and generally of no consequence. Neither species is native to North America.

Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00006-6 © 2020 Elsevier Inc. All rights reserved.

Host Plants. This insect is omnivorous and feeds on several types of plant and animal matter. Although its ­predatory habits somewhat offset its phytophagous behavior, on occasion the European earwig can inflict significant injury to vegetables, fruit, and flowers. Bean, beet, cabbage, celery, chard, cauliflower, cucumber, lettuce, pea, potato, rhubarb, and tomato are among the vegetable crops sometimes injured. Seedlings and plants providing the earwigs with good shelter, such as the heads of cauliflower, the stem bases of chard, and the ears of corn, are particularly likely to be eaten, and also likely to be contaminated with fecal material. Among the flowers most often injured are the dahlia, carnation, pinks, sweet william, and zinnia. Ripe fruits such as apple, apricot, peach, plum, pear, and strawberry are sometimes reported to be damaged. The European earwig is reported to consume aphids, spiders, moth eggs, caterpillar pupae, leaf beetle eggs, psyllids, scale insects, spiders, and springtails as well as vegetable matter. Aphid consumption is especially frequent and welldocumented (McLeod and Chant, 1952; Buxton and Madge, 1976a, b; Mueller et al., 1988), but it is abundant in orchards where it can choose among numerous potential prey. However, unlike many orchard-dwelling predators, it does not overwinter on fruit trees (Horton et al., 2002). In addition to the higher plants mentioned above, earwigs consume algae and fungi, and often consume vegetable and animal matter in equal proportions (Buxton and Madge, 1976a; Suckling et al., 2006). Natural Enemies. There are several known natural enemies, including some that were imported from Europe in an attempt to limit the destructive habits of this earwig in North America. Some authors have suggested that the most important natural enemy is the European parasitoid Bigonicheta spinipennis (Meigen) (Diptera: Tachinidae), which has been reported to parasitize 10%–50% of the earwigs in British Columbia. Others, however, report a

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low incidence of p­ arasitism (Lamb and Wellington, 1975). Another fly, Ocytata pallipes (Fallén) (Diptera: Tachinidae), also was successfully established, but it causes little mortality. Under the cool, wet conditions of Oregon, Washington, and British Columbia, the fungi Erynia forficulae and ­Metarhizium ­anisopliae also infect earwigs (Crumb et  al., 1941; Ben-Ze’ev, 1986). The nematode Mermis nigrescens appears to be an important mortality factor in Ontario, where 10%–63% of earwigs were infected during a 2-year period (Wilson, 1971). However, this nematode has not been reported from earwigs elsewhere. Avian predation can be significant (Lamb, 1975). Life Cycle and Description. One generation is completed annually, and overwintering occurs in the adult stage. In British Columbia, the eggs are deposited in late winter, hatch in May, and nymphs attain the adult stage in August. Overwintering females may also produce an additional brood; these eggs hatch in June and also mature by the end of August (Lamb and Wellington, 1975). In Washington, these events occur about 1  month earlier (Crumb et  al., 1941). Only a single brood of eggs is produced in colder climates such as in Quebec (Tourneur and Gingras, 1992). Egg. The egg is pearly white and oval to elliptical. The egg measures 1.13 mm long and 0.85 mm wide when first deposited, but it absorbs water, swells, and nearly doubles in volume before hatching. The eggs are deposited in a cell in the soil, in a single cluster, usually within 5 cm of the surface. The mean number of eggs per cluster is reported to range from 30 to 60 eggs in the first cluster. The second cluster, if produced, contains only half as many eggs. The duration of the egg stage under winter field conditions in British Columbia averages 72.8 days (range 56–85 days). The second or spring brood of eggs requires only 20 days to hatch. They are attended by the female, which frequently moves the eggs around the cell, and apparently keeps mold from developing on the eggs (Buxton and Madge, 1974). Females guard their eggs from other earwigs and fight with any intruders. Nymph. The nymphal stages, four in number, have the same general form as adults except that the wings increase in size with maturity. The cerci are present in all instars, growing in size with each molt. The body color darkens, gradually changing from grayish brown to dark brown, as the nymph matures. The legs are pale throughout. The wing pads are first evident in instar four. The mean head capsule width is 0.91, 1.14, 1.5, and 1.9 mm in instars 1–4, respectively. The mean body length is 4.2, 6.0, 9.0, and 9–11 mm, respectively. The number of antennal segments is 8, 10, 11, and 12 in instars 1–4. The mean duration (range) of instars under laboratory temperatures of 15–21°C is 12.0 (11–15), 10.2 (8–14), 11.2 (9–15), and 16.2 (14–19) days for instars 1–4. However, the development time is considerably longer under field conditions, requiring 18–24, 14–21, 15–20, and

FIG. 6.1  Adult European earwig. (Photo by L. Buss.)

about 21 days for the corresponding instars. Young nymphs are guarded by the mother earwig, which remains in or near the cell where the eggs are deposited until the nymph’s second instar is attained. Adult. The adult normally measures 13–14 mm long, exclusive of the pincher-like cerci (forceps), though some individuals are markedly smaller. The head measures about 2.2 mm wide. Adults, including the legs, are dark brown or reddish brown, though paler ventrally. The antennae have 14 segments. They bear a set of cerci at the tip of the abdomen. The pronounced cerci are the most distinctive feature of earwigs; in the male the cerci are strongly curved, whereas in the female they curve only slightly. They can use the cerci in defense, twisting the abdomen forward over the head or sideways to engage an enemy, often another earwig. Despite the appearance of being wingless, adults bear long hind wings folded beneath the abbreviated forewings. Although rarely observed to fly, when ready to take flight the adults usually climb and take off from an elevated object. The hind wings are opened and closed quickly, so it is difficult to observe the wings. Earwigs are nocturnal, spending the day hidden under leaf debris, in cracks and crevices, and in other dark locations. Their nighttime activity is influenced by weather. Stable temperature encourages activity, and it is favored by higher minimum temperatures but is discouraged by higher maximum temperatures. High relative humidity seems to suppress movement, whereas higher wind velocities and greater cloud cover encouraged earwig activity (Chant and McLeod, 1952). They produce an

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aggregation pheromone in their feces that is attractive to both sexes and to nymphs, and release quinones as defensive chemicals from abdominal glands (Walker et al., 1993). Social behavior is weakly developed in the European earwig. Males and females mate in late summer or autumn, and then construct a subterranean tunnel (nest) in which they overwinter. The female drives the male from the nest at the time of oviposition. Eggs are manipulated frequently, the female apparently cleaning them to prevent growth of fungi. She will relocate the eggs in an attempt to provide optimal temperature and humidity. Although the female normally keeps the eggs in a pile, as the time for hatching approaches she spreads the eggs in a single layer. After hatching, females continue to guard the nymphs and provide them with food. Food is provided by females carrying objects into the nest, and by regurgitation. Thus, there is parental care, but no cooperative brood care (Lamb, 1976). Thus, the adults are generally considered to display subsocial behavior (Kölliker, 2007). The most comprehensive treatment of European earwig biology was provided by Crumb et  al. (1941), though the publications by Jones (1917a, b) and Fulton (1924) were informative. Lamb and Wellington (1974) described methods for rearing. Keys to the western earwig species were provided by Langston and Powell (1975). The eastern species were considered by Hoffmann (1987). A synopsis of the European earwig, including keys to related Canadian insects, was given by Vickery and Kevan (1985).

Damage The economic status of earwigs is subject to dispute. Undoubtedly, earwigs sometimes damage vegetable and flower crops, both by leaf consumption and fruit injury. Foliage injury is usually done in the form of several small holes. Tender foliage may be completely devoured except for major veins. However, the physical presence of earwigs as crop contaminants is perhaps even more important, because most people find their presence and odor repulsive. The annoyance associated with their presence is exacerbated by the tendency of earwigs to aggregate, often in association with human habitations; most people simply find them annoying. Their propensity to consume other insects, particularly aphids, is an important element in offsetting their reputation as a crop pest (Moerkens et  al., 2009; He et al., 2008). However, augmenting the earwig population by field release, and/or providing them with additional shelter to enhance survival, have had mixed success in suppressing aphid populations (Carroll and Hoyt, 1984; Carroll et al., 1985; Logan et al., 2007).

Management Sampling. Population monitoring can be accomplished with baits and traps. Small piles of baits which can be

checked during the evening, distributed in dense ­vegetation, often attract large numbers of earwigs. Wheat bran or ­oatmeal can serve as a bait. Shelter traps take advantage of the natural tendency of earwigs to hide in crevices and dark spots, and can be used to detect the presence of earwigs and to estimate abundance. Shelter traps take many forms, including rolled cardboard, tubes, and capped sleeves, and can be used with or without food to encourage their shelter (Suckling et al., 2006). Insecticides. Residual foliar insecticides and baits containing toxicant can be used to suppress earwigs. Of the numerous baits evaluated, Crumb et al. (1941) suggested that wheat bran flakes plus toxicant and a small amount of fish oil was optimal. Fulton (1924) believed fish oil was unnecessary but suggested the addition of glycerin and molasses. Commercial products are rarely formulated specifically for earwigs because they rarely are a severe problem. Rather, products sold for grasshoppers, cutworms, slugs, and sow bugs are applied for earwig control. Bait is most effective if applied in the evening. Cultural Practices. On residential property or in small gardens, persistent trapping can be used to reduce earwig abundance, though this approach is not effective if the initial earwig density is high. Boards placed on the soil are attractive to earwigs seeking shelter. Even more earwigs accumulate if there are narrow grooves or channels in the board. A moistened, rolled-up newspaper placed in the garden in the evening and disposed of in the morning makes a convenient earwig trap for home gardens. A particularly effective collecting technique is to fill a flowerpot with wood shavings and invert it over a short stake that has been driven into the soil. Traps can also be placed in trees because earwigs favor this habitat.

Ringlegged Earwig

Euborellia annulipes (Lucas)

African Earwig

Euborellia cicticollis (Gerstaecker) (Dermaptera: Carcinophoridae)

Natural History Distribution. First found in the United States in 1884, the ringlegged earwig now is widespread in southern states. It is also known from many northern states, and the southernmost portions of British Columbia, Ontario, and Quebec in Canada. The ringlegged earwig occurs in Hawaii and has been transported to most other areas of the world, including both tropical and temperate climates. Its origin is uncertain, but it is now found widely in Asia, Africa, and Europe, in addition to North America. In cooler climates, however, its occurrence may be limited to greenhouses.

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The origin of E. annulipes is uncertain, but it is now found widely in Asia, Africa, and Europe, in addition to North America. In cooler climates, however, its occurrence may be limited to greenhouses. It is very similar and closely related to the ringlegged earwig. Its distribution is much more restricted, however, and known only from California, Arizona, and Texas. Host Plants. The ringlegged earwig is omnivorous in its feeding habits, taking both plant and animal material readily. It occurs as a minor nuisance in southern vegetable gardens and in greenhouses, where it nibbles on succulent plants such as lettuce. It is also documented to feed on roots or tubers of radish, potato, and sweet potato, and the pods of peanuts, though this is infrequent and normally of little consequence. The ringlegged earwig is a voracious predator of insects and sow bugs. This predatory behavior probably offsets the small amount of damage done to plants. It also is highly cannibalistic. The diet of the African earwig is not well characterized. Damage to vegetables is, so far as is known, limited to melons. Both seedlings and ripe fruit are consumed; feeding on nearly mature fruit can result in holes and scars on the rind of the fruit. The African earwig, like the ringlegged earwig, will feed on other insects, particularly aphids. Natural Enemies. The natural enemies of the ringlegged earwig seem to be undocumented, though they are likely about the same as those attacking the European earwig, Forficula auricularia Linneaus. Cannibalism of eggs and nymphs by adults is an important mortality factor. Life Cycle and Description. This insect seems not to have been studied under field conditions. Three generations were observed under greenhouse conditions in Ohio—one each in the spring, autumn, and winter months. A complete generation can be completed in 61  days (Klostermeyer, 1942). Lemos et  al. (2003) studied development at 25°C and 30°C, and reported that female longevity was 198 and 149 days at 25°C and 30°C, respectively. Thus, under field conditions, it seems probable that at least two generations occur, one each in spring and autumn, at least in warm climates. Although gross reproduction was similar at these two temperatures (about 90 eggs), net reproduction was about 65 and 40 eggs. In Illinois, adults can be found throughout the year except during winter when adults seek shelter deep in the soil. The African earwig also is a long-lived insect; adults may live for as long as a year. They are thought to have one or two generations per year. Eggs are found in June to August in California, and nymphal instars are present throughout the year. Both nymphs and adults overwinter. These insects favor undisturbed areas such as field margins, rather than cultivated areas. They often are found beneath plant debris or in the soil to a depth of up to 5 cm.

Egg. The eggs of the ringlegged earwig are nearly spherical when first deposited, and measure about 0.75 mm in diameter. As the embryo develops, however, the egg becomes more elliptical, and attains a length of about 1.25 mm. The egg is creamy white initially, becoming brown as the embryo develops. Females deposit 1–7 clutches of eggs with a mean clutch size of about 50 eggs. Total fecundity is estimated at 100–200 eggs. The duration of the egg stage is 6–17 days. The eggs of the African earwig are similar, though they are larger and the clutch size is less, averaging about 25 eggs. Initially, the eggs measure about 1.15 mm long, but increase to 1.55 mm. Their mean weight increases from 0.504 to 1.174 mg during the 13–14 day period of development. As might be expected for soil-dwelling insects, the eggs must be held at 100% humidity if they are to develop successfully. Nymph. The nymphs of the ringlegged earwig greatly resemble the adult in form, differing primarily in size (although due to differences occurring in nutrition, size may overlap). Wing pads are absent. The head and abdomen are dark brown. The pronotum is considerably lighter in color, usually grayish or yellowish brown. The legs are whitish, with a dark ring around the femur. The cerci are moderately long, and not strongly curved. Normally, five instars are found, but six are observed occasionally. Instars are difficult to distinguish and no single character is completely diagnostic. The number of antennal segments is most useful, though this is mostly effective among the early instars. The number of antennal segments is about 8, 11, 13, 14–15, 15–16, and 14–17 in instars 1–6. The head capsule width is 0.62–0.75, 0.70–0.91, 0.83–1.09, 1.04–1.56, 1.22–1.56, and 1.40–1.72 mm for instars 1–6, respectively. The body length is 3.0–4.7, 3.9–6.9, 5.7–7.7, 6.7–10.8, 8.7–13.2, and 9.8–12.9 mm, respectively, for instars 1–6. When reared at 21–23°C, Bharadwaj (1966) reported mean development times of 11.8, 10.6, 13.4, 16.3, 20.1, and 27.0  days for instars 1–6, respectively, for a total of about 99  days. However, he observed a considerably shorter mean nymphal development period, 83.6 days, when cultured at 20–29°C. Klostermeyer (1942) was able to rear the nymphal stage through to the adult in as little as 45 days, with an average of 81 days, when cultured at 18–29°C. The African earwig displays 4–7 instars, with the number affected by temperature. When reared at 22.2–26.6°C, the number of instars was reported to be 4–6, whereas at 26.6– 29.4°C it was 5–6 instars, and at 32.2–33.3°C it was 6–7 instars. The mean development time of the instars was 19.1, 15.8, 17.9, 27.8, and 34.1 days for instars 1–5, respectively, at 22.2–26.6°C, whereas it was 11.7, 8.7, 9.9, 11.1, 13.5, 13.0, and 17.5  days for instars 1–7, respectively, at 32.2– 33.3°C. Thus, the total nymphal development time averaged 115.5 days at 22.2–26.6°C and 74.3 days at 32.2–33.3°C.

Order Dermaptera—Earwigs Chapter | 6  209

FIG. 6.2  Adult of the ringlegged earwig. (Photo by J. Castner.)

Adult. The ringlegged earwig adults are elongate, dark brown, and wingless. They measure 12–16 mm long, with females averaging slightly larger than the males. The legs are pale, usually with a dark band around the middle of the femur, and often tibia, of each leg. Adults generally bear 16 antennal segments. The leg bands are the basis for the common name, and are readily apparent. Cerci of the adults can be used to distinguish the sexes. In the male the cerci are more curved, with the right branch of the forceps turned sharply inward at the tip. Males also possess 10 abdominal segments, whereas females possess eight segments. African earwigs also are dark brown to blackish, but the status of their wings is variable. Some adults are wingless, lacking any modifications of the mesonotum and metanotum. Others are short winged (brachypterous), displaying abbreviated extensions on the mesonotum and/or the metanotum, but lacking fully developed tegmina on the mesonotum or wings on the metanotum. Finally, some adults are winged, bearing normal tegmina and fully developed hind wings. In California, the brachypterous adults are the normal form. African earwigs average about 14 mm in length. Males have 10 abdominal segments whereas females possess 8 abdominal segments. The antennae are brown except for segments 15 and 16, which usually are white. The tegmina are tan or dark brown, and the legs yellowish but bearing smoky markings on the femora and tibiae. As in the ringlegged earwig, the forceps of males often are asymmetrical, the right process bent or curved more than the left. Females have only slightly curved forceps, as do the nymphs. Distinguishing characteristics useful for separating the ringlegged and African earwigs include: winged specimens are always African; short-winged specimens are African; the antennae of the African earwig normally consist of 17–20 segments, whereas those of the ringlegged earwig normally consist of 15–16 segments (occasionally 14 or 17). If some segments are white, the species can usually be separated

on this basis (the ringlegged earwig may have one or two white segments from 11 to 13, whereas the African earwig may have 15 or 16 white segments. Ultimately, the genitalia can be examined to determine the species. Unfortunately, the antennae of these earwig species sometimes lack white segments. Earwigs are nocturnal. Mating occurs 1–2  days after attainment of the adult stage. Oviposition commences 10–15 days after mating, and requires about 3 days to complete. The adults construct a small cell in the soil in which eggs are deposited. The female drives the male from the oviposition chamber before the eggs are produced. The female protects the egg clutch from mites, fungi, and intruders, cleaning and relocating them if necessary. Maternal care decreases soon after nymphs hatch, disappearing after about 10 days. The female cannot tolerate the presence of her progeny once she begins production of a subsequent egg clutch. Adults are long-lived, capable of living over 200  days in the case of the ringlegged earwig and over a year in the case of the African earwig. The biology of the redlegged earwig was given by Klostermeyer (1942), Neiswander (1944), Bharadwaj (1966), and Langston and Powell (1975). The temperature effects on biology are supplied by Lemos et  al. (2003). Culture was described by Bharadwaj (1966). This earwig was included in the keys by Langston and Powell (1975) and Hoffmann (1987). Knabke and Grigarick (1971) provide the most detailed study of the African earwig.

Damage These earwigs cause little direct injury to growing vegetable crops, but can feed on both the above-ground and belowground portions of plants. More commonly, they serve as a contaminant of produce, sometimes defecating on leafy green vegetables. They also can cause injury to stored products such as potatoes and carrots, and are important contaminants of food processing plants (Gould, 1948). These earwigs are important insect predators, and are documented to feed on such diverse prey as caterpillars, beetle larvae, and leafhoppers. Unfortunately, there seem to be no quantitative data on their relative importance.

Management These earwigs rarely warrant suppression, but are easily killed by most residual insecticides. They also accept bait formulations consisting of wheat bran, molasses, and toxicant, as well as many other baits (Neiswander, 1944). For additional information on earwig damage and management, see the section on the European earwig, Forficula ­auricularia Linnaeus.

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Chapter 7

Order Diptera—Flies and Maggots

FAMILY AGROMYZIDAE—LEAFMINER FLIES American Serpentine Leafminer Liriomyza trifolii (Burgess) (Diptera: Agromyzidae)

Natural History Distribution. This native leafminer has long been found throughout the eastern United States and Canada, northern South America, and the Caribbean. However, it has been introduced into California, Europe, and elsewhere, resulting in a fairly cosmopolitan distribution. There is increasing international traffic in horticultural crops, particularly flowers, which is thought to be the basis for the expanding range of this species. Liriomyza trifolii readily infests greenhouses, so range expansion seems likely to continue, including some northern climates. As a vegetable pest, however, its occurrence is limited principally to tropical and subtropical regions. Liriomyza trifolii is considered to be one of the three most-damaging polyphagous leaf miners of horticultural crops. All originated in the New World but all have been spread widely. The other two important species are Liriomyza sativae Blanchard (vegetable leafminer) and Liriomyza langei (Frick) (pea leafminer). The taxonomy of this group was greatly confused until 1980; consequently, many records before this time were incorrect or unsubstantiated. Confusion with vegetable leafminer, L. sativae Blanchard, has been especially frequent. Host Plants. Liriomyza trifolii is perhaps best known as a pest of chrysanthemums and celery, but it is highly polyphagous. For example, Stegmaier (1966b) reported 55 hosts from Florida, including bean, beet, carrot, celery, cucumber, eggplant, lettuce, melon, onion, pea, pepper, potato, squash, and tomato. Flower crops that are readily infested and that are known to facilitate spread of this pest ­include chrysanthemum, gerbera, gypsophila, and marigold, but there are likely many other hosts, especially among the Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00007-8 © 2020 Elsevier Inc. All rights reserved.

f­amily Compositae. Numerous broad-leaved weed species support larval growth. Schuster et al. (1991b) found that the nightshade Solanum americanum; Spanish needles, Bidens alba; and pilewort, Erechtites hieracifolia were suitable weed hosts in Florida. Natural Enemies. In North America, parasitic wasps of the families Braconidae, Eulophidae, and Pteromalidae play an important role in natural control when insecticides are not used; they usually keep this insect at low levels of abundance. At least 14 parasitoid species are observed from Florida alone. Species of Eulophidae such as Diglyphus begina (Ashmead), D. intermedius (Girault), D.  pulchripes, and Chrysocharis parksi Crawford are predominantly found in most American studies, though their relative importance varies geographically and temporally (Minkenberg and van Lenteren, 1986). Many of the parasitoids attacking L. trifolii also attack L. sativae. Predators and diseases are not considered to be important, relative to parasitoids. However, both larvae and adults are susceptible to predation by a large number of general predators, particularly ants. Life Cycle and Description. Leafminers have a relatively short life cycle. The time required for a complete life cycle in warm environments such as California and Florida is often 21–28 days, so numerous generations can occur annually in warm climates. Leibee (1984) determined the growth at a constant temperature of 25°C and reported that about 19 days were required from oviposition to the emergence of the adult. Development rates increase with temperature up to about 30°C; temperatures above 30°C are usually unfavorable and lead to a high larval mortality rate. Minkenberg (1988) indicated that the egg stage required 2.7 days with the temperature at 25°C for the development. The three active larval instars required 1.4, 1.4, and 1.8 days, respectively. The time spent in the puparium was 9.3 days. Also, there was an adult preoviposition period that averaged 1.3 days. The temperature threshold for the development of the various stages is 6–10°C except egg laying which requires about 12°C. 211

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Egg. The eggs are deposited in the middle or lower stratum of plant foliage. The adult appears to avoid immature leaves. The female deposits the eggs from the lower surface of the leaf, but they are inserted just below the epidermis. Eggs are oval in shape and small in size, measuring about 1.0 mm long and 0.2 mm wide. Initially, they are clear but soon become creamy white. Larva. Body length and mouth parts can be used to differentiate instars; the latter is particularly useful. For instar one, the mean and range of body and mouth part (cephalopharyngeal skeleton) lengths are 0.39 (0.33–0.53) mm and 0.10 (0.08–0.11) mm, respectively. For instar two, the body and mouth part measurements are 1.00 (0.55–1.21) mm and 0.17 (0.15–0.18) mm, respectively. For instar three, the body and mouth part measurements are 1.99 (1.26–2.62) mm and 0.25 (0.22–0.31) mm, respectively. A fourth instar occurs between puparium formation and pupation; this is a nonfeeding stage and is usually ignored by authors (Parrella, 1987). Pupa. The larva normally leaves the leaf mine and drops to the soil to pupate (or more technically, pupariate). The puparium initially is yellowish, then golden brown, but turns into darker brown as it matures. It is oval and slightly flattened ventrally. It measures about 1.3–2.3 mm long and 0.5–0.75 mm wide. The puparium has posterior spiracles on a pronounced conical projection, where each spiracle has three distinct bulbs, of which are elongate.

leafminer, the mesonotum is shining black and the hind margin of the eyes is darker. The small size of this species serves to distinguish it from pea leafminer, Liriomyza huidobrensis (Blanchard), which has a wing length of 1.7– 2.25 mm. Also, the yellow femora of American serpentine leafminer help to separate it from pea leafminer, which has darker femora. Adult longevity is 13–18 days. Leibee (1984), working with celery as a host plant, estimated that oviposition occurred at a rate of 35–39 eggs per day, for a total fecundity of 200–400 eggs. Parrella et al. (1983) reported similar egg production rates on tomato, but lower total fecundity, because tomato is a less suitable larval host. The female makes numerous punctures of the leaf mesophyll with her ovipositor and uses these punctures for feeding and egg laying. The proportion of punctures receiving an egg is about 25% in chrysanthemum and celery, both favored hosts, but only about 10% in tomato, which is less suitable for larval survival and adult longevity. Although the female apparently feeds on the exuding sap at all wounds, she spends less time feeding on unfavorable hosts. Adults are weak fliers and often are blown by the wind. Males live only 2–3 days, possibly because they cannot puncture foliage and therefore feed less than females, whereas females usually survive for about a week. Typically, they feed and oviposit during much of the daylight hours, but especially near mid-day. A good summary of American serpentine leafminer biology was published by Minkenberg and van Lenteren (1986). Keys for the identification of agromyzid leafminers were provided by Spencer and Steyskal (1986).

Damage

FIG. 7.1  Adult of American serpentine leafminer. (Photo by L. Buss.)

Adult. The adults are small, measuring  65%) levels of parasitism. Adjuvants that enhance nematode survival increase levels of leafminer mortality (Broadbent and Olthof, 1995). Cultural Practices. Because broadleaf weeds and senescent crops may serve as sources of inoculum, destruction of weeds and deep plowing of crop residues are recommended. The adults experience difficulty in emerging from anything but a shallow layer of soil. Cultural practices such as mulching and staking of vegetables may influence both leafminers and their natural enemies. Price and Poe (1976) reported that leafminer were higher in numbers when tomatoes were grown with plastic mulch or tied to stakes. At least part of the reason seems to be due to lower parasitoid activity in plots where tomatoes were staked. However, Wolfenbarger and Moore (1968) reported that aluminum mulch seemed to repel leafminers in tomato and squash plantings, whereas Webb and Smith (1973) observed this not to be the case.

Asparagus Miner

Ophiomyia simplex (Loew) (Diptera: Agromyzidae)

Natural History Distribution. This fly is native to Europe, where it occurs widely. It was first noticed in North America in 1861 and probably was introduced with asparagus plants. By the late 1800s and early 1900s, it was found generally in New England and the Middle Atlantic States and had been distributed to some locations in the Great Lake states. The asparagus miner was detected in California in 1905. It is now believed to occur wherever asparagus is grown and is a worldwide problem. Host Plants. Asparagus miner has a very restricted host range, feeding only on asparagus. It is rarely associated with asparagus spears (young shoots); instead, it is principally found on older stalks bearing “ferns.” Asparagus miner attacks young plants, however, if they are old enough to bear ferns.

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Natural Enemies. Little is known about the natural enemies of asparagus miner in North America. Two parasitoids, Chorebus rondanii (Giard) (Hymenoptera: Braconidae) and Thinodytes cephalon (Walker) (Hymenoptera: Pteromalidae), are relatively important in the United States and also found in Europe (Krombein et al., 1979; Morrison et  al., 2014b). However, in Michigan, at least 12 species of parasitoids have been reared from asparagus miner pupae (Morrison et  al., 2014b). Barnes (1937a), Barnes and Walton (1934), and Szelényi (1973) noted other parasitoid species in Europe. Life Cycle and Description. There are two generations annually throughout the range of this insect, with the pupal stage overwintering. The adults first become apparent in May, and within a few days of emergence they copulate and begin oviposition. First-generation larvae are common in June, completing their development and pupating in July. By late July or early August, second-generation adults have emerged and begin producing eggs. Second-generation larvae begin to mature in late August and early September. The larvae form puparia in the autumn, remaining in diapause until April or May of the following year. The lower development threshold for asparagus miner is 12.1°C and about 1500–2000 day degrees are required to complete the life cycle (Morrison et al., 2014a). Egg. The female deposits eggs beneath the epidermis of stems, usually near the base of the plant. The presence of the egg in the plant tissue is not apparent. The egg is white and elongate oval. One end tends to be more pointed than the other. The egg measures about 0.48 mm long and 0.22 mm wide. The duration of the egg stage is estimated to be 7–14 days.

Larva. The larva is whitish. It tapers slightly at both ends, and the anterior end bears black mouthhooks, while the posterior end bears a pair of black spiracles. There are three instars, and their length is about 0.4, 2.0, and 3.5 mm, respectively. Barnes (1937a) gave characters to separate the instars. The duration of the larval stage has been poorly documented, but it seems to be 7–14 days. Pupa. Pupation occurs within the larval mine. Firstgeneration larvae usually pupate above-ground, whereas second-generation insect tends to pupate in the stalk or roots below-ground, often at a depth of 5 cm or more. The puparium initially is light-brown, becoming dark-brown with age. It is somewhat flattened in form and measures about 4–5 mm long. The puparium bears hooks at both ends. The duration of the pupal stage is 14–21 days during the summer, and several months during the winter.

FIG. 7.3  Adult of asparagus miner. (Drawing by USDA.)

Adult. The adult is shiny black in appearance. It measures 3–4 mm long, with a wing span of 5–6 mm. The wings are colorless. Fecundity is unknown. The biology of the flies was discussed by Fink (1913) for New York and Barnes (1937a) for England. Ferro and Gilbertson (1982) and Lampert et al. (1984) provided bionomics in Massachusetts and Michigan, respectively.

Damage

FIG. 7.2  Larva and pupa of asparagus miner. (Drawing by USDA.)

The larvae feed just beneath the surface of the stem, interfering with photosynthesis. Burrowing starts at above soil level and proceeds upwards, forming mines. If several larvae are present in the same stalk the feeding activity of the larvae will girdle the plant. Infested plants become yellow in color. Discoloration may occur only at the base when few flies are present, or the entire plant may be affected. Burrowing by larvae also extends downward and even includes feeding on the roots. Larvae do not seem to damage young spears. The flies do not readily oviposit on this young tissue, and if they do, the eggs and larvae are too small to be observed at harvest, so they are not culled.

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The importance of asparagus leaf miner has been disputed. Eichmann (1943), for example, dismissed the possibility of injury, including the potential of interaction with crown rot disease caused by Fusarium spp. Although the direct effects of asparagus miner on yield are likely minimal, it is now apparent that their involvement with the disease is significant. The fungi Fusarium moniliforme and F. oxysporum are associated with all life stages of asparagus leaf miners, and increased incidence of Fusarium is associated with stem mining. Commercial asparagus fields in Massachusetts were found by Gilbertson et  al. (1985) to harbor 1.9 mines per stem and 2.9 pupae per stem. A significant decline in commercial asparagus in northeastern states has been attributed to the Fusarium and asparagus miner problem. Similarly, Morrison et  al. (2011) attribute the “early decline” of asparagus, which is a worldwide problem, to Fusarium diseases associated with this fly.

Management Sampling. Lampert et al. (1984) suggested that the pupal stage, particularly autumn puparia, was the best sampling unit. They also presented information on optimal sample size for fly populations with different densities. Ferro and Suchak (1980) studied the methods of adult sampling and recommended yellow or lime green sticky stakes because of the relatively large number of flies captured with these techniques. Insecticides. Insecticides applied to the stems of asparagus plants reduced the abundance of asparagus miners and the incidence of crown rot disease and thereby increased crop yield (Damicone et al., 1987). Soil fumigation also enhances yield due primarily to the destruction of fungi, but this approach is very expensive and useful only for the plant establishment phase of asparagus production. Cultural Practices. Destruction of overwintering stalks is sometimes recommended because puparia are harbored in the stalks. However, this is not completely satisfactory because some flies are associated with the roots. Wild asparagus is a major source of flies, and roadsides and irrigation ditches that might harbor asparagus should be checked for plants, and efforts made to eradicate them.

leafminer has been reported from most regions of the United States and is found as far north as Manitoba, Canada. However, it occurs as a pest principally in southern states. It likely is not native to North America. Host Plants. Cabbage leafminer attacks many plants in the family Cruciferae, including broccoli, cabbage, cauliflower, radish, turnip, watercress, and various mustards (Stegmaier, 1967). It is also known from peas (Leguminosae) and nasturtium (Tropaeolaceae). Although it is the most common leafminer attacking crucifers, other species are sometimes found on these crops. Natural Enemies. Stegmaier (1967) reported three species of eulophid wasps parasitizing cabbage leafminer in Florida: Diaulinopsis callichroma Crawford, Chrysocharis sp., and Pnigalio sp. (all Hymenoptera: Eulophidae). Oatman and Platner (1969) found seven species of Eulophidae, as well as one species each of Pteromalidae, Cynipidae, and Braconidae attacking this leafminer in California. In the latter study, the proportion of flies parasitized was lowest in January (26%) and highest in October (84%). The eulophid Diglyphus begini (Ashmead) was the predominant parasitoid and attacked the larval stage. Life Cycle and Description. Cabbage leafminers breed continuously in tropical climates. In more temperate climates such as California, L. brassicae occurs throughout the year, but it is much more abundant during the summer period (Oatman and Platner, 1969). Egg. Most feeding and egg deposition occur beneath the leaves, and along the margins of the leaves. The shiny white egg measures 0.16 mm wide and 0.28 mm long and hatches in about 3 days.

Natural History

Larva. The small larva tunnels through the mesophyll tissue, initially forming an elongate mine on the lower surface of the leaf. Accumulations of fecal material are usually evident along the center of the leaf mine, though this is more evident as the larva matures and produces a wider mine. The duration of the three active instars is about 6–7 days, with their duration averaging about 2, 2, and 3 days, respectively. The mouthpart (cephalopharyngeal skeleton) length of these instars measures about 0.09, 0.18, and 0.28 mm, respectively. About half-way through the third instar, the feeding behavior changes with the larva moving from the lower leaf surface to the upper leaf surface to feed. As the larva approaches maturity, it cuts a slit on the surface of the leaf and drops to the ground to form a puparium. A fourth larval instar occurs between puparium formation and pupation, but no feeding occurs so this instar is generally overlooked (Parrella, 1987).

Distribution. The cabbage leafminer is common throughout the tropical areas of North and South America, Africa, Australia, Asia, and Oceania, but it also occurs in some temperate regions. In North America, cabbage

Pupa. The yellow-brown puparium measures about 1.75 mm long and 0.8 mm wide. The puparium requires about 8–10 days. The adult emerges from a slit in the

Cabbage Leafminer

Liriomyza brassicae (Riley) (Diptera: Agromyzidae)

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a­ nterior region of the puparium, usually in the afternoon or ­evening hours. Mating occurs in the morning following adult emergence.

Corn Blotch Leafminer

Agromyza parvicornis Loew (Diptera: Agromyzidae)

Natural History

FIG. 7.4  Adult cabbage leafminer. (Drawing by USDA.)

Adult. These small black and yellow flies are easily confused with vegetable leafminer, Liriomyza sativae Blanchard, but can be distinguished using the male genitalia (Spencer, 1981). As in vegetable leafminer, the hind margin of the eyes is black, but cabbage leafminer differs in that it has less yellow color on the mesonotum. Vegetable leafminer generally has yellow on > 25% of the mesonotum surface, whereas cabbage leafminer has  cucumber > snap bean. Various weeds and field crops may favor the survival of whiteflies during vegetablefree periods (Coudriet et al., 1986). Wild lettuce, Lactuca serriola, and sowthistle, Sonchus spp., are examples of suitable weed hosts. Cotton, soybean, and to a lesser extent ­alfalfa and peanut are field crop hosts. As with most insects, host preference and suitability for whitefly growth and survival are highly, but not perfectly, correlated (Zalom et al., 1995). Despite the lack of suitability among certain hosts for whitefly growth and reproduction, even less suitable vegetable crops such as lettuce can sometimes be damaged when large numbers of dispersing whiteflies feed and oviposit. This is especially likely to happen when suitable crops supporting numerous whiteflies senesce or are harvested, forcing the whiteflies to disperse in search of food. Survival of whitefly may be enhanced by feeding on virus-infected plants, relative to healthy plants (Costa et al., 1991). Deterioration of host-plant quality will often prompt the dispersal of adults. Weather. Sweetpotato whitefly thrives under hot and dry conditions. Rainfall seems to decrease populations though the mechanism is not known. Sweetpotato whitefly is not a strong flier, normally moving only short distances in search of young plant tissue. However, under proper weather conditions dramatic, long-distance flights involving millions or billions of insects are observed. Such flights normally occur in the morning as the sun heats the ground, and most insects move downwind. Natural Enemies. Numerous predators, parasitoids, and fungal diseases of sweetpotato whitefly are known (Gerling, 1990a; Gerling et al., 2001; Cock, 1993). The general predators usually associated with Homoptera such as minute pirate bugs (Hemiptera: Anthocoridae), green lacewings (Neuroptera: Chrysopidae), and lady beetles (Coleoptera: Coccinellidae) are important, as are many parasitic wasps, particularly in the genera Encarsia and Eretmocerus (both Hymenoptera: Aphelinidae) (Polaszek et al., 1992). While these agents exert considerable control on whitefly populations in weedy areas or on crops where insecticide use is minimal or absent, they do not survive well in the presence of most insecticides. Fungi assume importance when both whitefly densities and relative humidity are high. Life Cycle and Description. Sweetpotato whitefly can complete a generation in about 20–30 days under favorable

weather conditions (Gerling et al., 1986). In tropical countries, up to 15 generations per year have been reported, and in the southwestern United States, 12–13 generations occur annually. Egg. The egg is about 0.2 mm long, elongate, and tapers distally; it is attached to the plant by a short stalk. The whitish eggs turn brown before hatching, which occurs in 4–7 days. The female deposits 90%–95% of her eggs on the lower surfaces of young leaves (Simmons, 1994).

FIG. 8.4  Sweetpotato whitefly nymphs. (Photo by L. Buss.)

Nymph. All instars are translucent and somewhat shiny. The flattened first instar is mobile and is commonly called the “crawler” stage. It measures about 0.27 mm long and 0.15 mm wide. Its movement is usually limited to the first few hours after hatch and to a distance of 1–2 mm (Price and Taborsky, 1992). The duration of the first instar is usually 2–4 days. The feeding site is normally the lower surface of a leaf, but sometimes more than 50% of the nymphs are found on the upper surface, and feeding location seems not to affect survival (Simmons, 1999). The second and third instars are similarly flattened, but their leg segmentation becomes reduced and the legs nonfunctional. The duration of these instars is about 2–3 days for each. Body length and width are 0.36 and 0.22 mm, and 0.49 and 0.29 mm, for the second and third instar, respectively. Although the early portion of the fourth instar is similar to instars 2–3, the latter portion is sessile and called the “pupa.” This term is not technically correct because some feeding occurs during this instar. The appearance of the fourth instar is variable, depending on the food plant; this stage tends to be spiny when develops on a hairy leaf but has fewer filaments or spines when feeding on smooth leaves. The fourth instar measures about 0.7 mm long and 0.4 mm wide. The duration of the fourth instar is about 4–7 days. Total preadult development time averages 15–18  days in the temperature range of 25–32°C, but increases markedly at lower temperatures. The lower- and

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upper-developmental thresholds are considered to be about 10°C and 32°C (Natwick and Zalom, 1984).

potato whitefly developmental biology on tomato was ­published by Salas and Mendoza (1995). Silverleaf whitefly was described by Bellows et al. (1994). The specific status of these whiteflies was discussed by Brown et  al. (1995), Boykin et  al. (2007), De Barro et  al. (2011), Liu et  al. (2012a,b), and others. Whitefly that easily can be confused with B. tabaci is greenhouse whitefly, Trialeurodes vaporariorum (Westwood). They can be distinguished in the field by the manner in which they hold their wings. Sweetpotato whiteflies hold their wings roof-like over their body while at rest, whereas greenhouse whiteflies hold their wings horizontally when at rest.

Damage

FIG. 8.5  Pupa of sweetpotato whitefly. (Photo by L. Buss.)

The form of the pupa is used to distinguish among whitefly species and can be used to separate greenhouse whitefly from the similar-appearing greenhouse whitefly, Trialeurodes vaporariorum (Westwood), from the Bemisia spp. Greenhouse whitefly is straight sided when viewed laterally, ovoid, and lacks a groove near the anal end of the body. In contrast, the Bemisia spp. are oblique-sided, irregularly oval, and possess a groove in the anal region.

FIG.  8.6  Adults and immature stages of sweetpotato whitefly. (Photo by L. Buss.)

Adult. The adult is white and measures about 1.0– 1.3 mm long. The antennae are pronounced and the eyes red. Oviposition begins 2–5 days after the emergence of the adult, often at about 5 eggs per day. Adults typically live 10–20  days and may produce about 50–150 eggs, though there are records of over 300 eggs per female. Females may produce male offspring without fertilization but males are common, so most females are probably fertilized. The biology and management of sweetpotato whitefly were comprehensively reviewed by Butler et  al. (1986), Cock (1986, 1993), and Gerling (1990b). A study of sweet-

The adult and nymphal whiteflies use their piercing-­sucking mouthparts to feed on the phloem of host plants. This results in direct damage, which is manifested in localized spotting, yellowing, or leaf drop. Under heavy feeding pressure, wilting and severe growth reduction may occur. Systemic effects are also common, with uninfested leaves and other tissue being severely damaged by whitefly feeding on other areas of the plant. A translocated toxicogenic secretion by nymphs, but not by adults, is implicated (Yokomi et al., 1990). The young and developing tissue is damaged by whiteflies while feeding on older tissue. Once the whiteflies are removed, new plant growth is normal if a disease is not transmitted. Damaged foliar tissue, however, does not recover once injured. Among leafy vegetables and crucifers, white streaking or discoloration, especially of veins, is common (Brown et al., 1992; Costa et al., 1993). In Texas, population densities of three adult whiteflies per leaf are estimated to inflict a 10% yield reduction in cantaloupe (Riley and Sparks, 1993). Other studies in Texas and Arizona (Riley and Palumbo, 1995a,b) demonstrated similar losses and indicated that yields could be optimized if plants were treated with insecticide at whitefly densities of three adults per leaf or 0.5 large nymphs per 7.6 cm2 of leaf area. A disorder called irregular ripening affects tomato fruit when whiteflies feed on tomato foliage (Schuster et  al., 1990). Although the tomato foliage is not damaged, the internal portions of the fruit do not ripen properly and the surface is blotched or streaked with yellow. Squash silverleaf, a disorder induced by sweetpotato whitefly on squash, has been known from Israel since 1963 but did not occur in the United States until about 1986, when this whitefly first became abundant in Florida. Silverleaf symptomology includes blanching of the veins and petioles, and eventually the interveinal areas of the leaf. The fruit of both yellow- and green-fruited varieties also may be blanched (Yokomi et al., 1990; Schuster et al., 1991a). In addition to direct damage, sweetpotato whitefly also causes damage indirectly by transmitting plant viruses. Over 60 plant viruses, most belonging to a group called

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g­eminiviruses, are known to be transmitted to crops by sweetpotato whiteflies (Markham, 1994). Some viruses, such as tomato yellow leaf curl virus, cause more damage than the insect feeding alone, so the effects are devastating. Unfortunately, unlike the case with the phytotoxemia caused by the whitefly salivary secretions, once viruses are inoculated into the plant there is no recovery by the host even if the whiteflies are eliminated. Lastly, whiteflies cause injury by excreting excess water and sugar in the form of honeydew. This sticky substance accumulates on the upper surface of leaves and fruit and provides a substrate for growth of a fungus called sooty mold. The dark mold inhibits the photosynthetic activity of the foliage, and may also render the fruit unmarketable unless it can be washed thoroughly and the residues are removed.

Management Sampling. The distribution of whitefly life stages on cantaloupe was studied by Tonhasca et al. (1994) and Gould and Naranjo (1999). The eggs tend to be concentrated on young foliage and mature larvae on older foliage. Large nymphs are considered a good stage for population assessment because they cannot move and are large enough to see without magnification. Adults sometimes are concentrated on lower leaves, but they move to young foliage during oviposition. Such distributions must be considered in population assessment before initiating management practices. Ohnesorge and Rapp (1986) recommended yellow sticky traps rather than direct visual counts for population estimates when insect densities were low. However, when Palumbo et  al. (1995) evaluated several sampling methods in cantaloupe, they found that visual observation of the lower-leaf surface and vacuum sampling were less time consuming, and sometimes more precise, than yellow sticky traps. Sampling techniques were reviewed by Butler et al. (1986) and Naranjo (1996). Insecticides. In southern states, where sweetpotato whitefly can be the most important insect problem on some vegetable crops, frequent applications of insecticides are often made to minimize the direct and indirect effects of whitefly feeding. Whitefly resistance to nearly all classes of insecticides is known, and the rotation of insecticide classes is encouraged. Mixtures of insecticides are often used, which is indicative of high levels of resistance. Most agriculturalists suggest that whitefly numbers be maintained at low levels because once they become abundant they are difficult to suppress. This continued selective pressure on the insect populations exacerbates the development of insecticide resistance, of course, so it is highly desirable to substitute nonchemical control whenever possible. The phytotoxemia and disease transmission potential of this insect exaggerates its damage potential, further justifying the frequent application of insecticides. Often the most ­effective approach to

effective management involves regional or area-wide suppression based on a combination of insecticides, weed management, and crop management. Sweetpotato whitefly feeds on the lower surface of foliage and is sessile throughout most of its life—habits that minimize contact with insecticides. Frequent insecticide application also disrupts naturally occurring biological control agents. In an attempt to minimize the cost and disruptive effects of insecticides, and to reduce the evolution of insecticide resistance, surfactants such as soaps and oils have been extensively studied for whitefly control (Liu and Stansly, 2000). The mechanism of control by surfactants is not clearly understood, but disruption of the insect cuticle, physical damage, and repellency are postulated. In any event, mineral and vegetable oils alone, or in combination with soaps and detergents, can provide some suppression of whiteflies. A combination of insecticide and oil often enhances whitefly control (Horowitz et al., 1997). Suppression usually increases with the concentration of the surfactants, but 0.5% detergent plus 0.5% vegetable oil, or 0.5% detergent alone, or 1% insecticidal soap alone, or 0.75%–1.0% light-mineral oil are often recommended initially until the phytotoxicity potential is known. Oil has more residual activity than soaps or detergents; the former is also more repellent to adults (Liu and Stansly, 1995). Cucurbits and crucifers seem especially prone to foliage damage by surfactants, and damage occurs frequently under high-temperature conditions. High gallonage enhances coverage and pest population reduction but increases the cost of control (Butler and Henneberry, 1990a,b; Butler et  al., 1993). Neem products and other growth regulators affect the survival of immature insects only (Price and Schuster, 1991), and are most effective by foliar application, but seed and soil treatments are also effective (Kumar et al., 2005; Kumar and Poehling, 2006a,b). Plant essential oils have numerous effects on Bemisia biology (Yang et al., 2010a,b,c). Biological Control. Although many predators, parasitoids, and fungal diseases are known to attack sweetpotato whitefly, no biotic agents are known to provide adequate suppression alone. Under greenhouse conditions, parasitoids can be released at high enough densities to provide some suppression, especially when insecticidal soap and other management techniques are also used (Parrella et al., 1991). Under natural field conditions, parasitism does not usually build to high levels until late in the growing season (Cock, 1993). Insecticides often interfere with parasitoids and predators, of course, and effective use of biological control agents will probably be limited to cropping systems where broad-spectrum insecticide use is minimized, and other management techniques used that favor action of predators, parasitoids, and disease agents. Some new insecticides are quite selective, killing whiteflies and yet preserving most natural enemies. Lecanicillium (Verticillium), Paecilomyces, Beauveria, and other fungi similarly show

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some promise under greenhouse conditions but are limited by low humidity and economic constraints under field conditions (Meade and Byrne, 1991; Cock, 1993; Faria and Wraight, 2001). A summary of predators and parasitoids and their roll in biological control of B. tabaci was published by Gerling (1990a,b); and Gerling et al. (2001). Cultural Practices. Cultural controls can be vitally important in managing sweetpotato whitefly. Incorrect crop management, in particular, can create or exacerbate whitefly problems. Whiteflies can move from crop to crop, and areawide crop-free periods help diminish populations. Thus, prompt tillage of land and destruction of crop residues after crop maturity is recommended. Similarly, weeds can harbor whiteflies, whitefly-transmitted diseases, and whitefly parasitoids, so weed management is a consideration. Whitefly populations typically consist of a mixture of whiteflies that will disperse long distances and others that are trivial fliers that do not disperse long distances (Byrne et al., 1996). Row covers and other physical barriers can decrease the infestation of crops, and infection with the disease (Cohen and Berlinger, 1986). Screen hole sizes of about 0.19 mm2 or smaller are required to successfully exclude whiteflies (Bethke et al., 1994). Colored and aluminum mulches provide only a temporary reduction in whitefly abundance and disease transmission (Cohen and Berlinger, 1986) or none at all (Powell and Stoffella, 1993). Orozco-Santos et al. (1995) reported effective whitefly exclusion on cantaloupe by using row covers and reduction in whitefly population levels with transparent mulch. Protecting young plants from infestation with these techniques, and of course, using insectfree transplants, will minimize loss, in part because young plants (especially the first 25–30 days) are more susceptible to geminivirus infection (Hilje et al., 2001). Greenhouses can be used to reduce whitefly and virus loss. Solid plastic roof structures and either solid or screened walls are typically used to exclude whiteflies. Maintaining good airflow in the greenhouse is important, but the mesh must be fine enough to exclude insects. Addition of UV light absorbing agents to the plastic can interfere with orientation of the whiteflies and reduce insect abundance and damage. Host-Plant Resistance. Host-plant resistance offers considerable potential, but currently, it is difficult to put into practice. Both very hairy and hairless cultivars are perhaps less suitable for parasitoid activity than plants with intermediate densities of plant hairs. Resistance to the toxic saliva and to viruses transmitted by whiteflies also are being sought among commercially acceptable hybrids. For example, McCreight and Kishaba (1991) evaluated numerous cucurbit species and cultivars for susceptibility to squash leaf curl virus. Susceptibility varied among and within cultivated species, but overall susceptibility among species was: Cucurbita maxima > C. pepo > C. mixta > C. moschata. All cultivated species and cultivars were at least moderately

s­ usceptible to squash leaf curl virus, and most were severely damaged in field tests. In contrast, many wild cucurbit species were unaffected. Among commercial tomato cultivars, there was less oviposition by whiteflies on plants with low trichome densities, but this relationship was not apparent for wild tomatoes (Heinz and Zalom, 1995). As the basis for resistance is understood, there is a good possibility of incorporating at least partial resistance into commercial cultivars. Disease Transmission. Growers generally rely on whitefly suppression to manage disease incidence. This is not entirely satisfactory, however, and roguing of virusinfected plants is often suggested to minimize the withinfield spread of viruses. As whiteflies may transmit disease from one crop to another, or from weeds to crops, vegetation management is important. As noted above, reflective mulches have not produced consistent economic benefits. Mineral and vegetable oils may inhibit virus transmission. Application of 10%–15% commercial whitewash solution is reported to be as effective as mineral oil, and more effective than some pyrethroid insecticides, in reducing virus disease incidence; however, phytotoxicity is sometimes a problem (Marco, 1993). Row covers or other physical barriers can substantially prevent disease transmission (Costa et al., 1994), but often they are not economical. UV lightabsorbing plastic has been suggested as a preferred medium for plastic greenhouses (Antignus et al., 1996). Apparently, the elimination of UV light decreases the attraction of plants to whiteflies or changes their feeding behavior, thereby suppressing feeding and disease transmission.

FAMILY APHIDIDAE—APHIDS Artichoke Aphid

Capitophorus elaeagni (del Guercio) (Hemiptera: Aphididae)

Natural History Distribution. This species is found in temperate regions throughout the world but probably is of European origin. It is the most common and damaging aphid affecting artichoke in California. Host Plants. The summer hosts of this aphid are artichoke and Cirsium and Carduus thistles. The winter hosts are Elaeagnus spp. trees and shrubs, principally Russian olive, E. angustifolia. Natural Enemies. Natural enemies are not well known, but Lange (1941) reported that ladybird beetles (Coleoptera: Coccinellidae) and flower flies (Diptera: Syrphidae) commonly preyed on these aphids. In Japan, the ladybird beetle Harmonia axyridis Pallas drove an artichoke aphid population to extinction by the end of May (Osawa, 1992). A wasp parasitoid, Ephedrus persicae Froggott (Hymenoptera: Braconidae), is known. The fungi Beauveria bassiana and

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Bionectria ochroleuca have been observed to effectively suppress artichoke aphid when sprayed on plants in Tunisia (Guesmi-Jouini et  al., 2014), and Fusarium sacchari and F. semitectum occur there naturally on C. elaeagni (Jouda et al., 2010). The fungi Entomophthora planchoniana and Zoophthora radicans were found in association with this aphid in Argentina (Scorsetti et al., 2006). Life Cycle and Description. In temperate regions this aphid spends the winter on Elaeagnus spp., dispersing to artichoke or thistles in the spring. This pattern is documented in Colorado, where artichoke and Canada thistle, Cirsium arvense, are suitable summer hosts. However, under the mild climatic conditions of California, where most North American artichoke is grown, the artichoke aphid remains on artichoke throughout the year. This is a fairly small aphid, the females measure only about 1.6–2.0 mm long and the males 1.5–1.8 mm. In the wingless (apterous) viviparous form, the head and abdomen are greenish yellow or frosty green with some darker patches dorsolaterally. The legs, antennae, and cornicles are pale though often tinged with brown or gray. The cornicles are quite long. On summer hosts, the winged viviparous forms (alatae) are darker, bearing blackish antennae, head, and pronotum. The abdomen is yellowish green or green, but with a large dark square patch dorsally, and three dusky patches laterally. The long cornicles are pale or greenish, but the tips are darker. The wing veins are dusky brown. The sexual forms also are similar in color. In the wingless oviparous female, the yellowish body and head are suffused with reddish brown or pink. A pair of large dark markings appears dorsally on the abdomen, accompanied by 5–6 dark spots laterally. The legs and cornicles are yellowish, but the cornicles are darker apically. In the winged male, the color of the head, thorax, and antennae is dark. The abdomen is yellowish, sometimes suffused with reddish brown. The abdomen also bears a row of large dark spots dorsally and laterally. The cornicles are pale, but darker apically. The legs are brownish. A good description of this poorly studied aphid can be found in Cottier (1953). A key to artichoke-infesting aphids was included in Palmer (1952) and Blackman and Eastop (1984); in the latter, it was included in the key to artichoke.

Damage Artichoke aphid is principally numerous during the summer months when the populations completely cover the lower surface of foliage and cause wilting of the plant. Large quantities of honeydew are secreted, causing the plant to blacken. The effect of aphid feeding and honeydew secretion is to delay the maturation of the artichokes for up to several months (Lange, 1941). This species is capable of transmitting several plant viruses, including the artichoke latent virus (Kennedy et al., 1962).

Management Insecticides applied for suppression of artichoke plume moth, Platyptilia carduidactyla (Riley), normally keep aphids at low levels. For more information on aphid management, see the sections on melon aphid, Aphis gossypii Glover, or green peach aphid, Myzus persicae (Sulzer).

Asparagus Aphid

Brachycorynella asparagi (Mordvilko) (Hemiptera: Aphididae)

Natural History Distribution. This species was first observed in North America in 1969, in New York. Presumably, it had been introduced accidentally from Eurasia, where it is found widely in eastern countries and along the Mediterranean. It dispersed (or was dispersed) quickly, attaining North Carolina in 1973, British Columbia in 1975, Missouri and Washington in 1979, Alabama, Georgia, Oklahoma and Idaho in 1981, and California in 1984. Although it is abundant across the northern portions of North America, and very abundant along the west coast including southern California, it is uncommon in the humid southeastern states. Host Plants. This aphid feeds only on species of Asparagus. In addition to garden asparagus, Asparagus officinalis, it is known to feed on ornamental Asparagus spp. Natural Enemies. Many native predators, parasitoids, and insect diseases affect asparagus aphid. In New Jersey and Delaware, Angalet, and Stevens (1977) documented 31 species of natural enemies—the most abundant were ladybird beetles (Coleoptera: Coccinellidae), green lacewings (Neuroptera: Chrysopidae), and a parasitoid of numerous aphid species, Diaeretiella rapae (M’Intosh) (Hymenoptera: Braconidae). Other species of some importance were brown lacewings (Neuroptera: Hemerobiidae), the predatory midge Aphidoletes aphidimyza Rondani (Diptera: Cecidomyiidae), flower flies (Diptera: Syrphidae), other wasps (Hymenoptera: Braconidae and Aphelinidae), and a fungus. From these studies, the authors concluded that the natural enemies kept the population in check, which is the case in Europe (Stary, 1990) and Michigan (Ingrao et  al., 2017) as well. In fact, this aphid has not proved to be a serious pest of asparagus in eastern states and provinces (Hayakawa et  al., 1990). Once attaining the arid western regions of North America, however, asparagus aphid developed into a severe pest and seemed less influenced by natural enemies, though important biological control agents such as D. rapae seem to be exerting greater effects with time (Pike et  al., 1999). This suggests that climate plays a significant role, perhaps in conjunction with weather-­ sensitive natural enemies such as fungi. The edges of asparagus fields support greater numbers of aphid natural enemies than are found within asparagus fields, s­ uggesting that plant

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d­iversity, ­especially the presence of weeds or flowering plants, also may influence aphid abundance (Ingrao et al., 2017). Beginning in the mid-1980s, parasitoids such as Binodoxys (Trioxys) brevicornis (Haliday) (Hymenoptera: Braconidae) were imported from southern Europe and introduced into California, where some establishment resulted (Daane et al., 1995; Zumoffen et al., 2013). Life Cycle and Description. Oviparous aphids deposit overwintering eggs on the asparagus ferns in September or later. Beginning in about March, eggs hatch into aphids that develop into stem mothers (fundatrices). Fundatrices move to asparagus spears and give birth to about 18 nymphs. Subsequent generations may be apterous (wingless) or alate (winged). Sexual forms are produced in the autumn, they mate, and the females deposit eggs on the asparagus plant. The duration of a complete generation is about 15–19 days at 25°C. Egg. The eggs initially are green, but turn shiny black within 1–2 days. Females produce, on average, 10.5 overwintering eggs during their life span. The elliptical eggs are deposited in the lower one-third of the asparagus canopy. Nymph. The grayish-green nymphs exhibit four instars, the durations of which are about 2 days each, regardless of the morph or sex. Thus, nymphal development time averages about 8.0–9.5  days. Each female produces about 55 nymphs at 23°C, but only 27 and 9 at 14°C and 32.5°C, respectively. Adult. Both the alate and apterous adult viviparous aphids are present throughout the summer months and reproduce parthenogenetically. The adults are relatively small, measuring 1.2–1.7 mm long, and have short antennae. They are elongate oval, and green or gray green, often covered with a whitish waxy secretion. They blend in well with asparagus foliage but impart a slightly bluish-gray tint when the infestation is heavy. The most important characters to distinguish viviparous asparagus aphid from other a­ sparagus-infesting species are the inconspicuous cornicles and long cauda of B. asparagi (Stoetzel, 1990). Egg-producing females survive up to about 20 days. The biology of asparagus aphid was discussed by Tamaki et al. (1983b), Wright and Cone (1988a,b), and Hayakawa et  al. (1990). Keys to distinguish B. asparagi from other asparagus-infesting aphids were published by Stoetzel ­ (1990), and Blackman and Eastop (1984); in the latter, it was included in the key to asparagus-infesting aphids.

Damage Aphids feed on the new growth, causing shortening of the internodes, rosetting, dwarfing, and reduced root growth (Capinera, 1974a). Asparagus aphids deplete the sugars, particularly in the roots, and to a greater extent some other

aphids (Leszczynski et  al., 1986). Heavily infested plants have a bushy or bonzai-like appearance. Aphid infestation can kill seedlings or small plants in a relatively short time. Older, well-established plantings may show damage and even death in the year following infestation by aphids, especially in cold winter. Freezing and aphid infestation are synergistic; together they decrease survival and vigor of dormant asparagus crowns greater than either aphid feeding or freezing alone (Valenzuela and Bienz, 1989). The threat of damage is much greater in western areas than in eastern North America.

Management Sampling. Egg hatching can be predicted from temperature models. Eggs can be separated from foliage by washing with the petroleum-cleaning solvent (Wright and Cone, 1988a). Nymphs can be extracted from plants by heat or methyl isobutyl ketone (Wright and Cone, 1983a). Aphid distribution tends to be clumped, with most aphids in basal regions of the plants. Wright and Cone (1986) studied the distribution of aphids and recommended a sample of about 140 branches per field for making management decisions. Insecticides. Foliar insecticides can suppress aphids, but multiple applications may be necessary, especially under high-density conditions. Granular systemic insecticides may provide long-term control (Vernon and Houtman, 1983; Wildman and Cone, 1988). Some research has also been done to demonstrate the possibility of delivering insecticides to asparagus through the irrigation system (Wildman and Cone, 1986). Cultural Practices. Several cultural practices for suppression of asparagus aphid were investigated in Washington (Halfhill et  al., 1984; Folwell et  al., 1990). Autumn and spring tillage reduce aphid overwintering but also reduce subsequent spear production. Mowing and herbiciding within asparagus fields can destroy early-season aphids and delay the buildup of damaging populations. However, the destruction of wild (volunteer) asparagus is the most important cultural practice available, because it eliminates overwintering sites and limits invasion of aphids into commercial, aphid-free fields. Herbiciding, burning, and removal of asparagus crowns by digging are all viable options to eliminate volunteer asparagus.

Bean Aphid

Aphis fabae Scopoli (Hemiptera: Aphididae)

Natural History Distribution. The bean aphid apparently is native to Europe, but it has been spread to most temperate areas of the world except for Australia and New Zealand. Although  it

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occurs in tropical areas of Africa, it is not a very serious pest in warm environments. Bean aphid occurs throughout the United States and southernmost Canada, and the cooler areas of South America. Host Plants. The bean aphid is probably a complex of closely related species or subspecies. Typically, they are distinguished by their choice of secondary (summer) hosts rather than their morphology. Not surprisingly, then, this aphid feeds on a wide range of hosts, though it seems to favor plants in the family Chenopodiaceae as summer hosts. Vegetables attacked include asparagus, beet, carrot, celery, corn, fava bean, leek, lettuce, lima bean, onion, parsnip, pea, potato, spinach, pea, rhubarb, squash, and tomato. It also attacks sugarbeet, and in Europe, it is considered to be a very serious pest because it transfers viruses to this crop. Flowers such as nasturtium and dahlia commonly support this insect, as do many weeds, including curly dock, Rumex crispus; lambsquarters, Chenopodium album; and shepherd’s purse, Capsella bursa-pastoris. The taxonomy of this insect is confused, and some host records may prove to be due to other closely related species. The winter, or primary, hosts of bean aphid are Euonymus spp. and Viburnum spp. In England, abundance of bean aphid is positively correlated with abundance of spindle tree, Euonymus europaeus (Way and Cammell, 1982). Similarly, in Colorado, bean aphid primarily affects crops grown adjacent to urban areas, where Euonymous and Viburnum spp. are cultivated as ornamental shrubs. Natural Enemies. Fungi are an important mortality factor, often suddenly sweeping through high-density aphid populations. Several species of fungi are involved, but Neozygites fresenii apparently is the most effective in suppressing bean aphid populations (Dedryver, 1978; Wilding and Perry, 1980; Rabasse et al., 1982). Parasitism by native parasitoids, especially by Lysiphlebus testaceipes Cresson (Hymenoptera: Braconidae), is usually apparent when aphids are abundant. General predators such as green lacewings (Neuroptera: Chrysopidae), ladybird beetles (Coleoptera: Coccinellidae), and flower fly larvae (Diptera: Syrphidae) often are found feeding in bean aphid colonies. Ants commonly attend bean aphid, harvest honeydew, and apparently interfere with predators and enhance aphid survival (Capinera and Roltsch, 1981). In studies in Germany, ant attendance was highest in plant’s high sugar concentrations, which is secreted in aphid honeydew. Sugar richness and occurrence of the sugar melezitose are critical factors in determining the degree of ant attendance (Fischer et al., 2005). Natural enemies are believed to play an important role in the population cycles of bean aphid. When aphids are numerous on early-season hosts they provide abundant food for natural enemies, which then reach high levels of abundance in late-season populations, and greatly reduce

the number of aphids that overwinter. Therefore, the aphid population in the following year is quite low. However, low early-season populations fail to support natural enemies, so the aphids are free to reproduce and attain high late-season and overwintering population densities. This, of course, means that the early-season population will again be high. Because weather interacts with the aphids and their natural enemies, the cycling is not very predictable, so populationmonitoring systems have been developed in England to aid in the prediction of aphid outbreaks. Life Cycle and Description. The eggs hatch in the spring, often as early as February, and produce 1–2 generations of apterous (wingless) parthenogenetic females. This is followed by a generation of alate (winged) females that fly from the primary host to secondary (summer) host plants, where females reproduce parthenogenetically all summer. Alate and apterous forms are produced according to density and host-plant conditions, with high densities and depleted plants tending to produce alatae. In the autumn, in response to short-day conditions, alatae is produced that colonize the primary hosts. On the overwintering host, the females mate and lay eggs among cracks in the bark and on bud axils. Egg. In temperate areas, bean aphid overwinters as an egg on one of its primary hosts. Initially green, the eggs soon turn shiny black. In warmer areas, the aphids reproduce continuously without producing eggs. Nymph. Nymphs are dark, but bear four pairs of transverse white bars on the dorsal surface of the abdomen. Most authors report four instars. Tsitsipis and Mittler (1976), for example, indicated durations of 2, 2, 1.5, and 2.5 days for instars 1–4, respectively when reared at about 20°C. OgengaLatigo and Khaemba (1985), however, reported only three instars, with durations of about 2.3, 3.0, and 2.5 days. Total nymphal development time requires 5–10 days at 28–17°C, respectively. Adult. Bean aphid is dark olive green to dull black in color. The body length is 1.8–2.4 mm in females, with males only slightly smaller. The appendages tend to be black, but the tibiae may be pale, in part. The wings are transparent. On some crops bean aphid may be confused with another black species, cowpea aphid, Aphis craccivora Koch; adults of this latter species are shiny black with whitish legs, and on an average, they are smaller in size. Reproduction commences soon after attainment of the adult stage, usually a period of about 3–6  days. Both alate and apterous females reproduce. The adult produces about 85–90 nymphs during her reproductive period, which is estimated at 20–25 days. Most offspring are produced in the first 5–10 days of the reproductive period. Reproduction i­ ncreases with temperatures, up to a threshold of about 24°C, and

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then decreases. The reproductive period is followed by a postreproductive period of about 7  days (Frazer, 1972). Apterous females give birth to more, and larger, nymphs than alatae (Dixon and Wratten, 1971). The winged aphids disperse freely, but their eventual disposition depends largely on wind and windbreaks because they do not have strong powers of flight. Thus, leeward sides of hills and windbreaks are the areas where aphids accumulate. They also are deposited more heavily on the edges of crops. Because small fields have proportionally more “edge,” aphid mean density also tends to be higher in small plantings. Sometimes, dispersal of up to 30 km occurs. An excellent summary of bean aphid ecology was given by Cammell (1981). Tsitsipis and Mittler (1976) provided information on rearing aphids on both plants and artificial media. Keys for identification of bean aphid are found in several host-based keys (e.g., bean, carrot, celery, cucumber, lettuce, pea, potato, squash-infesting aphids) in Blackman and Eastop (1984), and also in Palmer (1952). Stoetzel et al. (1996) published a key for cotton aphids that is also useful to distinguish bean aphid from most other common ­vegetable-infesting aphids.

Damage Aphids tend to reproduce rapidly and build to high numbers on plants that are actively growing. They concentrate their feeding on young tissue and may deplete nutrients needed for plant growth. In sugarbeet, for example, bean aphid infestation decreased foliage and root weights, and sucrose yield (Capinera, 1981). Symptoms of infestation include curling of leaves and stunted plants. Studies on bean suggest that aphid feeding during the preflowering stage is more damaging than later in development (Khaemba and Ogenga-Latigo, 1985). Research from England indicates that aphid suppression is warranted when 5% of faba bean plants are infested (Bardner et al., 1978). In leafy vegetables such as celery, however, contamination of foliage with aphid bodies or honeydew is also a very important factor, as these conditions will cause crops to be rejected (Godfrey and Chaney, 1995). Estimates of honeydew production indicate that aphids may produce up to 30–40 drops per day (Banks and Macaulay, 1964). Significant damage is, however, infrequent in North America. Bean aphid is capable of transmitting numerous plant viruses, including many common viruses of vegetable crops. Kennedy et al. (1962) listed over 60 diseases, mostly stylet-borne viruses, transmitted by bean aphid. Although not usually considered as important a vector as many other species (Fereres et al., 1993), some work indicated that this insect was quite effective at transmitting beet yellows virus (Kirk et al., 1991).

Management Sampling. This species often develops very high densities, perhaps 5000 aphids per plant. Because of the difficulties in dealing with such abundance, procedures for estimating aphid density tend to avoid counting actual numbers per plant and focus instead on counts of subsamples, determination of the number of leaves infested (Banks, 1954), or length of infestation (Capinera, 1981). In England, forecasts of aphid populations are made based on densities of overwintering eggs on the spindle tree. Considerable savings in insecticide use are possible in years when aphid densities are low (Way et  al., 1977; Gould and Graham, 1977). Suction traps may also be used in forecasting, particularly in determining the time of migration (Way et al., 1981). There does not seem to be a need for such a forecasting system in North America owing to the relatively minor pest status of this insect. To monitor invasion of celery, and most likely other vegetable crops, yellow water pan traps and yellow sticky traps can be used (Godfrey and Chaney, 1995) though sticky traps should be avoided because aphids are often damaged by the adhesive, making identification difficult. Insecticides. Insecticides are applied as foliar sprays or, for systemic insecticides, also as granules at planting time. However, the use of granular materials in short-season crops such as beans is sometimes impossible, because the insecticide residue is too high at harvest time. In some crops, the disruption of pollinators is a concern. Bean aphid possesses both an alarm pheromone and a sex pheromone (Dawson et  al., 1990). The alarm pheromone causes the aphid to become disturbed and may induce it to leave the host plant. This may prove useful in inhibiting virus transmission, increasing contact with insecticide residues, or by increasing aphid exposure to predators or unsuitable environmental conditions. The sex pheromone is active only at short distances, and its use is yet to be developed. Biological Control. Research in England has demonstrated that the introduction of entomopathogenic fungi could suppress aphid abundance on beans and increase yield (Wilding, 1981). However, Erynia neoaphidis and Neozygites fresenii fungi are effective only at high aphid densities and under cool, moist weather conditions. Cultural Practices. Faba beans have been examined for their resistance to bean aphid, and some cultivars differ in susceptibility. However, this species is more susceptible than more primitive Vicia species (Holt and Birch, 1984). The partial resistance of faba beans, when combined with intercropping with a nonhost plant (wheat and barley), can reduce the abundance of bean aphids (Hansen et al., 2008). Border plantings of scorpion weed, Phacelia tanacetifolia (Hydrophyllaceae), have been shown in England

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to enhance the predation of bean aphid, and other aphid ­species, by flower flies (Diptera: Syrphidae). Adult flower flies require nectar for energy, and pollen for sexual maturation and egg production, with this flowering annual, easily manipulated to provide these resources (Hickman and Wratten, 1996).

Bean Root Aphid

Smynthurodes betae Westwood (Hemiptera: Aphididae)

Natural History Distribution. This aphid is found throughout the world, though it is not often recognized as a pest. Its origin seems to be the Mediterranean region. Host Plants. The primary host of bean root aphid is pistachio and related trees in the genus Pistacia. Its secondary hosts are numerous dicotyledonous plants, particularly in the families Compositae, Leguminosae, and Solanaceae, but also including Chenopodiaceae, Cruciferae, Polygonaceae, and others. Monocots (grasses) rarely serve as hosts. The principal vegetable crops affected are bean, cabbage, okra, pea, parsnip, pepper, squash, potato, and tomato. It also occurs on cotton and pistachio. Natural Enemies. The natural enemies of this insect are poorly known, but gall-forming insects such as this aphid typically are not greatly affected by natural enemies because they are somewhat protected by the gall in which they feed. However, Monoctonia pistaciaecola Stary (Hymenoptera: Braconidae) was observed to affect an appreciable proportion (20%) of the galls in Israel (Wool and Burstein, 1991). Life Cycle and Description. Bean root aphid has a complex, 2-year life cycle when Pistacia is involved, though it can also reproduce parthenogenetically on secondary hosts, bypassing the sexual stage. During the first year, nymphs developing from overwintered eggs initially form small red galls on the midvein of the Pistacia leaflet. At maturity, each of these aphids produces about 20 nymphs parthenogenetically which disperse to the margins of the leaflets and form new galls. Within the second set of galls, two additional generations are produced, resulting in about 35 aphids in each gall by autumn, when winged forms emerge from the galls and disperse to secondary hosts. Offspring of the dispersants penetrate the soil and feed on the roots. During the following spring winged forms are produced on the secondary plants which disperse back to the primary host and parthenogenetically produce male and female forms. The sexual forms mate and produce eggs. Egg. Small black eggs are deposited in crevices on the trunk and major branches of Pistacia.

FIG. 8.7  Adult bean root aphids. (Photo by L. Depa, Poland.)

Adult. Wingless (apterous) adults are quite spherical in general body form. The antennae, head, legs, and abdomen are brownish or pinkish and are covered with a moderately dense covering of short hairs. Cornicles are not present. This small aphid measures about 1.5–2.7 mm long. The antennae, head, and appendages of the winged (alate) form are dusky to black. The abdomen is green or olive, with a broad transverse black band across the dorsal surface of each segment. The short hairs also cover the body of this form and are white. Alatae is slightly larger, 2.2–2.7 mm long, and bears wings with dusky veins. Bean root aphid was described by Cottier (1953). A summary of its life cycle was provided by Burstein and Wool (1993). Keys for identification of bean root aphid are found in several host-based keys (e.g., bean, cabbage, okra, pea, potato, squash, tomato-infesting aphids) in Blackman and Eastop (1984),

Damage This species is rarely considered a pest, but when it is numerous it causes wilting and stunting of plants, and may cause deformities of roots and tubers.

Management Management is rarely a consideration, but if it becomes necessary the approaches used for management of other root aphids such as lettuce root aphid, Pemphigus bursarius (Linnaeus) and the sugarbeet root aphids, Pemphigus betae Doane and P. populivenae Fitch, likely are applicable.

Bird Cherry-Oat Aphid

Rhopalosiphum padi (Linnaeus) (Hemiptera: Aphididae)

Natural History Distribution. This species probably originated in North America but is now found in Europe, Asia, and

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New Zealand as well as throughout the United States and southern Canada. Host Plants. Bird cherry-oat aphid may alternate between winter and summer hosts, or in mild-winter climates, it can persist throughout the year on summer hosts. The winter or primary, the host is Prunus spp. The summer or secondary, hosts are numerous species of grasses, including all the major cereals and pasture grasses. Sedges, iris, and a few other plants are occasionally reported as hosts, but these records are suspect. It is found principally on wheat, barley, oat, and rye, but in some locales, it is a major pest of corn, including sweet corn. Susceptibility of hosts to infestation and suitability for aphid reproduction have been assessed by many investigators, including Villanueva and Strong (1964), Dean (1973), Leather and Dixon (1982), and Weibull (1993). Among the best hosts are such wild or forage grasses as wheatgrass, Agropyron spp.; brome, Bromus spp.; ryegrass, Lolium and Lilium spp.; and wild oat, Avena fatua. Among crop plants, barley seems to be optimal in most areas, but sweet corn is very suitable in the Pacific Northwest. Natural Enemies. Bird cherry-oat aphid is susceptible to attack by many of the predators, parasitoids, and pathogens affecting other aphids. Lady beetles (Coleoptera: Coccinellidae), green lacewings (Neuroptera: Chrysopidae), brown lacewings (Neuroptera: Hemerobiidae), and flower flies (Diptera: Syrphidae) are the most common predators. In Europe, and presumably in North America, eggs suffer high mortality rates during the winter months owing to cold weather in most northern locations and predators in most mild locations. In small grain crops grown in Idaho, Aphelinus varipes (Forester) (Hymenoptera: Aphelinidae); Aphidius ervi Haliday, Diaeretiella rapae (M’Intosh), Lysiphlebus testaceipes (Cresson) and Praon sp. (all Hymenoptera: Braconidae) were recovered from bird cherry-oat aphid, with A. varipes most abundant (Feng et al., 1992b). Fungi have some significant effects on aphid populations, but this occurs mostly in irrigated fields. Precipitation, particularly heavy rainfall, also has direct negative effects on aphid survival during the summer months. Bird cherry-oat aphid can transmit viruses that cause plant disease, and the effectiveness of natural enemies in preventing transmission of plant disease is often debated. However, Smyrnioudis et al. (2001) addressed this in a laboratory study using barley yellow dwarf virus and wheat seedlings. The presence of the predatory ladybird beetle Coccinella septempunctata (L.) (Coleoptera: Coccinellidae) seemed to increase the incidence of barley yellow dwarf virus, whereas the presence of the wasp parasitoid Aphidius rhopalosiphi de Stefani Perez (Hymenoptera: Braconidae) seemed to decrease the incidence of virus, relative to the control. In time, however, nearly all plants were infected.

Life Cycle and Description. The biology of this aphid varies, depending on the availability of primary hosts and weather conditions. A generation can be completed in about 5  days when reared at 26°C, but it requires about 22 days at 13°C. The longevity of the individual aphid is often 18–20  days. Aphids developing on primary hosts undergo 2–3 generations before overcrowding induces the formation of winged (alate) forms which disperse to grasses. Aphids may reproduce continuously during the summer months, but reproduction is disrupted if the temperature exceeds 30°C for even a few hours daily. Aphids feeding on wild grasses and grains may migrate back to Prunus and produce eggs in the autumn, or they may remain on Graminae through the winter months. In Manitoba, winged aphids are found dispersing in late May and June, and then again in October (Robinson and Hsu, 1963). In the northern Great Plains region of the United States, however, little overwintering occurs, this area instead becomes reinvaded annually by winged migrants from the south. Some individuals crawl down the plant stems and survive below-ground during cold weather (Kieckhefer and Gustin, 1967), and though this does not appear an important method of overwintering under the cold winter climate conditions of Idaho (Feng et  al., 1992a,b), it likely is effective in warmer climates. Egg. The egg is green initially, but turns black with age, and is deposited on the bark of Prunus trees. The egg measures about 0.57 mm long and 0.26 mm wide. In England, there is a steady but low rate of attrition in eggs during the winter months primarily due to predation, followed by a fairly sharp increase in egg loss in the spring, resulting in loss of 70%–80% (Leather, 1981). Nymph. There are four instars. The mean body length of nymphs is 0.6, 0.9, 1.2, and 1.4 mm for instars 1–4, respectively. The development time of nymphs reared at 26°C, which is about optimal for development, is 1.4, 1.1, 1.1, and 1.4  days for instars 1–4, respectively. Nymphs are green, but there is pronounced orange or rust color posteriorly near the base of the cornicles. First instar nymphs have only four antennal segments, instars two and three have five segments, and last instar and adults bear six segments. Aphids reared under cool conditions are greenish black and those reared under warmer conditions are light green. Aphids feeding on Prunus live on bud and young leaf tissue; the latter may become deformed and form a pseudo-gall partly enclosing and protecting the aphid colony. Older leaf tissue also is attacked, and eventually, some of the adults are winged. The shift to winged, dispersing forms coincides with a decrease in the nutritional quality of Prunus foliage (Dixon, 1971; Dixon and Glen, 1971).

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is ­included in host-based keys in Blackman and Eastop (1984) and Stoetzel and Miller (2001). The biology of this insect in Europe was reviewed by Leather et  al. (1989). Developmental biology was given by Villanueva and Strong (1964), Elliott and Kieckhefer (1989), and Michels and Behle (1989).

Damage

FIG. 8.8  Bird cherry-oat aphid, wingless. (Drawing by USDA.)

This species is known principally for its damage to small grains. It builds to high densities and damages grain plants directly, and it is also an important vector of several plant viruses, most notably barley yellow dwarf. Both persistent and nonpersistent viruses are vectored (Kennedy et  al., 1962). However, in some areas, it is the most abundant aphid on corn. Corn also serves as an important link or bridge for aphids and barley yellow dwarf virus between spring and autumn small grain crops (Blackmer and Bishop, 1991).

Management

FIG. 8.9  Bird cherry oat-aphid, winged. (Drawing by USDA.)

Adult. The body of the wingless adults (apterae) is greenish or olive brown, with the head and prothorax yellowish brown. The legs are green or darker, the cornicles green but with the tips dusky. A rusty colored patch is found around the base of the cornicles. The body length is 1–2 mm, averaging 1.7 mm. In the winged forms (alatae), the abdomen usually is light green but darker medially, though sometimes it is entirely dark green. A rusty brown patch may be found around the base of the cornicles. The head and thorax are black, but the prothorax is green. The appendages are dusky to blackish. The body length is 1.8–2.0 mm. The prereproductive period of young adults is normally less than a day. Viviparous (parthenogenetic) adults produce 10–50 nymphs during their life span, with the host plant, plant age, and temperature affecting reproduction. However, normal fecundity is 40–50 at optimal temperature during a reproductive period of about 15  days. Young plants and cool weather enhance reproduction. This species is easily confused with several grain-­ infesting species, but fortunately is not too readily confused with the other common corn-infesting aphid, corn leaf aphid, Rhopalosiphum maidis (Fitch). The body of bird cherry-oat aphid is ovate, whereas the body of corn leaf aphid is elongate. Also, the rust-orange color about the base of the spiracles distinguishes bird cherry-oat aphid from the solid blue-green color of corn leaf aphid. This species was included in the aphid keys of Palmer (1952) and Olsen et  al. (1993). Bird cherry-oat aphid

This aphid is principally a pest of small grains, and sampling protocols have been developed for such crops, but not for corn. For example, Ekbom (1987) and Elliott et al. (1990) presented binomial or presence-absence protocols for R. padi sampling on small grains. Nevertheless, aphids may attain high numbers on sweet corn and stimulate the application of foliar insecticides. There is evidence of resistance to certain insecticides in Europe and China. Cultural practices may affect the development and abundance of R. padi, and ultimately their damage potential. Aqueel and Leather (2011) studied the effects of nitrogen fertilization on aphids feeding on fertilized wheat plants. Fecundity and longevity increased with fertilizer application, and nitrogen fertilizer reduced the time to attain maturity.

Buckthorn Aphid

Aphis nasturtii Kaltenbach (Hemiptera: Aphididae)

Natural History Distribution. Buckthorn aphid is widespread in North America, but as a pest, it is limited principally to eastern Canada and the northeastern United States. This species occurs throughout Asia and Europe. Its origin is uncertain but could be Europe. Host Plants. This insect feeds on numerous plants from many plant families. It is principally a pest of potato during the summer months. In Maine and New Brunswick it is often the most abundant aphid in potato fields. Other vegetables that may serve as summer hosts include bean, beet, cabbage, cucumber, parsnip, pepper, salsify, squash, and watercress. Interestingly, in Maine, this little-known

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aphid has been reported to be the most abundant aphid on cucumber and squash. Other crops that support buckthorn aphid include buckwheat, strawberry, alsike and red clover, and tobacco. It also affects flowers such as dahlia, hollyhock, nasturtium, pansy, sunflower, and zinnia. Among the weeds supporting growth of these aphids are Canada thistle, Cirsium arvense; common plantain, Plantago major; dock, Rumex spp.; lambsquarters, Chenopodium album; pigweed, Amaranthus retroflexus; purslane, Portulaca oleracea; shepherd’s purse, Capsella bursa-pastoris; sorrel, Oxalis spp.; smartweed, Polygonum spp.; and wild radish, Raphanus raphanistrum. The winter hosts are common buckthorn, Rhamnus cathartica, and alder-leaved buckthorn, R. alnifolia; both are shrubs, and the former is occasionally planted as a hedge, but the latter is more important in overwintering. Natural Enemies. Buckthorn aphid has many natural enemies, though it is less heavily parasitized than potato aphid, Macrosiphum euphorbiae (Thomas). Among the parasitoids reared from buckthorn aphid are Praon spp., Aphidius spp., Diaeretiella rapae M’Intosh and Ephedrus incompletus (Provancher) (all Hymenoptera: Braconidae). The most important parasitoid, however, is Aphidius nigripes Ashmead (Hymenoptera: Braconidae) (Shands et al., 1965). Most of the common lady beetles (Coleoptera: Coccinellidae), some lacewings (Neuroptera: Chrysopidae), flower flies (Diptera: Syrphidae), and the predatory midge Aphidoletes aphidimyza (Rondani) are reported to prey on buckthorn aphid. Ground beetles (Coleoptera: Carabidae) affect aphid populations, but not as greatly as canopy-level predators (Boiteau, 1986). Several fungi affect buckthorn aphid, sometimes causing population collapse (Shands et  al., 1965). Alkyokhin et  al. (2005) concluded, after reviewing over 50  years of data, that density-dependent factors were affecting buckthorn aphid populations, resulting inperiodicity of population cycles of about 6  years. However, over time the cycling and overall density of the aphids diminished, which they attributed to the introduction of alien lady beetles or new insecticides. Subsequently, these same authors (Alkyokhin et  al., 2011) deduced that buckthorn aphid was affected negatively by predators, but not by parasitoids. The weather had only modest effects on aphid abundance. Life Cycle and Description. In Maine, eggs overwinter on buckthorn and hatch in April. One or more generations consisting of apterous (wingless) parthenogenetic females are produced on the buckthorn leaves, but beginning with the second generation some alate (winged) females are also produced. In subsequent generations, larger and larger proportions of alatae are produced, with the winged individuals dispersing to summer hosts. The aphids developing on summer hosts are mostly apterous until August when winged males and females are produced. The winged sexual forms disperse to buckthorn where the females deposit nymphs that, upon maturing, mate with the males and deposit eggs.

Egg. The deposition of eggs on buckthorn commences in early September in Maine. They usually are deposited on the buds of alder-leaved buckthorn. The small oval eggs are green when first deposited, but soon turn black. They measure about 0.5 mm long and 0.25 mm wide. In a 24-year study conducted in Maine, egg density averaged 102 eggs per 100 buds (range 5–300 per 100 buds) in the autumn, and 72 eggs per 100 buds (range 2.5–300 per 100 buds) in the spring. Overwintering mortality of eggs ranged from about 30%–90%. Nymph. Five instars were reported by MacGillivray and Anderson (1958). They reported mean development times of 1.9, 1.9, 1.4, 1.7, and 0.3 days, respectively, when reared at 20°C. Wang et  al. (1997b) reported only four instars, with mean development times of 2.4, 2.9, 2.7, and 2.0 days, respectively, when reared at 20°C, and 1.4, 1.4, 1.2, and 1.2  days when reared at 30°C. The lower developmental threshold was reported to be 6.5°C, and the upper threshold at 35.6°C. The optimal range for development is 20–30°C. Adult. The apterous (wingless) parthenogenetic female of buckthorn aphid is small and measures 1.1–2.1 mm long. The body is flattened and yellow or green in color. The antennae are shorter than the length of the body, and the head lacks tubercles near the base of the antennae. The cornicles are short. The winged (alate) parthenogenetic form is similar but tends to have a dark head, thorax, and abdominal tip. The body length is 1.5–2.4 mm. The sexual forms that are produced in autumn display sexual differences in coloration—the females are yellow but the males dark. Each adult female feeding on potato produces, on average, about 64 nymphs during a reproductive period averaging 20 days. The biology of buckthorn aphid is not fully described, but good treatment was provided by Patch (1924) and Shands and Simpson (1971). MacGillivray and Anderson (1958) and Wang et  al. (1997a,b) gave developmental information. Buckthorn aphid was included in the keys by Palmer (1952) and the pepper and potato keys provided by Blackman and Eastop (1984).

Damage This species can be quite numerous and damaging to potato and cucurbits in eastern Canada and the northeastern United States. It also causes the deformity of buckthorn hedge foliage. It occurs on the lower surface of leaves, favoring the lower leaves of plants that are grown in full sunlight, but it disperses freely to terminal areas of plants growing in the shade. In this respect, it differs considerably from potato aphid, which favors terminal growth as feeding sites. At high densities, it causes stunting of potato and even death. Buckthorn aphid also may transmit plant viruses, including potato leaf roll, beet yellows, and cucumber mosaic virus (Kennedy et al., 1962).

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Management Sampling. The egg abundance on buckthorn was shown to be positively related to subsequent aphid abundance in potato, though the correlation is not strong (Shands and Simpson, 1971). Aphid populations normally are monitored by visual examination of crops or with yellow water pan traps. Insecticides. Foliar insecticides are commonly applied for aphid suppression. Broad-spectrum insecticides are sometimes used because of the other pests associated with solanaceous crops. Chemical suppression is not usually recommended unless half of the leaves are infested. Soaps and dishwashing detergents are also effective against aphids, but care should be taken not to burn the plants. It is important to apply the treatments carefully so that the lower leaf surfaces receive good coverage. Treatment of buckthorn hedges in the spring can destroy early-season aphid populations and reduce infestation potential in crop plants. Cultural Practices. Studies conducted in Maine demonstrated that planting practices could influence damage to potato. Delay of planting from early May to late May or early June resulted in up to a 90% reduction in aphid infestation. Early hilling operations, wherein row ridges are heightened with soil from the row middles, similarly deprived dispersing aphids of young plant tissue, resulting in lower aphid densities (Shands et  al., 1972). Delayed planting was also shown to be beneficial in New Brunswick (Boiteau, 1984). Reflective mulches, particularly aluminum mulch, are sometimes recommended for disruption of aphid invasion of crops. Evaluation of aluminum mulch in potato, however, showed that the beneficial effect of the reflective mulch was slight or of short duration, and therefore impractical (Shands and Simpson, 1972). Destruction of buckthorn, or treatment with insecticide, can decrease the risk of crop infestation.

Cabbage Aphid

Brevicoryne brassicae (Linnaeus) (Hemiptera: Aphididae)

Natural History Distribution. Cabbage aphid is found throughout the United States and Canada, and in most other countries with a temperate climate. This aphid was probably introduced accidentally to North America from Europe by early colonists, as records of its damage to crops extend back to the late 1700s. Host Plants. Virtually all crops in the family Cruciferae are suitable hosts for cabbage aphid, but feeding is restricted to this family. The vegetable crops most severely affected are broccoli, Brussels sprouts, cabbage, and cauliflower, but collards, kale, kolhrabi, mustard, radish, rape, swede, and turnip are occasionally attacked. Beirne (1972) reported that in ­western Canada the principal injury of cabbage aphid was

caused to early cabbage and cauliflower, whereas in eastern Canada turnip was the principal crop injured. In the United States, probably due to shifting dietary (broccolli has become more popular) cabbage aphid is an especially troublesome pest of broccoli. However, the principal form of damage is due to contamination of the “head” (which is the blossom) with live and dead aphids, as this affects the quality of the product. The infestation of the outer leaves is of little consequence, as this part of the plant is removed at harvest. Interestingly, Nieto et al. (2006) reported that when aphids arrive early into a broccoli field there is less likely to be a problem at harvest time than if they arrive later. They attribute this to the actions of natural enemies, and the early arrival of aphids affords greater time for the natural enemies [principally flower fly larvae (Diptera: Syrphidae) in California] to increase in abundance so they can regulate the aphid population. Seed heads of radish are suitable hosts, as are numerous cruciferous weeds, especially Brassica spp. and shepherd’s purse, Capsella bursa-pastoris. The mustard-oil glycoside, sinigrin, functions as a feeding stimulant for cabbage aphid, as it does for other insects that specialize in feeding on crucifers (Moon, 1967). Natural Enemies. Aphids normally are attacked by numerous predators and parasites, but cabbage aphid seems less susceptible to natural control than most species. Diaretiella rapae (Macintosh) (Hymenoptera: Braconidae) is invariably associated with this aphid and often builds up to high levels late in the season, when aphids are most numerous. However, its ability to attain densities sufficient to suppress cabbage aphid populations normally is hampered by the presence of several species of hyperparasites (Pimentel, 1961; Oatman and Platner, 1973). Supplemental releases of D. rapae to field plots can, however, greatly raise the level of parasitism (Zhang and Hassan, 2003). Flower fly larvae (Diptera: Syrphidae), principally Allograpta, Syrphus, and Scaeva spp., and predatory cecidomyiid Aphidoletes aphidimyza (Rondani) (Diptera: Cecidomyiidae) larvae are generally reported to be the most important natural control agents (Oatman and Platner, 1973; Raworth et  al., 1984; Nieto et  al., 2006). Lady beetles (Coleoptera: Coccinellidae), while sometimes found feeding on cabbage aphids, are unusually ineffective (George, 1957). Some authors have suggested that predators and parasitoids decreased cabbage aphid population size only after aphid population growth rate is reduced by host plant deterioration or other factors. Fungi sometimes affect cabbage aphids but causes epizootics only at very high aphid densities. Life Cycle and Description. Cabbage aphid may have numerous generations per year, depending on climate; 20 are reported from southern California. In the north, this species produces sexual forms and overwinters in the egg stage, whereas in the south, sexual forms and eggs are not observed. Trumble et  al. (1982b) reported that eggs were the overwintering stage in northern California, but that

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crop debris and weeds served as the overwintering site for females in the southern portion of the state. Unlike many aphids, cabbage aphid does not disperse to an alternate host to overwinter. Egg. Egg deposition occurs just 1 day after mating, and the oviparous (egg-laying) aphids may live for a month or more. The egg production is not well studied, but Herrick and Hungate (1911) estimated 5–7 eggs per female. The eggs are initially pale yellow or yellow green, but they become shiny black within a few days of deposition. They are generally found on the underside of leaves and measure 0.65 mm long and 0.15 mm wide. Unlike many aphids, cabbage aphids do not alternate between summer and winter hosts; the eggs are found on the plants fed upon by the summer populations. Eggs typically hatch in April. Nymph. There are four instars, each about 2–3 days in duration. Nymphal bodies initially lack the grayish white, waxy exudate that is so typical of this species but acquires this soon after they commence feeding. The waxy material is easily removed, revealing a grayish-green insect with two rows of black bars along the back.

FIG. 8.10  Cabbage aphids of various ages. (Photo by L. Buss.)

FIG. 8.11  Adult winged cabbage aphid. (Photo by L. Buss.)

Adult. During the spring and summer months, the ensuing adults are all female and they reproduce parthenogenetically. Adult wingless (apterous) females give birth to 30–50 nymphs in their life span, which is about 30–40 days. Initially, their reproduction is high but drops off markedly as they mature. Most aphids found during the summer months are wingless females; only at high densities or when the host plant deteriorates are winged forms produced. The wingless females are grayish green, with a dark head, and pale-brown legs. The cornicles are dark and measure about 0.16 mm long. There is a double row of dark bars on the back, and the body is covered with a white powder or mealy secretion. The body measures 1.6–2.6 mm long, averaging about 2.5 mm. The winged (alate) parthenogenetic females are similarly dull green with a dark head, but the legs are dark brown. The dorsal surface of the abdomen is marked with a single row of dark bars, and the veins of the transparent wings are dusky to black. The cornicles are dusky green to black. The entire body of this insect is dusted with a fine white powder. The winged parthenogenetic female measures 1.6–2.8 mm long, averaging about 1.9 mm. The winged forms, though not strong fliers, quickly disperse over crop fields and to adjacent fields. The winged females have a shorter life span than the wingless form; the former persists only about 10  days. They also produce relatively few offspring, about 6–8. In the autumn months, especially September and October, egg-laying forms may be produced. Apparently, the oviparous forms can be produced both by winged and wingless summer forms. The oviparous female has a palegreen or greenish-yellow body, with a row of dark bars located centrally on the back. This form is wingless and measures about 2.2 mm long. Males are also produced at this time, but unlike the wingless oviparous females, the males have transparent wings. Males are greenish brown or yellowish, and have dark legs and a double row of dark bars on the back. The body measures about 1.3 mm long. In autumn, when sexual reproduction occurs, females produce a sex pheromone that is attractive to males, and which also is attractive to parasitoids (Gabryś et al., 1997). Cabbage aphid is easily confused with turnip aphid, Lipaphis pseudobrassicae (Kaltenback), though turnip aphid generally can be distinguished by the very sparse occurrence of waxy exudate on the body and longer cornicles. In humid climates, however, there tends to be a greater accumulation of waxy secretions on the turnip aphid’s body. Therefore, Blackman and Eastop (1984) recommended using the shape of the cauda—a structure found at the tip of the abdomen—to differentiate these two species. When viewed from above, the cauda of cabbage aphid is triangular, about as wide as it is long. In contrast, the cauda of turnip aphid is slender, about twice as long as it is wide.

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A comprehensive treatment of cabbage aphid was provided by Herrick and Hungate (1911). Brief description of these aphids and keys for their identification are found in Palmer (1952), and Blackman and Eastop (1984) included cabbage aphid in their cabbage, radish, mustard, and swede keys. Cottier (1953) also provided a description of cabbage aphid.

Damage Cabbage aphid populations, if not controlled, often build to very high densities. Heavily infested plants acquire a grayish appearance due to the mass of aphid bodies on the foliage. Honeydew and sooty mold are often evident. High densities also cause the leaves to wrinkle and curl, usually cupping downward. Under dry conditions, aphids cause the plants to wilt and leaf tissue that has been fed upon may turn yellow. Cabbage aphid prefers the youngest tissue and highest portions of the plant but may occur on both the upper and lower surface of foliage. Flower heads of seed crops may be attacked, reducing the setting of seed. Contamination of the plants with honeydew and aphids can cause considerable loss. In California, cabbage aphid is the most serious aphid pest of Brussels sprouts and broccoli because they are hard to remove from the head of the plant and thereby contaminate the produce (Trumble, 1982b; Pickel et al., 1983). Wolfenbarger (1967) estimated the impact of aphid densities on cabbage yield. He projected that when aphid densities averaged one per square inch (0.16 per cm2) on the most heavily infested leaf of each plant, the proportion of plants in a field that was aphid-free would be 64%. Further, he estimated that when there was an average aphid density as low as one per plant, the proportion of unmarketable heads would be 12%. In addition to the direct effects of feeding by aphids on plant growth and the damage caused by aphid contamination of foliage, cabbage aphid can also be a vector of plant viruses. Over 30 viruses are known to be transmitted by B. brassica. Cauliflower mosaic and cabbage ring spot virus transmission were studied by Broadbent (1954); cabbage aphid transmits cauliflower mosaic more effectively because this virus concentrates in the young tissue of the plant, which is the preferred feeding site of the aphid. Hostplant resistance offers considerable potential, but currently, it is difficult to put into practice. Both very hairy and hairless cultivars are likely to be less suitable for parasitoid activity than plants with intermediate densities of plant hairs.

Management Sampling. Aphid distribution within crop fields is highly aggregated when overall densities are low but become more uniform when aphid densities increase (Trumble, 1982a). Populations of winged aphids are more effectively sampled with cylindrical sticky traps than by water pan traps, but

such populations are not consistently correlated with onplant densities (Trumble et al., 1982a) and aphids removed from sticky traps are difficult to identify. Sequential sampling protocols were presented by Wilson et al. (1983a). Insecticides. Commercial producers usually depend on chemical insecticides to maintain an insect-free crop. Granular forms of systemic insecticides are sometimes used at planting and may provide aphid suppression for several weeks. However, systemic and nonsystemic insecticides are also applied to the foliage, and frequent applications may be necessary to maintain a clean crop. Many crucifer crops have curly leaves or form heads, and once aphids are established within such protected locations it becomes difficult to obtain control. Thus, most growers prefer to apply insecticides as a preventative measure, rather than waiting for the infestation to occur and then seeking to cure the problem. It is often difficult to attain good control of cabbage aphid during cool weather ( 26 whiteflies per leaf. Shipp and Wang (2006) found that although D. hesperus was effective at suppressing Frankliniella occidentalis Pergande (Thysanoptera: Thripidae), tomato fruit damage occurred when the ratio of predators to prey exceeded 1:10. Tomatoes compensate quite well for fruit abortion, leading to production of fewer but larger fruit. Maximum compensation occurred at about 15% fruit loss, and significant yield reduction occurred only when fruit loss exceeded 27% (Sánchez and Lacasa, 2008). Although tomato bugs unquestionably injure plants, the dependency by tomato bugs to feed on plants seems variable. Nesidiocoris tenuis can be reared quite well using only E. kuehniella eggs. Plants are considered, by some, to be only a source of water and an oviposition substrate, and not important in the nutrition of the bugs (De Puysseleyr et al., 2013). Consistent with this, N. tenuis populations decrease after prey populations diminish, suggesting that plants alone are not adequately nutritional (Sanchez, 2008). However, D. hesperus did not survive well on a diet of E. kuhniella eggs unless they also had access to either tomato foliage or supplementary water (Gillespie and McGregor, 2000). Also, D. hesperus bugs lived longer and produced more eggs when either fruit or leaf tissue was complemented with prey (eggs of E. kuhniella) relative to plant tissue alone (Gillespie et al., 2012).

Management Sampling. Sampling is usually conducted by visual examination of plants. These bugs tend to be concentrated in the apical regions to the middle areas of the plants and not found in great abundance in the lower areas. Like many insects, their distribution among plants tends to be ­aggregated.

Due to their small size and cryptic coloration they are difficult to census. Sanchez et  al. (2002) recommend use of a presence-absence sampling plan rather than counting D. hesperus. This is feasible because there is a strong positive relationship between population density and the proportion of occupied plant units. Insecticides. Tomato bugs can be controlled with insecticides when warranted. Insecticides recommended for other mirids and stink bugs (Pentatomidae) are effective, including some of the microbial and botanical products (Dara, 2014). The tomato bug threat to tomatoes, especially greenhouse-grown tomatoes, can be managed by adjusting the release rate of the predatory bugs, or by controlling them if the ratio of tomato bugs to prey becomes too high. Insecticides differ in their toxicity to tomato bugs when they feed on insecticide-treated prey, and although some materials do not cause direct toxicity, these insecticides may induce negative sublethal effects (Wanumen et al., 2016).

FAMILY PENTATOMIDAE—STINK BUGS Brown Marmorated Stink Bug Halyomorpha halys (Stål) (Hemiptera: Pentatomidae)

Natural History Distribution. This stink bug has long been a problem in Asia (principally China, Japan, Korea, and Taiwan), and was first reported in North America near Allentown, Pennsylvania in the mid-1990s. Since that time, it has been detected across much of the northern regions of the United States, and portions of southern Canada. Thus far, however, in the United States the serious damage is localized in the northeastern states from New Jersey to Virginia, although this insect is now established in Washington. It also now occurs in Europe. Undoubtedly, this vagile, adaptable insect that will continue to expand its range and impact though not likely to much of the South and arid West. Host Plants. Brown marmorated stink bug has a wide host range, feeding on tree fruits, vegetables, field crops, ornamental plants, and native vegetation. Hosts vary in suitability, and suitability changes during the season, so these bugs often disperse among crops. Adults normally feed on fruit, whereas nymphs feed on leaves, stems, and fruits (Hoebeke and Carter, 2003). Nielsen and Hamilton (2009) suggest that in its native range it affects about 300 species of plants, whereas Lee et al. (2013) indicate that a total of 106 plants in 45 plant families are hosts, most often from the plant families Fabaceae and Rosaceae. Hamilton et  al. (2018) listed 175 hosts in North America. Clearly, this species feeds on a great number of crop plants. Zobel et al. (2016) suggested that eggplant, okra, and bell pepper were most suitable for reproduction. They also noted that

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among vegetables, these crops plus sweet corn experienced the highest rate of injury per stink bug. Vegetable crops known to be injured include sweet corn, asparagus, snap bean, lima bean, tomatoes, green pepper, eggplant, and okra. Among the fruit crops injured are apple, peach, pear, plum, Japanese apricot, Asian pear, cherry, grape, fig, mulberry, persimmon, citrus, and blackberry. Among field crops, soybean is a favored host, but field corn, cotton, millet, and sorghum are attacked. Among the ornamental plants serving as hosts for brown marmorated viburnum, and crabapple. Lee et  al. (2013) provided a long list of hosts, with many plants designated as suitable for food, or for food plus reproduction. Natural Enemies. Indigenous stink bug parasitoids have been found to affect this exotic pest now that it has become established in the United States, but high levels of parasitism have not been recorded. Ogburn et al. (2016) reported less than 3% parasitism averaged over a large number of sites in the northeastern United States. Herlihy et al. (2016) similarly reported only 7%–10% parasitism. However, these latter authors reported the discovery of Trissolcus japonicus (Ashmead) (Hymenoptera: Platygastridae), which previously was not known to occur in North America. It now has been found on both the west and east coasts of North America. In laboratory tests, it parasitizes at least seven species of pentatomids native to Oregon, so it could prove disruptive to indigenous species (Hedstrom et  al., 2017). In China, effective parasitoids have been identified, particularly the egg parasitoid Trissolcus japonicus (Ashmead) (previously T. halymorphae Yang) (Yang et al., 2009; Talamas et al., 2013). Similarly, Trissolcus mitsukurii (Ashmead) (Hymenoptera: Platygastridae) is important in Japan (Lee et al., 2013). A list of natural enemies of H. halys found Asia is given by Lee et al. (2013). Predation of brown marmorated stink bug by three generalist predators was assessed in Canada using Chrysoperla carnea (Stephens) (Neuroptera: Chrysopidae); Coleomegilla maculata De Geer (Coleoptera: Coccinellidae); and Podisus maculiventris (Say) (Hemiptera: Pentatomidae) (Abram et  al., 2015). This study revealed that brown marmorated stink bug eggs were most readily consumed by C. carnea. Under field and laboratory conditions in West Virginia United States, a surprisingly wide range of predators, including some not often thought to be important predators, were observed to consume H. halys eggs, most notably katydids (Orthoptera: Tetigoniidae), ground beetles (Coleoptera: Carabidae), earwigs (Dermaptera), jumping spiders (Araneae: Salticidae), and crickets (Orthoptera: Gryllidae) (Morrison et  al., 2016). In Europe, the ant Crematogaster scutellaris Olivier (Hymenoptera: Formicidae) was shown to affect the survival of all preadult stages (Castracani et al., 2017). Life Cycle and Description. In China and Japan, there generally is but one generation, but two generations (or one and a partial second generation), may occur annually in

southern latitudes. Up to five generations are reported annually in tropical climates (Nielsen and Hamilton, 2009; Lee et al., 2013). In Pennsylvania, United States, only a single generation occurs. Although there are enough degree-days for two generations to occur, the overwintering adults must first mature, thereby limiting the period when reproduction can occur (Nielsen and Hamilton, 2009). As this insect spreads south an additional generation will occur, as is well documented in Europe (Costi et al., 2017). The minimum temperature for development is about 12°C (Lee et  al., 2013). When reared in the laboratory at constant temperature, the mean (± SE) number of days required for development of the egg through fifth instar decreased from 121.5 (± 0.5) to 33.4 (± 0.5) days as the mean temperature was increased from 17°C to 30°C, respectively. Temperatures above 30°C were less suitable (Nielsen et al., 2008). Brown marmorated stink bugs overwinter as adults in reproductive diapause. As temperatures and day lengths increase in the spring they become active (in about April) and reproductively mature (in May) and commence egg production. Adults are not immediately reproductively mature; after attaining the adult stage, about 2 weeks are required for adults to become sexually mature. In univoltine environments, they become mature in the autumn but usually, do not reproduce in northern areas. The fourth and fifth instars are sensitive to diapause induction. In bivoltine environments, the overwintering adults begin ovipositing in May, first- generation adults produce eggs in late June and into July, and second-generation adults are found between late August and into September. Overwintering adults seek dark locations for overwintering, where they remain until overwintering nutrients in the fat body are depleted. Structures such as homes and barns are commonly used as a shelter, but natural shelters such as tree holes and under loose bark and leaf litter are acceptable. The overwintering stink bugs become active between March and May when the ambient temperature exceeds 10°C. When photoperiod exceeds about 14 h, ovarian development occurs. Procedures for the culture of brown marmorated stink bug have been published (Medal et al., 2012; Rosen et al., 2016; Dingle and Jackai, 2017). Taylor et  al. (2017) determined that marmorated stink bugs triggered to enter diapause need cold storage (9°C) for 7 weeks to break diapause. A comprehensive treatment of brown marmorated stink bug was published by Hamilton et al. (2018). Egg. The female of brown marmorated stink bug deposits eggs clutches that contain about 28 eggs, which are deposited on the abaxial (lower) surface of leaves. They deposit an average of 240 (range 100–500) eggs over about 40–100 days, with clutches deposited at about 4–5-day intervals. The eggs hatch after 5–6 days when held at 25°C. Although survival rates vary considerably with time and

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location, the egg stage seems to experience the highest level of mortality (Costi et al., 2017). Nymph. There are five instars in the nymphal stage. First instars are about 2.4 mm long, and mostly orange or red. First instars, and to some degree second instars, remain aggregated near the eggs, before dispersing to seek food. After the first instar they become darker, sometimes black. A good field character for recognition is the presence of white bands on the antennae, with the distal band broader than the proximal band. Eventually, usually beginning with the second instar, white bands also are present on the tibiae. As the instars progress, they take on more and more of the adult metallic coloration. Also, the margin of the nymphal abdomen, and continuing to the adult stage, is trimmed with alternating white and black areas. The wing pads become elongate in the later instars, and fifth instars are typically about 12 mm long. In the field, nymphal development typically requires 44–52  days, depending on temperature. When Nielsen et al. (2008) reared insects established from a population in Pennsylvania, United States, at a constant temperature of 25°C, mean (± SE) nymphal development times were 4.8 (± 0.1), 9.6 (± 0.2), 7.1 (± 0.2), 5.9 (± 0.3), and 7.8 (± 0.3) days for instars 1–5, respectively, or a total egg to adult development time of 44.9 (± 0.8) days. In Switzerland, the equivalent study produced very similar development times, with a total egg to adult development time at 25°C of 42.3 (± 0.5) days (Haye et al., 2014).

FIG. 8.50  Adult of brown marmorated stink bug. (Photo by L. Buss.)

Adult. As with other stink bugs, this species has a shieldshaped body and is capable of emitting a pungent odor if disturbed. In the adult stage it is 12–17 mm in length. The mottled coloration is mostly various shades of gray or brown, but adults bear the white bands on the antennae and tibiae that were present in the nymphs. Also, in the adult stage the dorsal surface is of this bug is infused with patches of metallic copper and blue coloration. The edges of the posterior abdominal segments protrude from beneath the wings

and are alternately color brown and white. This species resembles the other common brown vegetable-infesting stink bugs (all Euschistus spp.) but the humerus (lateral edge of the pronotum) is rounded in H. halys, whereas it normally is pointed in the Euschistus spp. (E. servus, E. conspersus, and E. variolarius) McPherson and McPherson (2000) provide a key to economically important stink bugs, and McPherson produced keys to stink bugs in northeastern North America although they do not contain H. halys because it is a recent introduction. Also, a key to the most important vegetable-affecting stink bugs (including H. halys) is found in Appendix A.

Damage Brown marmorated stink bugs pierce the leaves and fruit of their hosts and remove sap. This results in death of adjacent tissue, producing small necrotic spots. In the case of damaged fruit, as the healthy nearby tissue expands the necrotic tissue does not, leading to formation of a dimple-like distortion. Alternatively, fruit may drop prematurely. Wounds also are susceptible to infection by plant pathogens. Brown marmorated stink bug also attains nuisance status by invading structures to seek suitable overwintering sites. With the onset of cool weather, these stink bugs enter buildings by finding small holes around doors and windows.

Management Sampling. Population monitoring is accomplished using pyramid traps, blacklight traps, and beating of vegetation though pheromone-based monitoring will likely supplant these other techniques soon. Morrison et al. (2015) presented comparative efficacy data on traps, mostly variants of pyramid traps. Nielsen et al. (2013) assessed blacklight traps, and found them to be sensitive tool over a range of conditions. This species typically infests trees and shrub, so unlike some other species of stink bugs, monitoring population on grass plants is inadvisable. In China, a cross-attractant aggregation pheromone produced by males of Plautia stali Scott (Hemiptera: Pentatomidae) is used to attract brown marmorated stink bug (Lee et al., 2013). This was also observed in Pennsylvania (Khrimian et al., 2008). However, more recently an aggregation pheromone of the brown marmorated stink bug has been described (Khrimian et al., 2014; Weber et al., 2014). This two-component pheromone attracts nymphs and adults, especially with the addition of a synergist, throughout most of the season (Leskey et al., 2015). Insecticides. Presently, management of brown marmorated stink bug depends mostly on application of insecticides. Residual insecticides are needed to kill these stink bugs because they are mobile, often moving from uncultivated to cultivated plants. They are easier to kill early in the season, apparently because overwintered insects

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are more susceptible to insecticides (Leskey et  al., 2013). Applications of insecticides in the morning are sometimes recommended because the bugs are less active and more likely to receive a lethal dose. Because the bugs are also found on uncultivated vegetation, application to edges of woodlands adjacent to crops can provide some benefit (Lee et  al., 2013). Generally, broad-spectrum insecticides are used for suppression of H. halys, although this is not compatible with most integrated pest management systems (Kuhar and Kamminga, 2017). If insecticides are to be used, selective products would be desirable so as not to disrupt biologically based pest suppression efforts directed at other pests in the cropping systems. Microbial insecticides have potential for use (Gouli et  al., 2012) but have not been proven effective under field conditions. Penca and Hodges (2017) evaluated the effects of a juvenile hormone analog, pyriproxyfen, on reproductive biology of the stink bug and its parasitoid Trissolcus japonicus. Pyriproxyfen disrupted diapause, leading to continuation of reproduction though hatchability of the eggs produced by treated adults. When parasitoids were provided with eggs from treated adults they were accepted and the parasitoids inserted eggs into their host’s eggs, but the parasitoids could not compete development. Thus, pyrifroxyfen potentially can disrupt H. halys population biology by interfering with diapause, but use of this approach may interfere with biological control. Cultural Practices. Nielsen et  al. (2016) studied potential trap crops during mid-June to September, with sunflower proving to be quite attractive to H. halys early in the period of peak bug emergence. However, sorghum supplanted sunflower later in the season, becoming more attractive after the sunflower had senesced. Stink bugs can overwinter in attics and garages as well as inhabited areas of buildings, becoming a nuisance when they become active. They can be prevented from entering most structures by sealing openings. Caulk or screening can be used, but the insects can enter through narrow openings so exclusion efforts must be implemented carefully.

Brown Stink Bug

Euschistus servus (Say) (Hemiptera: Pentatomidae)

Natural History Distribution. This native insect is found throughout the United States and southern Canada, and extends south into Mexico. Two subspecies are recognized by some authors: E. servus servus (Say) occurs in the southern states, including Virginia, southern Illinois, Texas, Arizona, and southern California; E. servus euschistoides (Vollenhoven) occurs in the northern portions of the continent. They meet in a broad band of intergradation from Maryland to Kansas. E. servus is the most abundant member of the genus in southern states.

Host Plants. This species has a wide host range. Among the vegetables attacked are bean, cabbage, corn, cowpea, okra, pea, pepper, squash, and tomato. Other crops that serve as hosts include such field crops as alfalfa, clover, cotton, lespedeza, oat, soybean, sweet clover, and timothy, and such fruit crops as apple, citrus, peach, pear, raspberry, and tobacco. Some of the weeds fed upon by brown stink bug are cocklebur, Xanthium sp.; curly dock, Rumex sp.; fleabane, Erigeron annuus; goldenrod, Solidago sp.; horseweed, Erigeron canadensis; ragweed, Ambrosia sp.; mullein, Verbascum thapsus; pigweed, Amaranthus sp.; prickly lettuce, Lactuca scariola; yellow thistle, Cirsium horridulum; and Canada thistle, Cirsium arvense. Mullein is reported to be especially important as it is present early in the season before many other hosts are available. Natural Enemies. Brown stink bug is parasitized by Telenomus and Trissolcus spp. (Hymenoptera: Platygastridae), Hexacladia smithi Ashmead (Hymenoptera: Encyrtidae) and several flies, including Gymnosoma fuliginosum Robineau-Desvoidy, Trichopoda pennipes (Fabricius), Cylindromyia binotata (Bigot), C. euchenor (Walker), C. fumipennis (Bigot), Gymnoclytia unicolor (Brooks), G. immaculata (Macquart), Euthera tentatrix Loew (all Diptera: Tachinidae), and Sarcodexia sternodontis Townsend (Diptera: Sarcophagidae) (McPherson, 1982). Yeargan (1979) studied mortality of brown stink bug eggs in Kentucky, and reported that when egg masses were distributed in crops about 50% were parasitized, 12% were destroyed by predators, and 20% failed to hatch probably due to undetectable natural enemy feeding. When stink bugs were allowed to oviposit naturally on plants 60% were parasitized, 37% were destroyed by predators, and 2% failed to hatch. Obviously, natural enemies have the potential to destroy a high proportion of stink bug eggs. Life Cycle and Description. There are two generations annually throughout the range of this species though in northern locations many late-developing nymphs of the second generation perish with the onset of winter. The adult is the overwintering stage. Overwintered adults usually commence egg laying in April or early May and continue through mid-June. The adults from the first generation produce eggs beginning in August, and the nymphs from this generation mature but do not produce eggs before the onset of winter. Thus, there are three peaks in abundance annually, beginning with the overwintering adults from the previous autumn, but only two generations per year. Egg. Females deposit eggs in clusters of about 20, but counts of 14 and 28 eggs per mass are especially common, probably reflecting ovariole number. Overwintered adults produce, on average, about 120 eggs per female, with over 500 being produced by an occasional individual. The egg production by first-generation adults is lower, usually 40–80 per female. The eggs are yellowish white

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and slightly ­greenish initially. They are somewhat barrelshaped, ­attached one end to a leaf and with a row of 30–35 small processes ringing the upper end of the egg. They average about 1.13 mm in length (range of 0.98–1.18 mm) and 0.96 mm in width (range of 0.86–1.18 mm). During warm weather the incubation period is typically 5–6 days (range about 3–14 days). Nymph. There are five instars, the mean durations of which were 3.7, 5.1, 4.9, 6.9, and 6.7  days, respectively, when reared on beans in Arkansas (Rolston and Kendrick, 1961) Munyaneza and McPherson (1994) reported similar values when reared at 23°C on beans in Illinois. In contrast, Woodside (1946) reported mean instar durations of 5.3, 9.3, 10.3, 13.0, and 13.6 days when reared on peach in Virginia. The difference in total nymphal development time, 33 vs 51 days, is not likely due solely to food plant effects, because the long egg incubation time of about 9 days in the Virginia study suggests cooler rearing conditions. Nymphal lengths are reported to average 1.5, 2.4, 4.2, 8.5, and 10.4 mm, respectively. The young nymphs are light colored dorsally, with the thorax yellowish brown and the abdomen white or yellow. The abdomen also bears red markings. The antennae are reddish brown. The older nymphs have substantially the same appearance, though the abdomen is always yellow. The wing pads become evident in instar four, and the pads overlap abdominal segments in instar five.

entire summer before perishing. The adult measures about 11–15 mm long, and is brown or grayish yellow. The abdominal segments, when viewed from below, bear black spots at the lateral angles. The ventral surface of the abdomen is brown in individuals that are entering reproductive diapause (usually fall adults) but green in those that will reproduce (usually spring and summer adults) (Borges et al., 2001).The lateral edges of the pronotum, or “shoulders,” are rounded. A key to distinguish stink bugs commonly affecting vegetables is found in Appendix A. The most complete life-history study was conducted by Rolston and Kendrick (1961), but McPherson (1982) provided an excellent summary of biology. Munyaneza and McPherson (1994) provided developmental data, description of the nymphal stages, and information on rearing. Aldrich et  al. (1991b) provided information on the chemistry of a volatile that attracts nymphs and adults of brown stink bug and other Euschistus spp., as well as stink bug parasitoids. Useful keys to Pentatomidae that include brown stink bug include McPherson and McPherson (2000), and McPherson (1982).

Damage Nymphs and adults feed on tender shoot tissues, buds, and fruit. Most damage to vegetables takes the form of deformed fruit, aborted blossoms, or death of young tissue. Sedlacek and Townsend (1988) and Apriyanto et al. (1989a,b) found that young corn plants were damaged by stink bug feeding, and corn kernels are also damaged, with the damage reflected in discoloration and abortion of kernels (Ni et al., 2010). Symptoms also include chlorotic lesions, tightly rolled leaves, wilting, stunting, increased tillering, delayed silk production, and smaller grain weight. The first instars, however, feed little or not at all—a common condition among stink bugs.

Management

FIG. 8.51  Adult brown stink bug. (Drawing by USDA.)

Adult. Adults from the first generation do not all go on to produce eggs. Over 90% of insects reaching adulthood after August 1 enter reproductive diapause. Even adults produced in June may enter diapause but the incidence is much lower. Adults successfully overwinter under plant debris and amongst weeds. In Arkansas, survival of adults sheltering in mullein was especially high. Open habitats, rather than wooded environments, seem to be favored (Jones and Sullivan, 1981). Longevity of overwintered adults in the spring usually is about 2 months, but some persist for the

Phillips and Howell (1980) stressed the importance of weeds in stink bug biology and noted that damage was higher in weedy areas, especially following senescence of the weeds. Reduced tillage practices and presence of cover crops seem to contribute on increases in stink bug abundance, probably because weeds are more abundant. Thus, if stink bugs are prone to be a problem, it is important to monitor weed populations. Nearby crops can also be a source of adults that suddenly disperse into more suitable and susceptible crops. Pheromones specific to Eustichus spp. can be used to capture and monitor abundance of this stink bug (Aldrich et al., 1991a,b). Foliar insecticides are effective to minimize crop loss, with special care warranted to protect the blossoms and fruit from feeding damage. Substrate-borne vibrational signals (“songs”) are produced by many stink bugs (including brown stink bug), with

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males and females producing song that differ among species and sex. Also, individual bugs produce more than one song (McBrien and Millar, 2003). These songs potentially could be used for sampling, but thus far a means to use them has not been developed (Lampson et al., 2010).

Consperse Stink Bug

Euschistus conspersus Uhler (Hemiptera: Pentatomidae)

Natural History Distribution. Consperse stink bug occurs in western North America from British Columbia and Idaho south to California and Nevada. It is a native species. Host Plants. Consperse stink bug damages vegetable and fruit crops. Known principally as pest of tomato, it also feeds on apple, apricot, blackberry, fig, loganberry, peach, pear, plum, and raspberry. It also is found occasionally in alfalfa, barley, cotton, sorghum, and sugarbeet. Among the numerous weeds known to support consperse stink bug are bracken fern, Pteridium aquilinum; dock, Rumex sp.; horehound, Marrubium sp.; mallow, Malva sp.; mullein, Verbascum sp.; plantain, Plantago sp.; mustard, Brassica sp.; and the thistles Cirsium occidentale and C. mohavense. Natural Enemies. Stink bug eggs are parasitized by Trissolcus and Telenomus (Hymenoptera: Platygastridae). The level of parasitism varies greatly, and the first generation is typically lightly parasitized. Digger wasps, Dryudella sp. (Hymenoptera: Sphecidae), prey on nymphs, and spiders are important predators of adults. In a study conducted in Washington, in addition to platygastrid egg parasitoids, Krupke and Brunner (2003) reported that Gymnosoma filiola Lowe and G. occidentalis Townsend (Diptera: Tachinidae) also parasitized consperse stink bug, though at low levels. Life Cycle and Description. In California, there are two generations annually, but only one generation is present in cooler areas. The adults overwinter in reproductive diapause and commence production of eggs in March or April. First-generation nymphs are found from April to June, and adults from June to September. Second-generation eggs are present from July to August, nymphs from July to October, and adults until the subsequent spring. Egg. Eggs are deposited in clusters of 7–28 eggs, with most clusters consisting of 13–15 eggs. They normally are arranged in four short rows of 3–4 eggs. Adults are not very discerning in their oviposition habits, selecting dry leaves and plants unsuitable for nymphal growth as well as suitable host plants. Initially pearly white in color, the barrelshaped eggs bear a fringe of spines around the top. The eggs turn pink as they mature. Duration of the egg stage is about

5–30  days under field conditions, and averaged 6.2  days when reared at 27°C. Females may produce up to 640 eggs during their life span though mean fecundity is about 225 eggs. Nymph. There are five instars. The early instars are black and reddish, but later instars are various shades of yellow and brown. Duration of the nymphal stage ranges from 23 to 79 days under field conditions. When cultured at 27°C, mean development time was 25.6  days. Mean duration of the instars was 3.1, 5.6, 4.3, 4.9 and 7.7 days, respectively, for instars 1–5. Temperatures of about 27°C seems optimal for development and oviposition. The first instar does not feed, and young nymphs remain clustered around the egg cluster during this instar.

FIG. 8.52  Adult consperse stink bug. (Drawing by USDA.)

Adult. The adult measures 11–12 mm long and 6–6.5 mm wide. It is grayish brown or greenish brown dorsally but sprinkled with numerous small black spots. The ventral surface is greenish, yellowish, or brownish. The legs are yellowish, though they also bear some black spots. The antennae are yellow or reddish, and darker at the tip. Adults overwinter under weeds and trash in the field. In the spring, overwintered adults feed on weeds and commence oviposition on both weeds and crop plants. Second-generation adults commence egg laying after a preoviposition period of about 10–32 days. The biology of consperse stink bug was given by Borden et  al. (1952) and Hunter and Leigh (1965). Development was described by Toscano and Stern (1976). Toscano and Stern (1980) described reproductive biology. Alcock (1971) described aggregation and mating behaviors. Aldrich et al. (1991b) provided information on the chemistry of a volatile that attracted nymphs and adults of consperse stink bug and other Euschistus spp., as well as stink bug parasitoids. A useful key to Pentatomidae that includes consperse stink bug is provided by McPherson and McPherson (2000). A key to distinguish stink bugs commonly affecting vegetables is also found in Appendix A.

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Damage Stink bugs feed on developing fruit, usually green fruit, causing white corky tissue to form beneath the skin at the site of feeding (Michelbacher et al., 1952). However, pink and red fruit also is readily attacked (Zalom et al., 1997). Dimpling or distortion of the fruit is common. Stink bugs seem to favor the stem end of tomato fruit, but they also feed elsewhere. Some fruit drop also occurs and stink bugs are implicated in transmission of pathogenic yeast though plant disease transmission has not been demonstrated specifically for consperse stink bug. Damage is often localized along the margin of a field because the bugs encounter these plants first as they disperse into a crop from overwintering shelter or weed hosts (Pease and Zalom, 2010). Sudden damage sometimes occurs when heavily infested fields, such as cotton or wheat, senesce or are harvested, resulting in dispersal of the adults into nearby crops (Reisig, 2011).

Management Sampling. Stink bugs can be sampled with a sweep net or by visual observation, often in association with a beating sheet or tray. However, in a study conducted in California, components of a male aggregation pheromone placed in traps did not show high levels of correlation with plant canopy shake samples. Despite lack of precise population estimation with pheromone-based sampling, pheromone traps proved useful for detecting population trends (Cullen and Zalom, 2005). Also, capture of these stink bugs is greatly enhanced by the presence of attractive host plants (Krupke et  al., 2001), suggesting that it may be necessary to add host volatiles to pheromone traps to optimize capture. Another possible complication is the attraction of some parasitoids to aggregation pheromone (Krupke and Brunner, 2003), as this may interfere with parasitism. Scouting of fields for consperse stink bug can be optimized by use of a phenology model that estimates the stage of insect development (Cullen and Zalom, 2000). Substrate-borne vibrational signals (“songs”) are produced by many stink bugs (including consperse stink bug), with males and females producing song that differ among species and sex. Also, individual bugs produce more than one song (McBrien and Millar, 2003). However, this has not yet been exploited for sampling purposes. Insecticides. Insecticides are applied to foliage and young fruit to protect susceptible crops from injury though stink bugs are quite difficult to kill. It is important to obtain good penetration of canopy foliage down to the soil level because these stink bugs often are found on lower portions of the plant. Border treatments are especially important, because this is the portion of the field injured most severely.

Cultural Practices. Some predators and parasitoids benefit from the presence of hedgerows, and then are available to disperse into crops. Stink bug abundance and ­damage to tomatoes can be greater adjacent to weedy areas, suggesting that weed and crop management can alleviate stink bug damage (Pease and Zalom, 2010). In a California study, predation (apparently by rodents) and parasitism of stink bugs in crops was higher close to the hedgerows but dissipated away from the hedgerows (Morandin et al., 2014).

Green Stink Bug

Chinavia hilaris (Say) (Hemiptera: Pentatomidae)

Natural History Distribution. Although most of the literature refers to the green stink bug as Acrosternum hilare (Say), the name has been changed to Chinavia hilaris (Say). Green stink bug, which is native to North America, occurs throughout the United States and southern Canada. Chinavia hilaris is less damaging, and therefore less well known than southern green stink bug, Nezara viridula (Linnaeus). Although green stink bug is readily confused with southern green stink bug, the distribution of the latter species is limited principally to the warm-weather southeastern states, plus California, and Hawaii). The taxonomy of some stink bugs remain unsettled, and Barman et  al. (2017) suggested that based on genetic diversity there may be a cryptic species within the “Chinavia hilaris” population of California. Host Plants. Green stink bug has a wide host range, though it is known principally as a pest of soybean and tree fruit. Among vegetables it has been observed to feed upon are asparagus, cabbage, corn, cowpea, cucumber, eggplant, lima bean, okra, mustard, pea, pepper, squash, snap bean, tomato, and turnip. Beans, particularly lima bean, are often damaged. Other common hosts are fruit such as apple, apricot, blackberry, cherry, elderberry, grape, mulberry, orange, pear, and strawberry; trees such as ash, basswood, black cherry, black locust, dogwood, hackberry, honey locust, holly, maple, and redbud; and field crops such as alfalfa, cotton, and soybean. Green stink bug may also be found on such weeds as common elder, Sambucus canadensis; goldenrod, Solidago spp.; jimsonweed, Datura stramonium; mallow, Malva spp.; mullein, Verbascum thapsus; and rattlebox, Crotalaria sp. Schoene and Underhill (1933) suggested that suitability of host plants changed through the season, and that stink bugs relocated as necessary. Availability of appropriate wild hosts early in the season is a prerequisite for high densities of green stink bugs in crops. McPherson (1982) provided a comprehensive list of hosts. Schoene and Underhill (1933) and Jones and Sullivan

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(1982) provided less complete lists but they indicated relative preference in Virginia and South Carolina, respectively. Natural Enemies. Green stink bug is subject to attack by several natural enemies; McPherson (1982) and Jones et al. (1996) provided lists of the insect enemies. Eggs are parasitized by several wasps, but the most important seem to be Telenomus podisi Ashmead, Trissolcus euschitis (Ashmead), and T. edessae (Fouts) (all Hymenoptera: Platygastridae), and Anastatus reduvii (Ashmead) (Hymenoptera: Eupelmidae). The late instar and adult bugs are attacked by Trichopoda pennipes (Fabricius) (Diptera: Tachinidae), a common parasitoid of hemipterans. Predators known to affect green stink bugs include predatory stink bugs (Hemiptera: Pentatomidae), green lacewings (Neuroptera: Chrysopidae), and various birds, particularly quail, Colinus virginianus. Yeargan (1979) studied the fate of stink bug eggs in two cropping systems and noted that green stink bug suffered less natural mortality than some other species of stink bugs. Also, mortality from unspecified chewing predators was more common than from parasitism, and mortality due to sucking predators was negligible. Life Cycle and Description. The number of generations appears to vary. In Virginia (Underhill, 1934) and Ontario and Quebec (Javahery, 1990) it is reported to be one, but in Arkansas (Miner, 1966), Kansas (Wilde, 1969), and Virginia (Kamminga et al., 2009) apparently there are two generations annually. In Arkansas, the first generation often occurs on dogwood, with the second generation attacking principally soybean. It would be easy to overlook the first generation, which may explain some of the disagreement about generation number. Development time is long, especially the adult preoviposition period, so generation number is likely limited to one in the north, but the climate of southern states should allow a second generation. The optimum temperature for development is reported to be 28.4°C; at 15°C and 35°C this species was unable to complete its development (Da Silva and Daane, 2014). Photoperiod, rather than temperature, may determine generation number. Wilde (1969) reported that increasing day length stimulated egg production, whereas decreasing day length inhibited egg production. The life cycle requires about 100  days when stink bugs are cultured at 22°C. The adult is the overwintering stage. Adults become active in the spring when temperatures exceed about 21°C and feed on young leaves and stems of trees. Kamminga et al. (2009) reported that there were three peaks of flight yearly, as based on black light trap captures in Virginia, but the first seasonal flight results from the previous year’s second generation, which overwinters in the adult stage. Egg. Oviposition in both Virginia and southern Canada does not occur until about mid-June though in the midwestern states oviposition in early June has been

observed. Eggs are normally deposited in clusters of about 30 eggs, though clusters of up to 69 eggs have been observed. The first egg cluster produced by a female is the largest, with subsequent clusters diminishing in size. The female is not very selective in her choice of oviposition site, with eggs deposited on both foliage and fruit. Mean egg production per female was estimated by Miner (1966) to average 60, but egg production of up to 170 per female was observed. The rearing conditions of Miner’s study were suboptimal, and egg production likely suppressed. Javahery (1990) obtained oviposition of 130–150 eggs per overwintered female, and this is probably a better estimate of fecundity. The mean incubation period at 22°C is 12.7  days (range 9–19  days). Incubation length decreases as temperature increases, with eggs hatching in only 6  days at 30–33°C. The eggs are deposited on end and in fairly definite rows, but the rows are not as tightly interlocked as in most stinkbug species. They are light green when first produced but turn yellow and then pink as they mature. The cap of the egg becomes red immediately preceding hatch. The eggs measure 1.22–1.53 mm long, and 1.04–1.29 mm in diameter. The egg is somewhat cup-shaped, the top being slightly broader than the base. The top is ringed with a row of 45–65 small-flattened processes (Esselbaugh, 1946). Bundy and McPherson (2000) provides descriptions of eggs, and a key to the genera of stink bugs based on eggs. Esselbaugh (1946) also describes eggs from several species. Nymph. The eggs from the overwintering adults begin to hatch in July though overwintered females continue to produce eggs throughout the summer. Nymphs tend to remain aggregated during the first instar, but disperse thereafter. Mean duration (range) of the five instars is 7.0 (5–10), 8.9 (6–14), 7.9 (6–11), 8.9 (6–11), and 12.8 (8–20) days, respectively, when reared at 22°C on soybean (Miner, 1966). Development times are shorter when reared on apple, pear, and snap bean at 23°C, averaging 6, 6–7, 6–8, and 8–9 days, respectively, for instars 1–5 (Javahery, 1990). Simmons and Yeargan (1988a) studied development over a range of temperatures and observed fastest development at 27°C, with the egg to adult stage completed in about 40 days, whereas nearly 50 days were required at 24°C and 30°C, and 70 days at 21°C. The nymphs measure about 1.6–2.0, 2.3–3.2, 4.0– 5.2, 6.8–8.2, and 10.0–12.7 mm long for instars 1–5, respectively. In color, the dorsal surface of instars 1–4 tends to be brownish black on the head and thorax, with a yellow spot centrally and yellow at the lateral margins of the thorax. The abdomen is marked with transverse black and light-blue stripes, and large black spots centrally. The lateral margins of the abdomen bear a row of semi-elliptical spots, blackish bronze in color. The fifth instar differs from the earlier instars, as the abdomen lacks stripes, and is rather uniform yellow green in color.

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FIG. 8.53  Green stink bug. (Photo by J. Carr.)

Adult. Nymphs from the first generation begin to attain the adult stage in August. Adults begin feeding within a few hours of attaining the adult stage, but mean days to copulation by adults is 22.3 (range 11–30  days). Mating requires only a few minutes but sometimes persists for several hours. Females usually mate after each egg cluster is deposited. Another 21.7 (19–33) days are required, following copulation, until egg production commences. Duration of the adult stage is about 2 months during the summer, but several months during the winter. With the onset of cold weather, adults disperse to wooded areas. Flight activity can be predicted by mean monthly precipitation and the number of days below freezing (Kamminga et  al., 2009). Overwintering occurs in leaf litter and under bark of trees in wooded areas. The adult bugs measure 13–19 mm long. They are largely uniform green in color, though the dorsal surface is darker than the ventral, and the edges of the head, pronotum, and abdomen are marked with yellow. The three distal segments of the antennae are marked with brownish black. Southern green stink bug, N. viridula (Linnaeus), is easily confused with green stink bug, A. hilare. However, the two species differ ecologically and morphologically. Acrosternum hilare is usually associated with trees and shrubs, rather than the herbaceous vegetation fed upon by N. viridula. Also, A. hilare is found widely in North America though it is most abundant in the north. The two species can be differentiated by the shape of the abdominal spine. When viewed from below, A. hilare has a pointed spine protruding forward between the base of the hind legs whereas in N. viridula, the spine is rounded. Male green stink bugs produce an aggregation pheromone that attracts both males and females, other stink bug species, and the tachinid parasitoid Trichopoda pennipes (Aldrich et  al., 1989). The pheromone consists of the same chemicals found in the pheromone of southern green stink bug, but the ratios are different. Like other stink bugs, substrate-borne vibrational signals are an important part of

their communication system, with the songs differing between sexes and eliciting varying types of behavior (Cokl et al., 2001). The biology of the green stink bug was given by Whitmarsh (1917), Underhill (1934), Miner (1966), and Javahery (1990). Developmental biology was described by Simmons and Yeargan (1988a). Nymphs were described by Whitmarsh (1917) and Decoursey and Esselbaugh (1962). Culture on plant tissue was provided by many authors, such as Wilde (1968) and culture on artificial diet was presented by Brewer and Jones (1985). A recent summary of biology and management was provided by Kamminga et  al. (2012a). Useful keys to Pentatomidae that include green stink bug include McPherson and McPherson (2000) and McPherson (1982). A key to the common stink bugs found in vegetables is included in Appendix A.

Damage The green stink bug inserts its beak into fruit or foliar tissue and removes liquids. During feeding, the bug injects enzymes that liquefy plant tissue and cause the tissue at the feeding site to collapse. Subsequently, tissue adjacent to the feeding site continues to grow, but the tissues at the feeding site fail to grow, leading to dimples or similar deformities. Feeding sites also may become discolored, usually turning black and hardened, but sometimes white and spongy. Adults and nymphs inflict the same type of injury, but the nature of the injury varies among plants and its stage of growth. Damage potential varies considerably among insect stages. The first instar does not feed. Feeding frequency increases as the nymph matures, with the fifth instar feeding most frequently, an average of 3.4 times per day. Duration of each feeding session does not vary with bug age, however, averaging 1.5 h per session. The adults and fifth instars, and both sexes, displayed similar feeding behaviors (Simmons and Yeargan, 1988b). Plant responses to green stink bug feeding have not been well studied, but some states recommend treatment of beans if stink bug abundance attains one bug per 5 m of row. Stink bugs often move from crop to crop during the growing season, generally relocating to take advantage of the availability of new plant reproductive tissues. The behavior of the different species is not identical, however. For example, in a study of stinkbug infestations in Georgia farmscapes involving corn, cotton, and peanuts, Nezara viridula (L.) and Euschistus servus (Say) were more likely to oviposit initially on corn, then move to peanut and cotton. However, in these same fields, corn and peanut were not very suitable host plants for C. hilaris. All three species favored feeding on cotton (Tillman, 2013). Noncrop hosts are particularly important to C. hilaris, relative to these other two species (Cottrell and Tillman, 2015).

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Green stink bug harbors Nematospora coryli, a fungus that causes yeast-spot disease in beans and soybeans. Both adults and nymphs acquire the pathogen by feeding, retain it in their bodies, and excrete it in their saliva and feces (Foster and Daugherty, 1969; Clarke and Wilde, 1970). The pathogen greatly exacerbates the damage caused by green stink bug. Underhill (1934) reported the loss of entire lima bean crops due to this disease and its insect vector. Other fungi and bacteria can be transmitted by stink bugs (Russin et al., 1988), so the vector potential of green stink bug may not be limited to transmission of N. coryli.

Management Sampling. Stink bug populations tend to be highest at field margins, probably reflecting the tendency of adults to overwinter in wooded areas surrounding fields, and the early-season feeding preference for trees. Sampling in soybean is usually accomplished by shaking the plant over soil or a drop cloth, and counting the insects dislodged, but visual examination may be appropriate for crops with less vegetation. Sweep-netting, blacklight traps, and use of traps baited with aggregation pheromone (usually a yellow-based pyramid trap) are additional options for population monitoring. Unfortunately, the pheromone is expensive and also attracts southern green stink bug, N. viridula (Kamminga et al., 2012a,b). In all crops, sampling for eggs is done by visual examination. Insecticides. Stink bugs often are difficult to kill, but proper selection and timing of foliar insecticides can protect crops from injury. Susceptibility often varies with the age (stage) of the bugs. Residual materials are usually desirable because the stink bugs often develop outside the crop and enter at various times. It is most important to protect the crop during the blossom and early fruiting stage. Because bugs are entering from the crop margins, border treatments may be adequate. Cultural Practices. Stink bug populations increase during the season, with maximum densities and damage occurring late in the growing season. Stink bugs commonly enter crop fields from adjacent vegetation, either woodlands or weedy areas. Thus, a common recommendation is to plant as early as possible to avoid peak insect abundance. However, in studies conducted with soybean, early plantings (early June) were as damaged as late (early July) (McPherson et al., 1988). Jones and Sullivan (1982) suggested that the destruction of black cherry and elderberry would greatly reduce abundance of green stink bug. Trap crops are also sometimes recommended for stink bugs. For information on this subject, see the discussion on trap crops in the section on southern green stink bug. Pole beans are considered more susceptible than bush varieties (Underhill, 1934).

Harlequin Bug

Murgantia histrionica (Hahn) (Hemiptera: Pentatomidae)

Natural History Distribution. A native of Mexico and Central America, harlequin bug was first observed in the United States in Texas, in 1864. Its appearance, coinciding with the occurrence of Union troops during the American Civil War, earned it the name “Sherman-bug” and “Lincolnite” in parts of the south. It rapidly spread throughout the southern states, and eventually reached northern locales such as Colorado, Iowa, southern Michigan, Pennsylvania, and Massachusetts. It is considered to be a serious pest only in southern states, however, is not regarded as a problem in California. Harlequin bug is also a pest in Hawaii. DiMeglio et  al. (2016) assessed survival of harlequin bug during the winter months and in the laboratory at low temperatures. Temperatures below freezing could be lethal to most bugs, though by sheltering beneath leaf litter they were somewhat protected by the warmer temperatures found there. Eggs and young nymphs though not normally the overwintering stages, were more cold-hardy than older bugs. Thus, wintertime survival is better in warmer climates. Host Plants. Harlequin bug is principally a pest of crucifers (Brassica spp.), including broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, collards, kale, kohlrabi, mustard, radish, rutabaga, turnip, and watercress. This bug is reported to be especially fond of horseradish. In the southernmost states, crucifers do not thrive during the summer months and the bugs are forced onto other plants. Thus, they are sometimes found feeding on beans, okra, squash, tomato, and many other vegetables, but this is usually due to lack of normal food. Harlequin bug feeds readily on cruciferous weeds such as wild mustard, Brassica spp.; shepherd’s purse, Capsella bursa-pastoris; and pepperweed, Lepidium spp.; and related mustard oil-containing plants such as members of the family Capparaceae. Other weeds common in crops, such as pigweed, Amaranthus spp.; and lambsquarter, Chenopodium album; are also fed upon and reproduction occurs on these plants. Natural Enemies. Harlequin bug appears to be relatively free of natural enemies, other than for egg parasites and general predators. The egg parasitoids are Oencyryus johnsoni (Howard) (Hymenoptera: Encyrtidae), Trissolcus murgantiae Ashmead, and T. podisi Ashmead (both Hymenoptera: Platygastridae). The best-known species is O. johnsoni, which is reported frequently from harlequin bug eggs and caused up to 50% mortality during a harlequin bug outbreak in Virginia (White and Brannon, 1939). This parasite is widely distributed and apparently has other hosts. It attacks eggs in all stages of embryonic development, and prevents

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them from hatching (Maple, 1937). However, O. johnsoni is not the only effective parasite, as T. murgantiae was observed to parasitize 45% of harlequin bug eggs in North Carolina, at locations where O. johnsoni parasitized only 30% of eggs (Huffaker, 1941). Because of its effectiveness, T. murgantiae was introduced into California (DeBach, 1942). Life Cycle and Description. Harlequin bug breeds continuously in the southern portions of its range. During mild winters, all stages have been observed as far north as Virginia. In colder climates, only the adults survive the winter in sheltered locations. They seek shelter in and near fields, among overwintering crop plants, and in other organic debris such as dead leaves and bunches of grass. Development from the egg to adult stage requires about 48 days at 24°C (Zahn et al., 2008b). Two or three generations per year seem normal, but Paddock (1915a) indicated four generations in south Texas. These insects produce chemicals from their scent glands that apparently inhibit predation (Aldrich et al., 1996). Egg. The adults begin depositing eggs about 2  weeks after becoming active in the spring. Eggs are deposited beneath leaves, usually in clusters of 12 arranged in two rows of six, at intervals of 5–6  days. As the female nears the end of her life, the egg batches get slightly smaller and the arrangement less regular. The eggs are barrel shaped and measure about 1.30–1.38 mm long and 0.90–0.92 mm in diameter. They are light gray or pale yellow, and generally are circled by two black bands. They may also bear small black dots or spots, and the top has a semicircular black marking. The average number of eggs is reported to be 115 per female. Egg deposition may occur over a period of about 40–80 days. Eggs hatch in about 4–5 days during warm weather, while 15–20  days may be required during cool weather. Nymph. Upon hatching, young nymphs stay clustered near the old eggs for 1–2 days. The newly hatched nymphs are pale green with black markings but soon become brightly colored—black or blue, with red and yellow or orange markings. Paddock (1918) indicated that there were six instars in Texas, and that nymphal development could be completed in just 30 days. He gave average development times as 3.4, 3.2, 4.7, 4.7, 7.0, and 4.3 days, respectively, for the six instars developing under summer conditions. Under spring conditions, development times were increased by about 30%. At 24°C, Zahn et al. (2008b) reported that mean development time of the nymphal instars was 3.3, 4.5, 8.9, 12.9, and 14.4 days for instars 1–5, respectively. This is consistent with the report of White and Brannon (1939), who observed a development time of about 40–60 days during the summer, and slightly longer, perhaps 70 days, during cool weather.

FIG 8.54  Harlequin bug. (Photo by J. Capinera.)

Adult. The adults usually live about 60  days, but may live considerably longer during the winter. They measure about 8.0–11.5 mm long. The adults are brightly colored, similar to the large nymphs, principally black and yellow or black and red. The color pattern varies, with the spring and summer bugs more brightly colored than the overwintering insects. As with many stink bugs, harlequin bugs produce a disagreeable odor if disturbed, and birds avoid eating them. The biology of harlequin bug was provided by Chittenden (1908b), Paddock (1918), and White and Brannon (1939). Reviews of biology and management were published by Wallingford et  al. (2013) and McPherson et  al. (2018). Useful keys to Pentatomidae that include harlequin bug include McPherson and McPherson (2000) and McPherson (1982). A key to distinguish stink bugs commonly affecting vegetables is found in Appendix A.

Damage The piercing-sucking feeding behavior of this insect results in white blotches at the site of feeding. Wilting, deformity, and plant death may occur if insects are abundant. Mild winters are said to favor survival and subsequent damage (Walker and Anderson, 1933). Once considered the most serious crucifer pest in the south, this insect has been relegated to minor status in commercial production and persists mostly as a home garden pest. However, the broad-spectrum insecticides once used for crucifer insect control are no longer so popular, and some of the more selective insecticides are not effective on harlequin bug (Wallingford et al., 2013).

Management Sampling. Harlequin bug abundance is normally determined by visual examination of plants. However, they are attracted to black and green, so trapping devices might be useful for these insects. Also, males produce an aggregation pheromone that attracts both males and females (Zahn et al., 2008a).

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Chemical Control. Insecticides are applied to the foliage for suppression of this bug. Harlequin bug can be difficult to control with insecticides; targeting the young bugs and thorough coverage are recommended (Rogers and Howell, 1973). Adjuvants are often used to enhance sticking of liquid insecticides to crucifer crops. Soil applications of systemic insecticides have also become popular. Soap applied alone or in combination with rotenone was reported to provide good control (Walker and Anderson, 1933, 1934), but many biorational or selective insecticides are not very effective against this insect. Cultural Practices. Trap crops, usually consisting of early-planted mustard, rape, or kale are sometimes recommended to divert the overwintering bugs from the principal crop. A comparison of these three crops demonstrated that mustard and rape prevented harlequin bugs from attaining broccoli plots, though they were effective only at low bug densities (Ludwig and Kok, 1998). Wallingford et al. (2013) also found mustard to protect collards from damage. Such trap crops must be sprayed or destroyed, however, or the adults will soon move to the main crop. Destruction of crop residues, on which the insect may overwinter in the north or oversummer in the south, is an important cultural practice to alleviate harlequin bug damage. Host-Plant Resistance. Sullivan and Brett (1974) studied the relative susceptibility of different crucifer crops to harlequin bug in North Carolina. They reported that mustard and Chinese cabbage were the most susceptible; turnip, kale, rutabaga, and some radishes were intermediate; and cauliflower, cabbage, broccoli, collards, Brussels sprouts, kohlrabi, and most radish varieties were fairly resistant. Cabbages were the most resistant crop, but considerable variation among cultivars was evident.

Onespotted Stink Bug

Euschistus variolarius (Palisot de Beauvois) (Hemiptera: Pentatomidae)

Natural History Distribution. Onespotted stink bug is native to North America. It is widely distributed in the United States; it is absent only from the southwestern states, apparently including California. In the midwest and other northern areas, it is considered to be the most common stinkbug species (Esselbaugh, 1948). In Canada, it is known from Quebec and Ontario west to British Columbia. Host Plants. This species is reported to feed on numerous plants. Among vegetable crops attacked are asparagus, bean, cantaloupe, corn, cowpea, mustard, onion, pea, potato, squash, and tomato. Field crops that serve as hosts are alfalfa, clover, cotton, oats, rye, sugarbeet, timothy, tobacco, and wheat. Such fruit crops as gooseberry, grape, peach, pear, and raspberry are damaged. Some of the numerous weeds known to support this species are burdock, Arctium sp.; curly dock,

Rumex sp.; goldenrod, Solidago sp.; horseweed, Erigeron canadensis; milkweed, Asclepias sp.; mullein, Verbascum thapsus; pigweed, Amaranthus sp.; ragweed, Ambrosia sp.; and thistle, Cirsium sp. Trees are commonly reported as hosts; among them are black walnut, elm, pine, poplar, sassafras, tulip tree, and willow (McPherson, 1982). Natural Enemies. Onespotted stink bug is parasitized by Telenomus and Trissolcus spp. (Hymenoptera: Platygastridae and several flies, including Gymnosoma fuliginosum Robineau-Desvoidy, Trichopoda pennipes (Fabricius), Cylindromyia binotata (Bigot), C. fumipennis (Bigot), Gymnoclytia occidua (Walker), Cistogaster immaculata Macquart, and Euthera tentatrix Loew (all Diptera: Tachinidae) (McPherson, 1982). Yeargan (1979) studied mortality of brown stink bug eggs in Kentucky and reported that when egg masses were distributed in crops, about 50% were parasitized, 13% were destroyed by predators, and 25% failed to hatch probably due to undetectable natural enemy feeding. When stink bugs were allowed to oviposit naturally on plants, 71% were parasitized, 26% were destroyed by predators, and 1% failed to hatch. Natural enemies clearly have the potential to destroy a high proportion of stink bug eggs. Life Cycle and Description. There is only a single generation per year throughout the range of this species. The eggs are deposited in late April to June, with adults produced by August. The adults overwinter, and both new and overwintering adults can be found late in the summer. Egg. The eggs are barrel shaped, and attached on end to the underside of a leaf. The upper end of the egg is ringed with a single row of 27–33 small spines. Initially, the eggs are light green, but soon turn creamy white. They measure about 1.1 mm long and 0.9 mm wide. Females deposit the eggs in clusters; the average number per cluster is about 14. The incubation period of eggs is about 6  days (range 3–10 days).

FIG. 8.55  Onespotted stink bug.

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Nymph. There are five instars. The mean length (range) was reported by Munyaneza and McPherson (1994) to be 6 (6), 11 (9–14), 10 (8–12), 12.6 (7–15), and 10.6 (8–17) days, respectively. Esselbaugh (1948) reported mean (range) development times of 3.5 (2–11), 7.5 (4–16), 7.9 (5–17), 7.7 (5–13), and 10.0 (8–16), respectively. The first instars are grayish brown on the thorax, the abdomen similar but lighter and marked with red. First instars measure about 1.5 mm long. Second instars are whitish, and conspicuously marked with black spots on the head and thorax and red on the abdomen. They measure about 2.8 mm long. Third instars are similar to second instars in appearance, except that the abdomen is greenish and they measure about 4.1 mm long. Fourth instars are greenish, with a row of red spots along the median line of the abdomen. The wing pads are evident in this instar, which measures about 6.7 mm long. The fifth instar has a dark yellow head and thorax, and a green abdomen. The wing pads are well developed, overlapping the first three abdominal segments. This instar measures about 10.3 mm long. The nymphal instars of E. variolaris are very similar to those of brown stink bug, E. servus (Say). Adult. The adult bugs are grayish brown. They measure 11.0–15.0 mm long. The lateral edges of the pronotum, or “shoulders,” generally taper to a sharp point. In males of this species the ventral surface of the tip of the abdomen is marked with a single dark spot, which serves as the basis for the common name. Because it is sex specific, it is not a particularly useful diagnostic character. The adults overwinter under dry leaves and other plant debris, under logs, and beneath mullein leaves. Winter mortality is high, with 80% or more of the population perishing. Bugs begin mating a few days after emerging from diapause in the spring. The average time from copulation to egg laying is 27 days. The biology and description of the nymphs were given by Parish (1934) and Munyaneza and McPherson (1994). The latter authors also gave information on rearing and characters to distinguish onespotted stink bug nymphs from brown stink bug nymphs. Useful keys to Pentatomidae that include onespotted stink bug include McPherson and McPherson (2000) and McPherson (1982). A key to distinguish stink bugs commonly affecting vegetables is also found in Appendix A.

Damage Nymphs and adults feed on tender shoot tissue, buds, and fruit. Most damage to vegetables takes the form of deformed fruit, aborted blossoms, or death of young tissue. Annan and Bergman (1988), Sedlacek and Townsend (1988), and Apriyanto et  al. (1989a,b) found that young corn plants were damaged by stink bug feeding. Symptoms included are chlorotic lesions, tightly rolled leaves, wilting, stunting, increased tillering, delayed silk production, and smaller grain weight. Even a single day of feeding by stink bugs reduced corn growth and yield. The first instars, however, feed a little or not at all—a common condition among stink bugs.

Management Phillips and Howell (1980) stressed the importance of weeds in stink bug biology and noted that damage was higher in weedy areas, especially following the senescence of the weeds. Reduced tillage practices and the presence of cover crops, especially wheat, contributed to an increase in stink bug abundance. Foliar insecticides are effective, with special care needed to protect the blossoms and fruit.

Painted Bug

Bagrada hilaris (Burmeister) (Hemiptera: Pentatomidae)

Natural History Distribution. Painted bug (sometimes called “bagrada bug”) has long been known from eastern and southern Africa, the Middle East, and southern Asia (Pakistan and India). It was first described from specimens collected in India. It is abundant and damaging mostly in temperate areas. In North America, it was first observed in 2008 in southern California and soon spread to Hawaii, Arizona, Nevada, Utah, New Mexico, and Texas (Taylor et al., 2015). It is expected that it will develop into a pest in other areas of North and Central America. It is now reported to be a pest in parts of southern Europe and Mexico as well. This insect acquired the name bagrada bug almost immediately upon discovery in California, but this name is a poor choice because it is based on the genus name. This insect has been moved from the genus to the genus over time (a common occurrence with scientific names) so it may be moved again. Thus, a name independent of the genus name, such as painted bug, has been suggested (Palumbo et al., 2016), and “painted bug” is used widely in Asia. Host Plants. The painted bug reportedly has a wide host range. Taylor et al. (2015) and Palumbo et al. (2016) provided host lists (feeding or association) of over 70 plant species from over 20 plant families, and Bundy et al. (2018b) provided a list of 96 hosts. It is mostly known as a pest of brassica (cole) crops (Brassicaceae or Cruciferae), including vegetables such as arugula, broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, collards, kale, radish, rutabaga, and turnip, and field crops such as oilseed brassicas. During periods when painted bug are very abundant, they have been reported to be quite destructive to some grains such as maize (corn), pearl millet, and wheat. However, these latter food plants are not suitable for development, but serve to keep the insects alive until more suitable hosts are located or become available. Certain ornamental Brassicaceae such as sweet alyssum, Labularia maritime; candytuft, Iberis spp.; and stock, Matthiola incana, also are readily infested. Important weed host include wild mustards such as perennial pepperweed, Lepidium latifolium; shepherd’s purse, Capsella bursa-pastoris; tansy mustard,

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Descurainia sophia; wild radish, Raphanus raphanistrum; and London rocket, Sisyrmbrius irio. The seasonal abundance often is linked to the period of growth of annual wild mustards. Lambert and Dudley (2014) reported that perennial pepperweed was especially favored by painted bug in the autumn months, and even suffered high levels of feeding injury from this insect in southern California. Reed et  al. (2013) conducted feeding studies using plants typically found in southern California. They reported that painted bug feeds readily on all hosts in the plant family Brassicaceae. In the presence of brassica hosts, the bugs generally avoided feeding on plants in the families Asteraceae, Cucurbitaceae, Malvaceae, and Solanaceae. Some feeding occurred on plants in the families Fabaceae and Poaceae. In fact, corn and Sudan grass were significantly damaged. Huang et  al. (2014) also provided painted bugs with a choice among several prospective hosts in different forms of choice tests and noted that radish was especially attractive, followed by cabbage, broccoli, kale and cauliflower, and then by other plants. There was not a consistent difference displayed in these studies for plant age (cotyledon versus one-, two-, and four-leaf stages), although authors commonly state that the youngest plants are most damaged. Natural Enemies. Knowledge about natural enemies of painted bug is quite limited. Several parasitic wasps (Hymenoptera) have been identified, mostly from the genera Telenomus, Paratelenomus, Trissolcus, Gyron (all Playgastridae), and Ooencyrtus (Encyrtidae). In India and Pakistan, parasitism rates of 15%–30% have been observed, with soil-covered eggs less susceptible to parasitism than those above-ground. Native parasitoids are not adapted to parasitizing eggs in the soil, so it seems unlikely that they will be of great benefit. The effectiveness of dipteran parasitoids that affect nymphs and adults of painted bug also need to be determined. Research is underway, however, to import and screen candidate parasitoids from Asia (Palumbo et al., 2016). Native predators are deterred by the chemical defenses of stink bugs, but this is a general phenomenon, not limited to painted bug. Thus, they cannot be expected to exert important levels of control. Life Cycle and Description. The life cycle of painted bug is not much different from that of other pentatomids, except that they are aposematically colored, so they often form aggregations because they seemingly are protected from predation. Also, they often are active throughout the year if there are plant hosts available. They are favored by warm, dry conditions, however, and normal development generally requires a temperature of 16–40°C. Survival is reported to be optimal at temperatures from 24°C to 35°C, and development occurs fastest at 35°C (Reed et al., 2017). The number of degree days needed for full development is estimated to be 284. With favorable temperatures and suitable food, the life cycle can be completed in 14–18 days, theoretically resulting in up to 10 generations per year. Because

in the United States they presently occur only in areas with relatively warm winters, a period when their hosts are most abundant, they are most abundant and most active during the winter. However, with the exception of eggs, all stages can be present throughout the year in New Mexico (Taylor et al., 2015). In New Mexico, they alternate predominately between London rocket and broccoli in the winter and mesa pepperwort, Lepidium alyssoides (Brassicaceae) in the summer. Adults can be found throughout the year on pepperwort, but the other stages occur discontinuously. In southern New Mexico, only two generations (and possibly a possible partial third generation) occur per year. Thus, despite the potential for more generations, availability of food resources likely constrains the biotic potential of this species. The activity and seasonality of this insect varies depending on the cropping system, and often there are two peaks of activity corresponding to the planting/germination stage and the harvest stage of brassica crops (Palumbo et  al., 2016). In warm season areas, senescence of brassica crops is followed by movement of the bugs to non-brassica host. Feeding and mating have been observed to occur mostly in the afternoon. This corresponds to the warmest period of the day, and temperature is a more reliable indicator of activity than other environmental factors (Huang et al., 2013). Egg. The barrel-shaped eggs are usually deposited singly or in small numbers in loose soil, and only occasionally on the leaf, flower or stem, or on debris such as dried leaves. The female actively buries the eggs. Soil particles typically adhere to the chorion. With the exception of this species, the pentatomids found in the New World deposit clusters or rows of eggs on foliage. Soil deposition of eggs occurs with some other Old World pentatomids, as well. When first deposited, the eggs are whitish or light brown, turning dark pink within 5 days. When maintained at 25°C, mean duration of the egg stage was 7.5 days (range 6–9  days), but in as little as 4  days under warmer conditions, and up to 20  days during winter. Mean egg length is 0.93 ± 0.03 mm (± SE) (range 0.87–1.0 mm) and width is 0.77 ± 0.02 mm (range 0.55–0.75 mm). Females can deposit 100–150 eggs over a 3-week period. Nymph. There are five instars in the nymphal stage. The head and thorax are reddish brown, with the abdomen reddish dorsally. After molting, the nymphs are orange red, but the legs, head, and thorax soon darken. Beginning with the third instar, two color forms, a lighter or a darker form, may occur. The mean lengths (± SE) of instars 1–5 are 1.19 ± 0.03 mm, 1.50 ± 0.05 mm, 2.30 ± 0.04 mm, 3.22 ± 0.05 mm, and 4.62 ± 0.21 mm, respectively. Other descriptive characters are provided by Taylor et al. (2015). Mean duration of the instars (± SE) when reared on mesa pepperwort at 25°C is 3.35 ± 0.05, 7.08 ± 0.13, 6.39 ± 0.13, 7.33 ± 0.18, and

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10.25 ± 0.16 days for instars 1–5, respectively. Duration of the egg through instar 5 is 41.85 ± 0.36 days, with male and female development times being essentially the same. The nymphs tend to remain aggregated, and also aggregate with adults. The first instar does not feed.

Sometimes the plants are deformed, so head-forming crops such as broccoli can experience significant losses. On older plants, insertion of their piercing-sucking mouthparts and secretion of digestive enzymes causes stippling and wilting of the plant tissue. On leaves and stems, starburst-shaped lesions often develop. When the bugs are abundant, and especially when the supply of brassica crops or weeds declines, these insects may move onto crops of other plant families and cause injury. This is often viewed simply as “bridging” behavior, allowing the insects to survive until a reproductive host is available.

Management

FIG. 8.56  Painted stink bug. (Photo by J. Carr.)

Adult. The adults are shield shaped, as with most pentatomids. They are black with orange and white markings, about 5–7 mm long, and 2.5–4.0 mm wide. They resemble harlequin bug, Murgantia histrionica (Hahn), with which they are easily confused, but they are considerably smaller than harlequin bug (which is 8–11.5 mm long). The females are larger than the males, averaging about 7 and 6 mm in length, respectively. Adult longevity is up to about 30 days when feeding on preferred host plants, but can be extended to nearly 100 days when feeding on nonpreferred hosts (Palumbo et al., 2016). Females generally survive longer than males. Typically, the female becomes receptive to males 1–2 days after attaining the adult stage, and oviposition commences 4–5 days after the initial mating. McPherson and McPherson (2000) provide a key to economically important stink bugs, although it does not include B. hilaris. A comprehensive treatment of painted bug was published by Bundy et al. (2018b). Also, a key to the most important vegetable-affecting stink bugs (including B. hilaris) is found in Appendix A.

Damage Painted bug has rapidly developed into a very serious pest in the southwestern United States, damaging a number of brassica crops. They are particularly damaging the small plants, so direct-seeded crops are especially susceptible, though transplants are also damaged. They feed on leaves, stems, flowers, and seeds. They feed by repeatedly inserting their stylets between the epidermal layers of the leaf, damaging the cellular tissue, and removing sap. Infested plants wilt and desiccate, and the apical meristem is killed.

Sampling. Visual examination of plants and weeds can be used to assess damage potential by counting the bug population. Inspections should be made immediately upon emergence of seedlings from the soil. Unlike most stink bugs, this species does not spend all of its time on host plants, instead seeking shelter beneath weeds, debris, and clumps of soil at night and during the heat of the day. This has serious implications for population assessment, as it would be easy to underestimate bug density. Also, ­starburst-shaped lesions are diagnostic of painted bug feeding. Painted bugs can be attracted to pyramid-shaped stink bug traps, but the lure normally used for stink bugs is not attractive to painted bug, so the traps should be baited with an attractive plant such as sweet alyssum. Studies with crossvein traps suggest that black traps are more attractive than purple, red, yellow, or white traps (Joseph, 2014). There is evidence for volatile and contact pheromones (Guarin et  al., 2008). Females are attracted to odors from sexually mature males. The sex pheromone may also function as a defensive compound. The chemistry of painted bug communication is, as yet, not well defined and a pheromone lure has yet to be synthesized and evaluated (Palumbo et al., 2016). Insecticides. Currently, the principal method of protecting crops from damage by painted bug is application of insecticides. Several insecticides or insecticide rotations have been identified that provide effective control, but after suppression, plants are always susceptible to reinvasion if there are untreated populations of bugs nearby. Also, it is imperative to protect the very young plants, so systemic insecticides are applied immediately after seeding, or to transplant plugs (Joseph et al., 2016). Insect growth regulators can be effective in suppression of painted bug, but they must be applied to the nymphs because the adults are not affected (Joseph, 2017). Botanical insecticides have provided erratic results. Cultural Practices. Use of transplants rather than direct seeding of crops can help avoid some of the direct mortality that typically results when bugs feed on seedlings. Mesh or floating row covers can be used to prevent access by painted bug to small plots of susceptible crops. However, it is important that the mesh may not be in contact with the plant or the

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bugs will insert their mouthparts through the mesh to feed. Favored weed species, especially in the family Brassicacea, should be removed or killed. Fallen leaves and other debris should also be removed, as these materials provide shelter for the bugs. Timely destruction of crop residues and timing of planting can also be used to minimize injury. Indian mustard has been used as a trap crop. By planting this attractive plant around other, less attractive brassica crops, the insects are lured away. High levels of nitrogen fertilization exacerbate painted bug problems, so should be avoided.

Redbanded Stink Bug

Piezodorus guildinii (Westwood) (Hemiptera: Pentatomidae)

Natural History Distribution. This is a neotropical stink bug, and is found from the southern United States through Central America to Argentina, including many of the Caribbean islands. In the United States, where it has occurred since at least the 1960s (McPherson and McPherson, 2000), it occurs from South Carolina and Florida west to New Mexico, but it is becomingly increasingly important in important soybean-producing states such as Mississippi and Arkansas. In Brazil, where it has long been a serious problem, it seems to be displacing southern green stink bug, Nezara viridula (Linnaeus) as the major soybean pest. Host Plants. Nymphs feed on legumes (Fabaceae) including soybean, green bean, pea, lentil, alfalfa, red clover, and birdsfoot trefoil (Zerbino et  al., 2015). It also occurs on leguminous weeds, including Sesbania, Crotalaria, Cajanus, and Indigofera spp. (McPherson and McPherson, 2000). Panizzi and Slansky (1985b) reported that Indigofera hirsuta Linnaeus and Crotalaria lanceolata E. May (both Fabaceae) were as good as, or better than, crops for supporting growth and reproduction. In fact, soybean is less suitable as a reproductive host than preferred noncrop hosts, but they are of no commercial value so the focus of attention tends to damage the crop plants. Adults are reported to occur on many other plant families (Panizzi and Slansky, 1985a), but it is not always apparent from many reports whether these are reproductive hosts, only adult hosts, of the adults are simply resting on these other plants. Bundy et al. (2018a) provide a list of hosts, and distinguish between reproductive and incidental hosts. Natural Enemies. The natural enemies of this insect present in North America are poorly known. Not surprisingly, Trichopoda pennipes (Fabricius) (Diptera: Tachinidae) can attack this species. In addition, Euthera tentatrix Loew (Diptera: Tachinidae) and Trissolcus basalis Wollaston (Hymenoptera: Platygastridae), Telenomus podisi Ashmead (Hymenoptera: Platygastridae) parasitize redbanded stink bug in North America. Panizzi and

Slansky (1985a) list two (in addition to T. basalis already mentioned) additional parasitoids: Telenomus mormideae Lima (Hymenoptera: Platygastridae) and Trissolcus scuticarinatus Lima (Hymenoptera: Platygastridae) from South America. General predators such as ants (Hymenoptera: Formicidae), grasshoppers (Orthoptera: Acrididae), katydids (Orthoptera: Tettigoniidae), and many predatory Hemiptera feed on redbanded stink bug (Panizzi and Slansky, 1985a). Mermithid nematodes in the genus Hexamermis have been observed to occur naturally in redbanded stink bug, but their incidence is low (Kamminga et al., 2012a,b). Life Cycle and Description. The adults overwinter in sheltered locations, often litter under leaves. They become active early in the spring, usually initially feeding on clover and other early-season legumes and produce eggs. They often disperse to crops after the seedpods begin to form. Females can deposit as many as 10 egg clutches during their 30-day adult life span. The entire life span is 30–60 days, depending on the temperature. Four or more overlapping generations occur in the southern United States annually, with more generations in tropical climates. The abundance of redbanded stink bugs typically increases and attains damaging numbers later in the season (often in August or September). Bundy et al. (2018a) provide a recent review of the biology and management of redbanded stink bug. Egg. Eggs can be found throughout the warm months, a result of the overlapping generations. The cylindrical eggs are 0.88–1.08 mm long and 0.58–0.80 mm in diameter. The covering of the egg is unusually spiny. The egg color is reddish brown with black spines, and a light transverse stripe often occurs medially. The eggs are laid on the end in two parallel rows, though the last egg produced is laid on its side (Bundy and McPherson, 2000). Egg clusters typically consist of about 15 (but up to 28) eggs, and usually are deposited on preferred foods, such as pods, and also on leaves. Oviposition begins on an average (range) of 22.6 (16–65) days after females attain adulthood. Eggs reared at 24°C requires an average of 7.5 days to hatch. Bundy and McPherson (2000) provided a description of eggs, including a key to genera. Nymph. As is normal for stink bugs, the first instars cluster on or around the eggs and egg remnants after hatching, and do not feed. The second and third instars are strongly gregarious, but the fourth and fifth instars disperse. In early instars, the pronotum and head are often black and the abdomen is red with black markings. Later instars are green with black markings and a thick red and black stripe (or stripes) dorsally on the abdomen. The mean (range) duration at 24°C of the instars was reported to be 4.3 (3–5), 5.9 (3–8), 5.5 (3–8), 6.1 (5–8), and 9.7 (7–13) days for instars 1–5, respectively. The total development time from hatching to the adult stage is 31.5 (21–42) days (Panizzi and Smith, 1977).

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Both temperature and photoperiod affect nymphal development time and eventually reproduction (Zerbino et  al., 2013). The greenish nymphs vary in color, especially the fifth instar, which can be rather light or dark. The immature bugs are about 1.3, 2.25, 2.58, 4.6, and 7.87 mm long in instars 1–5, respectively. Adult. The adults are yellowish to light green, but often bear a reddish band across the scutellum, which is the basis for the common name. However, the band on the female is much more variable in color than in the male and may not be red; often it is cream colored, pink, orange, or purple. Photoperiod and reproductive status are correlated with band color; during autumn and winter, the band is light colored and the females are in reproductive diapause (Zerbino et al., 2015). A long spine is found ventrally between the base of the legs. The body length (mean ± SE) is slightly less in males (10.1 ± 0.04 mm) than in females (10.5 ± 0.07 mm) (Zerbino et al., 2015). Depending on the suitability of the host, adult longevity is often 50–90 days. Like most stink bugs, redbanded stink bugs use substrate-borne vibrations for communication. The vi­ bratory signals produced by the stink bugs are mostly ­short-distance female calling, male courtship, and male rivalry songs (Moraes et al., 2005). McPherson and McPherson (2000) provide a key to economically important stink bugs. Also, a key to the most important vegetable-affecting stink bugs is found in Appendix A.

Damage Redbanded stink bug feeds mostly on the seed pods of legumes, although occasionally they feed on the stems and leaves. They secrete saliva during their feeding, which damages the plant tissue, often resulting in tissue death at the site while feeding. Externally, this results in discoloration of the tissue, but internally it causes death of the developing seeds. As a result of this feeding decrease in yield and delay in plant maturity occur. The feeding also results in infection by plant pathogens, especially fungi, but also viral and bacterial pathogens (Husseneder et al., 2016). In the United States, thus far, redbanded stink bug is mostly a soybean pest, but in South America it damages green bean, lentil, and pea crops.

Management Sampling. Stink bug monitoring is often accomplished either by visual examination of plants, by sweeping plants with a net, or by using a vertical beat sheet (Drees and Rice, 1985). Yellow or pyramid traps can also be used to attract and capture stink bugs, although they are more effective if baited with aggregation pheromones (Hogmire and Leskey, 2006; Cottrell and Tillman, 2015).

A specific sex pheromone or aggregation pheromonebased lure for trapping redbanded stink bug has not yet been available, although a male-produced sex pheromone has been identified. It is only attractive to females during the evening hours (Borges et al., 2007). However, P. guildinii responds well to the pheromone lure for Euschistus heros (Fabricius) (Hemiptera: Pentatomidae) in South America, so this lure can also be used to attract and trap redbanded stink bug. Insecticides. Management of redbanded stink bug mostly depends on the application of insecticides. Residual insecticides are needed to kill these stink bugs because not only are they mobile, often moving from uncultivated to cultivated plants, but also because they are inordinately difficult to kill. Soybean growers often apply tank mixes of two insecticides to obtain better control. As the bugs are also found in uncultivated vegetation, application to weedy areas adjacent to crops can be beneficial. Microbial insecticides have the potential for use, with suppression demonstrated using the fungi Metarhizium anisopliae and Beauveria bassiana (Sosa-Gomez and Moscardi, 1998). Cultural Practices. Early planting and harvesting can help the crop to avoid the population increase that typically occurs late in the summer. Suppression of weedy legumes can help to suppress the population increase in the spring.

Redshouldered Stink Bug

Thyanta spp. (Hemiptera: Pentatomidae)

Natural History Distribution. This genus is poorly known, and authorities debate the status and distribution of these insects. The important “species” seem to be Thyanta accerra McAtee, T. calceata (Say), T. custator (Fabricius), and T. pallidovirens (Stål), but they also have been regarded as subspecies rather than species (McPherson, 1982; Rider and Chapin, 1992; McPherson and McPherson, 2000). Thus, the regional occurrence and damage caused by the different species is confusing. Presently, most authorities consider T. custator (including what was formerly considered T. accerra and T. calceata) to be a transcontinental species and T. pallidovirens to be limited in distribution to west of the Rocky Mountains. Differences in male-produced pheromones and substrate-borne vibrational signals between T. custator and T. pallidovirens seem to justify separation of at least these two species. Host Plants. A number of crops, weeds, and native plants are hosts of Thyanta spp. Among vegetables damaged are bean, lima bean, eggplant, cowpea, corn, asparagus, pea, lentil, and especially tomato. Other crops affected are alfalfa, buckwheat, canola (rape), cotton, sorghum, soybean, sugar beets, oats, clover, timothy, wheat, and pistachio.

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Often vegetables are damaged when the stink bugs disperse away from another crop that is more favorable for their development; their relocation typically is due to senescence or harvest. Weeds can also function as an initial host that supplies insects to vegetable crops. Some common weeds that host Thyanta spp. are pokeweed, Phytolacca americana; mullein, Verbascum thapsus; ragweed, Ambrosia sp.; milkweed, Asclepias sp.; goldenrood, Solidago sp.; bigpod sesbania, Sesbania emerus; wild blackberry, Rubus sp.; rabbitbrush, Ericameria nauseosa; and coralberry, Symphoricarpos orbiculatus. Natural Enemies. As with other stink bugs, the important natural enemies tend to be egg parasites of the family Platygastridae such as Telenomus podisi Ashmead and Trissolcus thyantae Ashmead and tachinid flies (Tachinidae) that attack the nymphs and adults, such as Cylindromyia fumipennis (Bigot), Gymnoclytia occidua (Walker), and Euclytia flava (Townsend). Life Cycle and Description. This species complex is not well studied. Eggs are described by Esselbaugh (1946), nymphs by Decoursey and Esselbaugh (1962), and some of the biology is given by Esselbaugh (1948). It is also summarized and expanded by McPherson (1982). There are two 2 generations per year. A male-produced pheromone of T. pallidovirens that attracts females has been described (McBrien et  al., 2002b). Substrate-borne vibrational signals are also important in communication of this species (McBrien et al., 2002a). Egg. The barrel-shaped eggs from this group vary somewhat, measuring from 0.88 to 1.5 mm in length and 0.76 to 1.25 mm in diameter, depending on what “species” is considered. They are yellowish or yellow green in color, darkening with age. The upper end is ringed with 45–65 processes. Egg clusters are deposited on fruit or foliage. Esselbaugh (1946) reported (in T. custator) that egg number per cluster was up to 70 eggs but with a mean of about 35. Schotzko and O’Keeffe (1989b), on the other hand, reported (in T. pallidovirens) a mean of about 24 eggs per cluster on pea and lentil pod-bearing plants. Egg production totaled up to 300 eggs per female when fed pea pod-bearing plants but was only about 100 when fed lentil pod-bearing plants. Seedling plants and plants in flower were not suitable hosts for reproduction. In general, egg clusters produced initially are the largest, and egg gradually diminishes in number as the female ages. As with other stink bugs, the eggs are placed on end, and in several definite rows. They are not tightly interlocked, however, unlike many species. Esselbaugh (1948) indicated that the egg required an average of 4.5  days to hatch, within a range of 3–8 days. A description of eggs, including a key to the genera of stink bugs, is found in Bundy and McPherson (2000). The eggs of Thyanta custator accerra McAtee, as well as eggs of several other stink bugs, are described by Bundy and McPherson (2000).

Nymph. Mean durations (range) of the nymphal instars given by Esselbaugh (1948) were 3.8, 6.9, 8.3, 9.8, and 12.0  days for instars 1–5, respectively. Thus, duration of the nymphal stage averaged 40.8 days. The range in length of the nymphs is 0.88–1.06, 1.16–1.84, 1.80–3.20, 3.10– 4.60, and 4.40–8.20 mm for instars 1–5 (Decoursey and Esselbaugh, 1962). The shape is elliptical or oval and they are usually brownish or bronze in color.

FIG. 8.57  Redshouldered stink bug. (Photo by L. Buss.)

Adult. The adult dorsally is generally green in the summer months, although often brown in the autumn and spring. Ventrally, this bug is yellow green. The anterolateral pronotal margins sometimes are black. As suggested by the name “redshouldered,” there are varying amounts of red (often faint) on the pronotum between the humeral angles. Sometimes the red coloration extends to the scutellum. They measure about 9–13 mm in length.

Damage Feeding by Thyanta spp. on fruits and pods causes depressions or pits in the growing fruit, and a lighter coloration or mottling of the appearance. The cells beneath the feeding site apparently are destroyed by the enzymes secreted by the bug. Thyanta pallidovirens is especially damaging to tomato fruit in California, and also can be quite damaging to pistachio fruit. In California, the abundance of T. pallidovirens and Euschistus conspersus Uhler, and their damage to tomatoes, were found to be greater near weedy borders (Pease and Zalom, 2010).

Management Management of this species seems to be little different from other stink bug species. Population monitoring is usually accomplished by visual examination or sweep netting. Dispersal from crop to crop is expected as plants mature and become attractive to these fruit and seed-feeding bugs. Weeds are an important source of dispersing bugs,

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and weed management can alleviate damage (Pease and Zalom, 2010). Insecticides are commonly applied, especially to address nymphal populations, although stink bugs are difficult to kill.

Say Stink Bug

Chlorochroa sayi (Stål)

Uhler Stink Bug

Chlorochroa uhleri (Stål) (Hemiptera: Pentatomidae)

Natural History Distribution. These stink bugs are native to western North America. Say stink bug is reported throughout the western United States from Montana and western Texas west to Oregon and California. Chlorochoa sayi does not occur in Canada (Scudder and Thomas, 1987), and reports of its occurrence in Canada are based on misidentification. Uhler stink bug is found from the Saskatchewan and the Dakotas, Nebraska, and New Mexico west to the Pacific Ocean. Host Plants. Say and Uhler stink bugs feed on the fruit and seeds of many plants. They are known principally as a pest of grains, and prefer to attack the seed head of such crops as alfalfa, barley, oat, rye, and wheat. However, on occasion they damage such vegetables as asparagus, bean, cabbage, lettuce, pea, and tomato. They also feed extensively on such weeds as broom snakeweed, Gutierrezia spp.; lambsquarters, Chenopodium album; mallow, Malva spp.; musk thistle, Carduus nutans; pigweed, Amaranthus spp.; prickly pear cactus, Opuntia sp.; Russian thistle, Salsola kali; tansymustard, Descurainia pinnata; toadflax, Linaria vulgaris; tumble mustard, Sisymbrium attissimum; sage, Artemisia spp; saltbush, Atriplex spp.; and feathergrass, Stipa spp. A related species, C. ligata (Say), is known as the “conchuela.” It is also found in western North America, and is associated with some of the same host plants (McPherson and McPherson, 2000). However, it is rarely reported to be a pest. Natural Enemies. Several parasitoids are known. An egg parasitoid, Telenomus utahensis Ashmead (Hymenoptera: Platygastridae), is an important mortality factor of Say stink bug, sometimes causing 60% mortality or greater late in the season (Jubb and Watson, 1971a,b). When this wasp discovers an egg clutch, few if any eggs escape parasitism. Several generations of the parasitoid occur annually (Caffrey and Barber, 1919). The other known wasp parasitoids, Telenomus podisi Ashmead (Hymenoptera: Platygastridae) and Ooencyrtus johnsoni (Howard) (Hymenoptera: Encyrtidae), also attack eggs. Among fly species reared from Say stink bug are Cylinromyia armata Aldrich, C. euchenor (Walker), and

Gymnosoma fuliginosum Robineau-Desvoidy (all Diptera: Tachinidae). Although the Cylinromyia spp. seem to be of little importance, the latter fly species attacks up to 25% of late instar nymphs and adults. Parasitoids of Uhler stink bug are less well known, though Telenomus utahensis caused higher levels of parasitism in Uhler stink bug than in Say stink bug (Jubb and Watson, 1971b), and G. fuliginosum is also known from C. uhleri. General predators, including the assassin bug Sinea spinipes (F.) (Hemiptera: Reduviidae), and the ambush bug, Phymata erosa Stål (Hemiptera: Phymatidae), feed on nymphs. The soft-winged flower beetle, Collops bipunctatus Say (Coleoptera: Melyridae), feeds on stink bug eggs. Songbirds and lizards sometimes consume both Say stink bug and Uhler stink bug (Knowlton, 1944; Knowlton et al., 1946) despite the fact that some vertebrate predators learn to avoid these odor-producing and presumably distasteful insects. Life Cycle and Description. Three or four generations of Say stink bug are known from New Mexico. The overwintering adults deposit eggs in late April or May, with firstgeneration adults appearing in June, second in August, and third in September. A small fourth generation sometimes occurs, though many nymphs from this generation perish with the onset of cold weather. The adults from generations 2–4 enter diapause during late October or early November, and reemerge the following spring. A complete generation requires about 80 days. Uhler stink bug’s biology and morphology are largely undescribed, but presumably about the same as Say stink bug. Egg. The eggs of Say stink bug are barrel shaped, and deposited on end. The height is 1.1–1.2 mm and diameter about 0.6–0.9 mm. When viewed from above, the egg is marked with three white circles which alternate with gray. On one side of each egg is an irregular patch of gray. Mean duration of the egg stage is about 9 days (range 4–13 days), though during the warmest months egg hatch tends to occur in 5–7 days. Eggs darken in color immediately preceding hatch. Egg clusters are placed on the underside of some object. Overwintering adults may deposit eggs on the dead plants or rubbish comprising their overwintering quarters, but later generations tend to oviposit on green plants. They often are placed in parallel rows, usually two or four rows, with rows consisting of 7–20 eggs. This tendency is most pronounced when oviposition occurs on the narrow stems of grasses which constrain oviposition, whereas on broad surfaces the eggs may consist of several rows. The number of eggs per cluster averages about 26 (range 13–43 eggs). Nymph. There are five instars in Say stink bug; the mean (range) duration is 5.0 (5), 8.2 (7–9), 6.4 (5–7), 8.2 (7–10), and 14.9 (13–17) days, respectively, during the summer months. Young nymphs usually remain on or near the egg mass for the duration of the first instar. The first

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instar ­measures 1.1–1.5 mm long. Its color is predominantly black, though a protruding area at the lateral margins of the thorax and abdomen bears some yellow pigment, and there are yellow-white spots on the abdomen. During the second instar, nymphs wander away from the egg mass to feed, but generally they remain fairly aggregated. The second instar measures 2.2–2.5 mm long and is colored like the first instar, except for also bearing yellow-white spots on the thorax. The later instars are solitary and move freely about the plant to feed. The nymphs feed during the morning and late afternoon, seeking shelter during the heat of the day and cool of the night. They also seek shelter during periods of high wind and rainfall. The third instar measures 3.1–3.2 mm long. The anterior region of the bug is black, but fading to dark green toward the posterior. The lateral margins turn from white to orange during this instar, and some yellow spots remain on the abdomen. The fourth and fifth instars measure 5.2–6.6, and 8.9–10.6 mm, respectively. They are pale green, with the lateral margins white with an orange edge, and some yellow spots on the abdomen. The wing pads become evident during the final two instars, especially the latter.

is estimated to average 54 per female, with a maximum of 107 eggs. This seems to be a fairly small number of eggs, however, and may represent the adverse consequences of caging. Overwintering adults and first-generation nymphs feed principally on early-developing native plants. The second-generation adults feed extensively on grain, and cause considerable damage. Later generations also feed on native plants because grains have matured and have been harvested. Unlike the nymphs, which seek shelter during the heat of the day, adults are active throughout the daylight hours. The adults are long lived, reportedly persisting for 3–4  months in the summer. During the winter, of course, adults must survive for several months. Adults are often found overwintering under plant debris, bales of hay, and dried livestock droppings. As with many pentatomids, chemical and vibrational communication among the bugs is important in their biology. Aggregation pheromones tend to be more important than sex pheromones, but defensive chemicals and substrate-borne vibrational signals are also important (Millar et al., 2010; Bagwell et al., 2008; Ho and Millar, 2001). The most complete research of Say stink bug biology, conducted in New Mexico, was published by Caffrey and Barber (1919). Patton and Mail (1935) contributed some observations from Montana. Buxton et al. (1983) described both species, and provided a key to close relatives, as does McPherson and McPherson (2000). A key to distinguish stink bugs commonly affecting vegetables is also found in Appendix A.

Damage

FIG. 8.58  Say stink bug. (Photo by J. Capinera.)

Adult. The adults of Say stink bug are about 12–16 mm long. Their color is green, but the shade varies considerably. The lateral borders of the pronotum, three spots on the anterior border of the scutellum, and the apex of the scutellum are yellow, orange, or red. The apical portion of the front wings is marked with small purple flecks along the veins. The adult of Uhler stink bug greatly resembles Say stink bug, though it lacks purple flecks on the membranous portion of the front wings and usually lacks the orange color common at the tip of the scutellum in Say stink bug. Newly emerged adults are unable to oviposit immediately. A period of 20–30 days normally is required before adults mate and begin to deposit eggs. Females deposit eggs over a period of about 1 month. Total fecundity

Nymphs and adults prefer to suck liquids from seeds and fruits, but if these are not available they feed readily on young leaf and stem tissue. Once seeds, including grain, begin to harden, the bugs no longer are able to feed. Thus, rapidly growing seeds are preferred. Seed heads of grains that have been attacked acquire a dull yellowish-white color, in contrast to the green appearance of undamaged heads. Such seeds are hollow, or nearly so. When fruit is attacked, the tissue adjacent to the feeding puncture does not develop as the fruit grows, leaving a blemish in the form of a depression. Foliar tissue that has been fed upon becomes wilted or discolored, and often dies.

Management Sampling. Monitoring typically consists of visual inspection, often accompanied by collection using a sweep net or beating tray to facilitate collection. However, the principal component of an aggregation pheromone produced by males of both species will attract males and females of both species, offering an alternative and more selective method of population estimation (Millar et al., 2010). Substrate-borne vibrational signals are also important in the communication

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of these stink bugs (Bagwell et al., 2008), but thus far has not been used for population monitoring. Insecticides. In the past, broad spectrum chemical insecticides targeted at more important pests also suppressed stink bugs, reducing them to minor pest status. The availability of more selective control techniques, including more specific insecticides, has allowed stink bugs to become more important pests because they are often not affected by these newer products. Thus, insecticides and cultural control aimed directly at stink bugs have assumed greater importance. Cultural Controls. These stink bugs are dependent on weeds for development of the first generations of the season. Thus, suppression of weeds is a recommended management practice to suppress stink bugs. However, with the immense amount of rangeland supporting suitable food plants in the western states, it is improbable that weed management would be effective in many areas. Many growers, therefore, solely depend on the application of foliar insecticides for stink bug control. It is useful, however, to monitor weedy areas and field margins for stink bugs, because they are infrequently abundant enough to damage vegetable crops.

Southern Green Stink Bug

Nezara viridula (Linnaeus) (Hemiptera: Pentatomidae)

Natural History Distribution. Southern green stink bug has a worldwide distribution. It is found on all continents where agriculture is practiced, but absent or rare from regions with cold winters. Its origin is probably eastern Africa, and was first observed in the western hemisphere in 1798. Distribution in North America is limited primarily to the southeastern United States—Virginia to Florida in the east, to Ohio and Arkansas in the midwest, and to Texas in the southwest. However, it has become established in Hawaii (in 1961) and California (in 1986), and occasional specimens have been found elsewhere outside the generally infested southeast. Southern green stink bug is a strong flier, and its range is expanding in many parts of the world. Host Plants. The host range of this insect includes over 30 families of plants, though it shows a preference for legumes and crucifers. Preference among plants varies during the year, with this stink bug most attracted to plants that are producing pods or fruit. As plants senesce, the bugs move to more succulent hosts. Among vegetables attacked by southern green stink bug are artichoke, bean, Brussels sprouts, cabbage, cauliflower, collards, corn, cowpea, cucumber, eggplant, mustard, okra, pea, pepper, potato, radish, squash, sweet potato, tomato, and turnip. Outside of North America, this insect is most commonly known as

“green vegetable bug,” an indication of its most frequent host selection. However, it feeds readily on other crops, and is known to attack field crops such as corn, clover, cotton, peanut, soybean, sugarcane, rice, and tobacco as well as fruits such as blackberry, grapefruit, lime, mulberry, orange, and peach. In many tropical areas, this bug is considered as a limiting factor in soybean production. Weeds commonly serve as hosts. Some of the wild plants fed upon by this bug are beggarweed, Desmodium tortuosum; castor bean, Ricinus communis; dock, Rumex sp.; nutgrass, Cyperus esculentus; lambsquarters, Chenopodium album; passion flower, Passiflora incarinata; pigweed, Amaranthus spp.; rattlebox, Crotalaria usaramoensis; wild grape, Vitus sp.; and wild plum, Prunus sp. Esquivel et al. (2018) provide an updated list of host plants. Southern green stink bug, a long lived and strong flier, moves readily among host plants. Some of the host associations are not true expressions of preference or suitability, but reflect availability. Also, host plants that are suitable for one stage may be unsuitable for another. Velasco and Walter (1992), for example, showed that castor bean was good for adult survival and egg production, but poor for nymphal survival. Corn also favors adult survival, but inhibits reproduction. Wild crucifers are not optimal hosts, but early in the season they are abundant and are the best hosts available, so they favor population increase. Soybean favors survival of both nymphs and adults, but is not especially attractive to these insects. Natural Enemies. As might be expected of an insect with a worldwide distribution, numerous parasitoids and predators are known. Over 50 species of parasitoids are known, most as egg parasitoids, and most not specific to southern green stink bug (Jones, 1918a). The most common parasitoids in North America tend to be the egg parasitoid Trissolcus basalis (Wollaston) (Hymenoptera: Platygastridae) and the nymphal and adult parasitoid Trichopoda pennipes (Fabricius) (Diptera: Tachinidae). Trichopoda parasitizes high proportions of green stink bug populations on several crops, with the proportion parasitized increasing through the growing season (Todd and Lewis, 1976; Buschman and Whitcomb, 1980; McLain et al., 1990). This insect is thought by some to be a complex of cryptic species, however, so the ecological relationship is uncertain. Trichopoda spp. were introduced into Hawaii and were credited with providing effective biological control on most crops. Other North American parasitoids include Anastatus sp. (Hymenoptera: Eupelmidae), Ooencyrtus sp. (Hymenoptera: Encyrtidae), Telenomus spp. (Hymenoptera: Platygastridae), and others; Jones et al. (1996) provided a list of parasitoids. Hoffman et  al. (1991b) reported on the successful introduction of Trissolcus basalis (Wollaston) (Hymenoptera: Platygastridae) to California to aid in the suppression of southern green stink bug.

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Predators of southern green stink bug are numerous, and Stam et al. (1987) gave a detailed assessment of predation in Louisiana soybean. Common egg predators were red imported fire ant, Solenopsis invicta Buren (Hymenoptera: Formicidae), grasshoppers (Orthoptera: Acrididae and Tettigoniidae), and southern green stink bug nymphs. Common nymphal predators were big-eyed bug, Geocoris spp. (Hemiptera: Lygaeidae); damsel bug, Reduviolus roseipennis Reuter (Hemiptera: Nabidae); and other insects in the orders Hemiptera and Coleoptera. Spiders sometimes caused significant predation. Predators, including birds, were also discussed by Drake (1920). Ehler (2002) conducted a detailed assessment of N. viridula biological suppression on tomatoes and bean in California. In this study, predation of eggs was less than predation of young nymphs, and parasitism was more important than predation. The relative importance of egg predators and parsitoids was examined by Shepard et al. (1994) in South Carolina. Stink bug egg clusters were introduced into several crops and their fate determined. The principal egg parasitoid was Trissolcus basalis, accounting for about 95% of parasitism. Predation (disappearance of eggs) and parasitism varied in relative importance among crops, seasons, and years; on average, predation and parasitism were about equally important. The action of natural enemies sometimes resulted in complete destruction of egg clusters. Diseases of this stink bug are poorly known. However, viruses affecting southern green stink bug were found in South Africa (Williamson and von Wechmar, 1995). Life Cycle and Description. The southern green stink bug overwinters in the adult stage, and enters diapause when the length of photophase falls to 12–13 h. The number of generations annually is estimated at 4–5, with considerable overlapping late in the season. About 45  days are required for a complete life cycle during the summer months. Todd and Herzog (1980) indicated the following “typical” pattern of stink bug phenology in the southeastern United States: inactive adults overwinter until February or March, and when they become active they feed on crucifers and small grains; first-generation nymphs and adults are present in March and April and feed on clover; second and third are present from May through July and feed on tobacco, corn, and vegetables, especially tomato; third-generation adults disperse to soybean, where fourth-generation nymphs and adults and fifth-generation nymphs feed and develop until October or November; fifth-generation adults disperse to crucifers or other late season hosts, and overwinter. With the onset of warm days in the spring, overwintered adults begin to copulate. The process is protracted, often lasting more than a day, and multiple matings are commonplace. Following 2–3 weeks of feeding the egg production commences. Esquivel et al. (2018) provide a recent review of this species, including both biology and management.

Egg. The eggs are laid in clusters, generally on the lower surface of foliage, and with about 60 eggs (range 30–130 eggs) per cluster. Females normally produce 1–3 egg clusters, with total production ranging from 45 eggs when adults are provided only with relatively poor hosts, to 160 eggs when provided with good host plants. The eggs are deposited in regularly shaped, hexagonal clusters, with the individual eggs ordered in regular rows and glued together. They are affixed to the plant on end, with the top or visible end somewhat flattened, and the bottom end rounded. The eggs measure about 1.3 mm long and 0.9 mm wide. They are yellowish white to pinkish yellow and the top, or cap, is clearly indicated by a ring and 28–32 min spines. The eggs darken in color as incubation proceeds, and hatching occurs after about 5  days. Hatching occurs synchronously in the egg cluster, so within 1.0–1.5 h all are hatched. Nymph. There are five instars. At hatching the nymph is yellowish orange, but soon it becomes brown. Each segment of the abdomen is marked, both dorsally and ventrally, with a pair of light spots. The appendages are yellow. The body length is about 1.6 mm. During most of this first instar, the nymphs cluster tightly around the egg cluster and do not feed. Finally, just before the young insects are about to molt, the nymphs move from the cluster and commence feeding. In the second instar the head and thorax are black, with the abdomen reddish black. Both the thorax and abdomen are marked with yellowish spots. The appendages are black. The body length is about 3.2 mm. Second instars remain near the egg cluster, and remain aggregated. The third instar is very similar to the second, differing principally in size, and measuring about 3.6 mm long. Third instars disperse away from the egg cluster, but remain aggregated. The fourth instar also may be relatively unchanged in appearance from the preceding instars, differing principally in body length of about 4.2 mm. However, the color of the fourth instar is highly variable, and may instead be greenish; the thorax is light green with black markings, and the abdomen is darker green with salmon shading and white spots. The appendages are brownish, with the tip of the antennae greenish. Unlike the preceding instars, fourth and fifth instars do not aggregate. The fifth instar is also highly variable, the head, thorax, and wing pads ranging from light green to almost black. The abdomen tends to be colored light or dark, corresponding to the shade of the thorax, and marked with rose spots dorsally, and whitish spots laterally. The body length of fifth instars is about 10 mm. Mean duration (range) of instars 1–5 is about 4 (3–5), 5 (3–10), 5 (3–9), 8 (6–10), and 10 (7–13) days, respectively. Thus, total nymphal development time is about 32 days, and egg to adult development requires 35–37 days, depending on the temperature and suitability of food. The optimal temperature for the development is about 30°C.

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FIG. 8.59  Southern green stink bug. (Photo by L. Buss.)

Adult. The adult is generally uniform light green, both dorsally and ventrally, though the ventral surface is paler. Adults measure about 13–17 mm long and 8 mm wide. The thorax is the widest part of the body, and is rounded laterally. Sometimes the adults differ in having yellow or brown on the thorax or scutellum, and the brown color may extend onto the forewings. During the summer months, females begin egg production about 14–20 days after attaining the adult stage. Southern green stink bug, N. viridula, is easily confused with green stink bug, Chinavia hilaris (Say). However, the two species differ ecologically and morphologically. Chinavia hilaris is usually associated with trees and shrubs, rather than the herbaceous vegetation fed upon by N. viridula. Also, though C. hilaris is found widely in North America, it is most abundant in the north. The two species can be differentiated by the shape of the abdominal spine. When viewed from below, C. hilaris has a pointed spine protruding forward between the base of the hind legs, whereas in N. viridula the spine is rounded. Useful keys to Pentatomidae that include southern green stink bug include McPherson and McPherson (2000) and McPherson (1982). A key to distinguish stink bugs commonly affecting vegetables is found in Appendix A. In coastal areas of the south, the adults of southern green stink bug are scarce during the winter months but not all are dormant; a few being found feeding on succulent plants. Even in these relatively warm areas, however, no nymphs are found. Overwintering occurs in sheltered locations, most commonly under bark of trees, but also beneath fallen leaves and in Spanish moss. Considerable winter mortality among southern green stink bug can occur at the northern limits of its range, and winter weather is considered to be an important variable in population abundance. High levels of mortality occur when bugs are exposed to − 10°C and − 5°C for 3 and 55 h, respectively (Elsey, 1993). Southern green stink bug displays a complex chemical ecology. Nymphs produce a bifunctional pheromone

that functions as an aggregation pheromone at low concentrations and a dispersant at high concentrations. Adult males produce an aggregation pheromone that attracts females, other males, and the parasitoid Trichopoda pennipes (Fabricius) (Diptera: Tachinidae). Presumably the principal function of this chemical is to enhance mate finding. Not surprisingly with an insect that occurs so widely, the aggregation pheromone blend varies slightly from location to location (Aldrich et al., 1987). However, there also are visual, tactile, and acoustic stimuli that are a necessary prelude to mating (Harris et  al., 1982b). The substrate-borne vibrational signals differ between sexes and elicit varying types of behavior (Cokl et  al., 2001). Stink bugs also secrete a defensive chemical that apparently is repellent to predators. Descriptions and summaries of life history were provided by Jones (1918a) and Drake (1920), the developmental biology was given by Harris and Todd (1980), and a useful review of behavior and ecology was presented by Todd (1989). Information on stink bug culture was provided by Harris and Todd (1981) and Brewer and Jones (1985).

Damage Southern green stink bug punctures plant tissue with its piercing-sucking mouthparts and removes sap, often feeding at night. Stem, leaf, blossom, and fruit tissue may all be attacked, but fruiting structures are preferred and invariably the tissue selected is young and succulent. The feeding site, as it heals, becomes hard and darkened. Seeds may be shriveled, deformed, and shrunken, or may simply bear a dark mark and depression at the feeding site. Fruit may be deformed or dropped from the plant. Under high levels of feeding pressure, young plants and plant shoots may perish. Stink bugs are capable of introducing bacteria and yeasts into plants as they feed. Wounds also allow entry of secondary plant pathogens. Vegetable crops are easily damaged by southern green stink bug. Studies of tomato fruit attack in Louisiana indicated that small fruit was preferred over large fruit, and green over red fruit (Lye and Story, 1988). There was an inverse linear relationship between bug density and fruit quality; fewer than two bugs on a tomato fruit for 1 day, or one bug for 2 days, resulted in decrease in fruit quality (Lye et al., 1988a,b). In corn grown in Louisiana, as few as two bugs per ear of corn during early ear development could result in damage (Negron and Riley, 1987). Cowpea and lima bean were studied in Georgia, and 2 days of feeding by one bug on a pod significantly reduced pod size and mean seed weight (Nilakhe et al., 1981a,b). In South Carolina, feeding by N. viridula on cowpea was more damaging to blossoms and pods when the plant was attacked early in the development of these plant organs, relative to later in development (Schalk and Fery, 1982; Abudulai and Shepard, 2001). It also can vector the fungal disease Eremothecium coryli (formerly Nematospora coryli).

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Management Sampling. A comprehensive review of stink bug sampling was published by Todd and Herzog (1980), though soybean rather than vegetables is emphasized. Sweep nets and direct visual examination are techniques most useful for stinkbug sampling. Lye and Story (1989) studied southern green stink bug sampling in Louisiana tomatoes. They reported that bugs were moderately aggregated and that young fruit clusters were the most efficient sampling unit. They also developed a sequential sampling protocol. Insecticides. Insecticides are commonly applied to foliage, especially at blossoming and pod set, to protect susceptible crops from stink bug damage. Systemic insecticides applied to the soil tend to be less effective (Chalfant, 1973a). Insecticide resistance has become a problem in some areas of the world, so steps to minimize the development of resistant populations should be followed. Insecticides can interfere with naturally occurring biological control, but research has shown that insecticides with a short residual life have a minimal impact on egg parasitoids (Orr et al., 1989). The important nymphal and adult parasitoid Trichopoda pennipes (F.) (Diptera: Tachinidae) is more susceptible to many insecticides than is N. viridula, necessitating caution in the application of insecticides if biological control is to be maximized (Tillman, 2006a,b). Cultural Techniques. Consideration of early-season feeding behavior is an important component of management. Cruciferous crops and weeds are potential sources of stink bugs for later legume crops, and should be monitored, treated, or destroyed. Legume cover crops or forage crops are another source of bugs that might invade vegetables, and should be treated similarly. Farmers and entomologists have long attempted to use early-season crops, or late-season crops planted early, as trap crops to lure stink bugs from the principal crop. Early in the spring or late in the autumn cruciferous plants such as collards, mustard, radish, rape, and turnip are attractive to bugs, whereas late in the spring and during the summer legumes such as bean and cowpea are most attractive. Drake (1920) recommended radish and collards in the spring and rattlebox in the summer to protect tomato. Similarly, Rea et al. (2002), working in New Zealand, showed that black mustard could attract and hold N. viridula for 2 weeks, allowing sweet corn to mature without damage. McPherson and Newsom (1984) and Todd and Schumann (1988) reported that early-planted soybean or cowpea could serve as a trap crop for late-planted soybean. Sorghum is also highly attractive to N. viridula (Tillman, 2006a,b; Gordon et al., 2017), as is sunflower, and sunflower is highly attractive to pollinators and natural enemies. Stink bugs will disperse from trap crops, however, so usually they must be treated with insecticide before dispersal occurs in the adult stage.

Plant cultivars resistant to attack by stink bug generally are not available commercially, but genetic sources of resistance have been identified for cowpea (Schalk and Fery, 1986).

Tomato Stink Bug

Arvelius albopunctatus (DeGeer) (Hemiptera: Pentatomidae)

Natural History Distribution. This neotropical insect is found widely in the Western Hemisphere from Argentina north to the United States and including most of the Caribbean region. In the United States, its distribution is presently limited to southern California, Arizona, and Florida. It has increased in importance in Brazil since the 1990s. Similarly, although problems with this insect are not widespread, Panizzi (2015) notes that it is becoming more important on tomatoes in Baja California, Mexico. Host Plants. Tomato stink bug is primarily a pest of tomatoes, though it feeds on other plants in the plant family Solanaceae including potato, pepper, and eggplant. Not surprisingly, weeds in this family such as Solanum capsicoides All., S. bonariense L., S. gracile Sendtn., S. palinacanthum Dunal, S. viarum Dunal, and Datura sp. are also susceptible to infestation. Other crops are sometimes mentioned as hosts, but these need confirmation. The status of this insect is controversial. In most areas, it is of secondary importance, and has even been viewed as a beneficial predator. Natural Enemies. The natural enemies are poorly known, though it is parasitized by Trichopoda pennipes (Fabricius) (Diptera: Tachinidae) and Hexacladia smithii Ashmead (Hymenoptera: Encyrtidae). Life Cycle and Description. Duration of the life cycle is about 80 days in Argentina. Egg. The female deposits her eggs on the plant. The eggs are light yellow, and barrel shaped. Martínez and Folcia (1999) reported that the mean number of eggs (± SD) per cluster was 28.2 ± 13.9, and duration of the egg stage was 16.9 ± 1.7 days when reared at a daytime temperature of 25°C in the daytime but falling to 14°C at night. Nymph. The nymphs are oval in shape, and highly varies in general coloration, generally yellowish but also including green, orange, and red (Campos et  al., 2007). The lateral margin of the abdominal segments is marked with black spots, and except for the first instar there are three large black spots medially on the abdomen. There are five instars. The mean, standard deviation, and range for nymphal development times were reported by Martínez and Folcia (1999) to be 5.18, 0.96, 4–6 days for instar 1; 13.84, 3.35, 8–26 days for instar 2; 11.70, 2.20, 8–19 days for ­instar 3;

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12.74, 3.13, 8–19  days for instar 4; and 18.63, 3.42, ­14–24 days for instar 5. Grazia et al. (1984) reported that the mean and range of body lengths for the nymphs were 1.86 and 1.76–1.97 mm for instar 1; 2.45 and 1.99–2.91 mm for instar 2; 4.20 and 3.42–4.98 mm for instar 3; 6.21 and 5.22–7.20 mm for instar 4; and 9.50 and 8.36–10.64 mm for instar 5. As with other stink bugs, the first instars remain aggregated after hatching, then dispersing as second instars.

and placement of susceptible crops can be used to reduce the level of infestation. For small plantings, screening can be used to keep these large bugs from reaching plants and fruits.

FAMILY PSEUDOCOCCIDAE—MEALYBUGS Cotton Mealybug

Phenacoccus solenopsis Tinsley (Hemiptera: Pseudococcidae)

Natural History

FIG. 8.60  Tomato stink bug. (Photo by L. Buss.)

Adult. The adult is yellow to green, like many stink bugs. However, it has very pronounced spines (usually ­yellow-tipped) laterally on the pronotum. In addition, there are numerous small dark spots on the pronotum and scutellum and white or spots on the hemelytra. The antennae have alternating light- and dark-colored bands, usually green and white. The head is usually edged with black. The adults are 11–12 mm long. McPherson and McPherson (2000) provide a key to economically important stink bugs, although it does not include A. albopunctatus. Also, a key to the most important vegetable-affecting stink bugs (including A. albopunctatus) is found in Appendix A.

Damage Damage results from feeding by nymphs and adults feeding on the fruit. Depression form at the points of feeding, resulting in the formation of fruit of inferior quality. Feeding sites are also points of entry of infections by plant pathogens, though they are not known to be vectors.

Management Insecticides. Contact and systemic insecticides are applied to the foliage as necessary, preferably when the insects are still young. Cultural Practices. Growers principally depend on insecticides for control, but populations can be minimized by elimination of solanaceous weeds, and selective timing

Distribution. Cotton mealybug (also frequently called solenopsis mealybug) was first described from the United States, but has now spread throughout Central and South America, much of Africa, southern Europe, the Middle East and southern Asia, and recently it was introduced to Australia. Most of the spread has occurred in the last two or three decades, so this is a “new” pest for much of the world. It has proved to be most damaging under tropical and subtropical conditions. It is said to be favored by hot, dry conditions, whereas high humidity and rainfall are deleterious. Thus, it has caused injury mostly in Pakistan, India, and nearby countries. Except in greenhouses, it does not normally survive freezing temperatures. Host Plants. Like many mealybugs, cotton mealybug displays a wide host range, and is documented to feed on over 200 host plants. Among the vegetable crops affected are okra, tomato, tomatillo, eggplant, pepper, chilli, potato, sweetpotato, watermelon, honeydew melon, squash, pumpkin, gourd, and corn. Other crops include cotton, fig, guava, grape, pigeon pea, sunflower, bottle mango, papaya, pomegranate, tobacco, basil, cashew, and castor bean. Numerous other plants, including weeds and ornamental plants, are susceptible to injury, though cotton grown under arid conditions is the crop usually identified as most seriously injured. Among the weeds supporting cotton mealybug are purslane, Portulaca oleracea (Portulacaceae); Canadian fleabane, Conyza canadensis (Asteraceae); bindweed, Convolvulus arvensis (Convolvulaceae); lambsquarters, Chenopodium album (Amaranthaceae); dandelion, Taraxacum officinale (Asteraceae), and common ragweed, Ambrosia artemisiifolia (Asteraceae). In general, this mealybug displays a preference for Asteraceae, Euphorbiaceae, Fabaceae, Malvaceae, and Solanaceae. Kosztarab (1996) reports that it is found widely on grasses and ragweed in eastern North America. Cotton is usually considered to be a favored host plant, but cotton mealybug is usually an induced pest that survives the heavy insecticide usage often found in cotton fields, and is unaffected by Bt cotton, so the relationship of this mealybug with cotton is complex.

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Badshah et al. (2015) determined the growth rate and fecundity of cotton mealybug reared on four vegetable crops (okra, eggplant, potato, and tomato) and compared them to a favored host, Hibiscus rosa-sinensis. The mealybugs completed their development fastest on Hibiscus (17.4  days) and slowest on tomato (19.6  days), though development times were quite similar for all the vegetable crops. Weight gain in females and egg production followed the same pattern as development time, with maximum weight and egg production associated with Hibiscus and minimum weight and egg production on tomato. Nevertheless, all plants demonstrated suitability and potential for considerable population increase. Natural Enemies. Natural enemies can play an important role in suppressing cotton mealybug populations. Several wasp parasitoids (Hymenoptera), ladybird beetles (Coleoptera: Coccinellidae), and lacewings (Neuroptera: Chrysopidae) attack these insects, although the species vary regionally. A ladybird beetle known as the mealybug destroyer, Cryptolaemus montrouzieri Mulsant is sometimes released to aid in mealybug population reduction. Ants interfere with the activity of beneficial insects. The benefits associated with parasitoids in India were documented by Ram and Saini (2010). Fand et al. (2010) published a comparison of four ladybird predators associated with cotton mealybug in India. Rashid et al. (2012) compared predation of cotton mealybug by the lacewing Chrysoperla carnea (Stehens) (Neuroptera: Chrysopidae) and the ladybeetle C. montrouzieri (Coleoptera: Coccinellidae) and reported that although both species consumed all instars of the mealybug, they preferred to feed on younger instars. Fand and Suroshe (2015) listed 28 species of natural enemies (12 predators and 16 parasitoids), but noted that Aenasius bambawalei Haayat (Hymenoptera: Encyrtidae) was most important. Life Cycle and Description. The life cycle of a female Madeira mealybug consists of the egg stage, three nymphal instars, and the adult stage. The male, however, has an additional nymphal instar. Those destined to be males construct a loosely woven filamentous cocoon after the second instar and undergo a molt to the prepupal stage before molting again to the pupal stage. Thus, the extra instar of the male is called a prepupa. In both sexes, the first instar is usually called a crawler and the terminal instar is called a pupa. The first instar is the dispersive stage of females. The biology is summarized by several authors, including Joshi et al. (2010), Vennila et  al. (2010), Prasad et  al. (2012), Huang et al. (2012), and Fand and Suroshe (2015). The total development period from egg to adult is usually reported to be 22–40 days, with males requiring slightly longer to complete development. This relatively short life cycle leads to multiple generations per year. In India, for example, up to 12 overlapping generations are reported annually in some regions, and damage to cotton occurs where 6–8 generations occur annually. Both temperature and host

plant can affect mealybug development times (Prasad et al., 2012; Mamoon-ur-Rashid et al., 2012). The development of cotton mealybug when reared on tomato in the greenhouse was studied by Huang et al. (2012). When cultured at 25°C and 70% RH, they reported that mean development times were 3.5, 6.3, 5.6, and 6.4  days for instars 1–3 and the pupal stage, respectively. The range of development times was reported as 2–5, 4–12, 4–9, and 5–9 days for these same periods. The mean total development times (range) were given as 16.2 (16–20) and 15.4 (13–19) days for males and females, respectively. Under field conditions, however, generation time is usually given as 28–35 days (Fand and Suroshe, 2015). The mean lengths and widths of the nymphs and pupae were reported to be 0.622 and 0.22; 1.13 and 0.47; 1.68 and 0.72; and 1.58 and 0.59 for instars 1–3 and pupae, respectively. In these studies, adult males lived only about 1.8 days, whereas females lived about 42.6 days. Mean fecundity (production of crawlers) was 134.6 offspring. Males are much smaller (mean of 0.99 mm in length) than females (mean of 3.12 mm in length). In contrast, Prasad et al. (2012) conducted rearing studies at 10 temperatures using cotton as the host plant. When reared at 25°C, mean development times were 7.4, 9.9, 7.9, and 25.2 days for females in the egg plus instar 1, instar 2, instar 3, and the total development time, respectively. The rate of development increased until a temperature of about 32°C, which appeared to be optimal, then began to decrease as the temperature increased, but there was incomplete development at 39°C. In these studies, fecundity ranged from 154.3 to 244.9 offspring per female, with the maximum fecundity at 30°C. Egg. The female deposits her eggs in a structure called the ovisac. The white ovisac is composed of fluffy, loosetextured wax strands, and contains yellowish eggs measuring 0.3–0.4 mm in length. Egg production is temperature sensitive and ranges from 150 to 600 eggs per female. The optimal temperature for oviposition is 25–32°C. Duration of the egg stage is reported to be unusually short in duration, usually less than 2 h. Nymph. The small, oval nymphs are somewhat flattened, and covered with a white, powdery, waxy secretion. They also possess waxy filaments extending outward from around the margin of the body. So in many respects, they resemble the adult females. Immature females have three instars, but males have four. Those destined to be males construct a loosely woven filamentous cocoon after the second instar and undergo a molt to the prepupal stage before molting again to the pupal stage. Adult. The male and female are distinct, with the males not often observed. As with other mealybugs, the adult

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f­ emales are oval, somewhat flattened, covered with a white, powdery, waxy secretion, and lacking wings. The body beneath the wax is dark green to almost black. They also tend to bear narrow, transverse, dark bands, especially across the thoracic region, although this character is variable and not reliable for diagnostic purposes. Adult females range from 2.3 to 5.5 mm long and 1.5 to 4.3 mm wide. A series of waxy filaments extends outward from around the margin of the body, with the pair of longest terminal filaments. The adult male resembles a gnat (small fly) and bears two wings and two pronounced terminal filaments. Some aspects of reproduction are not clearly understood or inconsistent. Prasad et al. (2012) failed to observe oviposition in the absence of mating, whereas Fand et al. (2015) report that unmated females produce eggs but they are not viable, and Vennila et al. (2010) claimed that parthenogenetic reproduction occurred under field conditions.

Damage Mealybugs damage plants by extracting sap. They feed on foliage, flowers, stems, and roots. Loss of sap causes plants to become chlorotic and drops leaves from the plants. Fruit may be similarly affected. The leaves, fruit, and even the entire plant may take on a deformed, stunted form. This, of course, can result in reduced yield by affected plants. Mealybugs also excrete honeydew, which supports the growth of sooty mold and attracts ants. Growth of sooty mold on the leaf surface can inhibit photosynthesis. When ants are attracted to honeydew, they typically defend the producers of the honeydew from predation and parasitism; this can lead to a population explosion of the mealybugs because they are protected from natural enemies. Honeydew on the plants can also interfere with harvesting of cotton. Some ants will relocate these mealybugs to other sites. Presently, it is not considered a very damaging pest in the United States, but based on its ability to cause injury elsewhere, it remains a threat. Its reputation as a serious pest of cotton is likely due, at least in part, to the intensive use of insecticides on this crop in some areas. Except the first instar, mealybugs are generally covered with a waxy layer that repels liquids, including liquid insecticides.

Management Sampling. These mealybugs can be found in all parts of the plant, including below-ground, so thorough inspection is needed to detect their presence accurately. The presence of waxy masses, honeydew, or deformed leaves are often an indication of infestation. Visual and hand inspection are usually used for routine monitoring, though yellow sticky traps are sometimes used to detect airborne dispersants. In visual inspection, look particularly closely at the junction of the petiole and leaf blade, especially on new growth, and beneath leaves. Check nearby weeds as well as crop plants.

It is also important to monitor sites before planting for residual populations of mealybugs on: crop debris that has still not been tilled, crops that are unharvested, plants that have sprouted from spilled seed, or for mealybugs living on weeds. Any populations found during inspection should be eliminated before they move to harvest a new crop. The presence of a sex pheromone has been suggested by Xu et al. (2016). However, none has been identified. Insecticides. Insecticides can be effective, but timing is important. Older mealybugs have a dense layer of wax that serves to protect the insect from coming into contact with aqueous insecticide, so timing the spray to affect the less waxy instars, particularly the crawlers, is critical in obtaining control of mealybugs (and other honeydew-­ producing insects).

Madeira Mealybug

Phenacoccus madeirensis Green (Hemiptera: Pseudococcidae)

Natural History Distribution. The origin of Madeira mealybug is believed to be from Mexico or from the southern United States, and it is thought to have been accidentally transported to tropical areas of Central and South America, then to Madeira and West Africa (Williams, 1987). More recently, it has established in Bermuda, North Africa, southern Europe, and southern Asia. In the United States it occurs naturally in the southern states from Florida to California, and in Hawaii. It is easily transported, usually on g­ reenhouse-grown ornamental plants, so it can appear elsewhere. It can be a pest in greenhouses nearly everywhere. Host Plants. Madeira mealy bug has a very wide host range; Muniappan et  al. (2009) report that it is known to affect 44 plant families. It has been collected from crops such as eggplant, potato, tomato, pepper, okra, various types of bean, cassava, cotton, and fruit such as citrus, pineapple, mango, and passion fruit. Among the food crops, only potato, eggplant, peppers, and basil have been seriously affected, but it is rapidly spreading to new areas and is likely to affect additional crops. Ornamental plant hosts are more commonly noted as hosts, and include such diverse plants such as mandevilla, Dipladenia (Mandevilla) splendens (Apocyanaceae); croton, Codiaeum variegatum (Euphorbiaceae); garden verbena, Verbena x hybrid (Verbenaceae); scarlet sage, Salvinea coccinea (Lamiaceae); chrysanthemum, Dendranthema spp. (Asteraceae); sunflower, Helianthus sp. (Asteraceae); geranium, Geranium sp. (Geraniaceae); Transvaal daisy, Gerbera jamesonii (Asteraceae); schefflera, Schefflera actinophylla (Araliaceae); peregrina, Jatropha integerrima (Euphorbiaceae); hibiscus, Hibiscus spp. (Malvaceae); blue mistflower, Chromolaena odorata (Asteraceae);

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­common coleus, Plectranthus scutellarioide (Lamiaceae); rose, Rosa spp. (Rosaceae); and lantana, Lantana camara (Verbenaceae). Natural Enemies. In general, the natural enemies of this mealybug are poorly documented. Generalist predators such as the mealybug destroyer, Cryptolaemus montrouzieri Mulsant (Coleoptera: Coccinellidae), likely can affect biological suppression. However, the parasitic wasps Anagyrus spp. (Hymenoptera: Encyrtidae), have been the principal focus of attention thus far (Chong and Oetting, 2008). Life Cycle and Description. The life cycle of a female Madeira mealybug consists of the egg stage, three nymphal instars, and the adult stage. The male, however, has an additional nymphal instar. Those destined to be males construct a loosely woven filamentous cocoon after the second instar and undergo a molt to the prepupal stage before molting again to the pupal stage. Thus, the extra instar of the male is called a prepupa. In both sexes, the first instar is usually called a crawler and the terminal instar is called a pupa. Madeira mealybug reproduces sexually, and the ratio of males to females is 1:1 (Chong et al., 2003). The relatively short life cycle leads to multiple generations per year. In Sicily, for example, 5–6 overlapping generations are reported annually (Papadopoulou and Chryssohoides, 2012). Overwintering occurs as young nymphs on the underside of leaves or in crevices in bark. When females were reared at 25°C on four different host plants, Tok et  al. (2016) reported that the mean durations of the egg, first nymphal, second nymphal, third nymphal, and egg to adult periods required 4.0–6, 6.2– 10.6, 4.9–9.8, 5.2–10.1, and 20.4–35.6 days, respectively. Males required about the same amount of time (the prepupa and pupal stage were combined and reported as the third nymphal period in this tabulation): 4.0–6.2, 6.6–12.5, 4.6–10.4, 6.7–7.9, and 21.9–35.1 days, respectively. These authors also reported the mean length of time required for preoviposition by adults, the oviposition period, adult longevity, and the numbers of eggs produced when reared on the four host plants were 7.1–10.0, 4.2–5.0, 3.7–4.7 (males) or 12.8–15.2 (females) days, and 132.3–233.4 eggs, respectively. Chong et al. (2003) conducted a similar study and reported comparable development times when reared on chrysanthemum at 25°C, but also studied growth and survival at other temperatures. Perhaps the most interesting aspects of this study were that the immature insects did not develop successfully at 30°C, and that maximum egg production occurred at 20°C. However other studies have reported survival and development at 30°C. Nevertheless, although Madeira mealybug is widely distributed in tropical areas, it seems to be well adapted for subtropical and perhaps temperate conditions.

FIG. 8.61  Madeira mealybug eggs. (Photo by L. Buss.)

Egg. The female produces white wax strands and forms a tubular structure called ovisac, in which she lives and deposits her yellow eggs. The eggs measure from 0.3 to 0.5 mm in length. The number of eggs varies, but usually is in the 100s per ovisac. Egg production is temperature dependent, with a mean number of 388 eggs per female at 15°C, 491 per female at 20°C, and 288 per female at 28°C (Chong et al., 2003).

FIG. 8.62  Nymph of Madeira mealybug. (Photo by L. Buss.)

Nymph. First instars are called crawlers because they are quite mobile; this is the most important dispersive stage of mealybugs. The crawlers are yellow, about the same size as the eggs, and is oval in shape. Older nymphs can move too, but do not travel very far. The second instars are slightly larger than the crawlers and begin to accumulate wax, taking on a whitish or gray tint. The third instar females are larger than the second instars, yet have more wax on the surface of the body, and show faint gray lines on the dorsal surface. The males are smaller than the females in the third instar, and they begin to produce waxy filaments that form a loose cylindrical covering in which they molt into a fourth instar.

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The filamentous covering produced by the males is smaller (often about 2.5 mm long) and the filaments are less dense, than the ovisac (which measures up to 6 or even 8 mm long). Although the sexes are indistinguishable during the first and second instars, in the third they can be distinguished, and of course the fourth instar is limited to males. After the third instar (females) or fourth instar (males), the mealybugs molt into adults. Adult. Adult females are wingless and retain the oval body form of the third instar, lacking wings. The body beneath the wax is greenish. Two darker lengthwise stripes may be present, though the pigmentation is weak. The female is 2–3 mm long. Males look quite different, however, resembling gnats (small flies) and bear wings and pronounced terminal filaments. The sex ratio is usually about 1:1. Madeira mealybug, like other mealybugs, has the body nearly covered with small particles of white wax. Also, when viewed from above, the periphery is ringed by small waxy filaments. Those at the posterior end are longer, up to 3–4 times as long as those occurring elsewhere, but not greatly elongated. Adult females produce large amounts of filamentous wax, forming a tubular structure called an ovisac. They lay their eggs within the ovisac. A good description of Madeira mealybug is provided by Williams (1987). Madeira mealybug, P. madeirensis, is easily confused with Mexican mealybug, Phenacoccus gossypii Townsend and Cockerell. Mexican mealybug is limited in distribution, and occurs mainly in Mexico, Texas, and Florida. Many records from around the world attributed to P. gossypii are, in fact, P. madeirensis. Morphologically, P. madeirensis is distinguished by the absence of dorsal multilocular disc pores on the mid-thorax, whereas P. gossypii has these pores present on the mid-thorax. Because these characters are difficult to discern, species determination is best left to experts. A key for the identification of these and related mealybugs found in North and South America was published by Williams (1987).

Damage Madeira mealybug feeds on plant sap (phloem) from both the leaves and stems of plants. They tend to be concentrated on the stems and the principal veins of the leaves. Their feeding causes wilting, plant distortion, yellowing of the plant, and ultimately it may cause death of the host. Honeydew can be deposited on foliage, discoloring the tissue, and interfering with photosynthesis.

Management Sampling. Mealybugs tend to establish on apical, unexpanded leaf tissues and at the intersection of stems or stem and leaf petioles. Thus, they can be well hidden

from casual observation and it takes careful inspection to detect them early in an infestation. A sex pheromone has been identified that can be used to attract males (Ho et al., 2009, 2011). Insecticides. The waxy secretions of mealybugs can interfere with insecticide adhesion, so timely application of insecticides for the earliest instar feasible is generally recommended, as wax coverage is less. Less toxic products such as insecticidal soaps, horticultural oil, or neem oil can be applied to mealybugs, with some suppression obtained, especially against younger nymphs that have less wax accumulated on their bodies. Systemic insecticides can be used to reduce mealybug numbers on some plants. These are available as liquids for spraying or drenching the roots, as spikes that are inserted into the soil, and as granules that release their toxicant when watered into the roots. Cultural Practices. Ants are often associated with mealybugs, with the mealybugs providing honeydew to the ants, and the ants providing the bugs with protection from predators. Disrupting the ant population often is an important element in obtaining control of mealybugs (and other honeydew-producing insects).

Pink Hibiscus Mealybug

Maconellicoccus hirsutus (Green) (Hemiptera: Pseudococcidae)

Natural History Distribution. This insect is probably native to southern Asia, but has been introduced to many tropical areas of the world, including Africa and northern Australia, and most recently to the Western Hemisphere. It was observed in Hawaii in 1983 and the Caribbean region in 1994, then California in 1999, and Florida in 2002. Its potential range in North America includes much of the southern United States. Host Plants. As its common name suggests, hibiscus is a favorite host of this mealybug. In fact, many woody ornamental, fruit, and forest trees support high population densities of hibiscus mealybug. Among the important economic hosts are such tropical fruits as avocado, banana, carambola, citrus, custard apple, grape, guava, mango, mulberry, passion fruit, and soursop; and ornamentals such as croton, heliconia, and hibiscus. It is also a minor pest of cotton. Vegetables are infested mostly when the mealybug population density on trees and shrubs is high, with incidence of infestation of vegetables declining as overall mealybug densities decrease. Among the vegetable hosts, most susceptible to infestation are bean, beet, carrot, cowpea, cucumber, okra, pepper, pigeon pea, squash, and tomato, but among the other vegetables occasionally infested are asparagus, cabbage,

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lettuce, onion, potato, pumpkin, sweet potato, and yam. In the presence of natural enemies, however, the threat to vegetable crops is greatly reduced. Natural Enemies. In Asia and Africa, many predators and parasitoids are reported to attack pink hibiscus mealybug, and some have been introduced to other countries to implement biological suppression (Mani, 1989). Nevertheless, it remains a pest in Egypt and India (Mani, 1989). The parasitoid Anagyrus kamali Moursi (Hymenoptera: Encyrtidae) was apparently accidentally introduced to Hawaii simultaneously with the mealybug nearly around 1982, and has held this pest in check. When pink hibiscus mealybug attained the Caribbean area in the mid-1990s, considerable damage was caused in Grenada, Trinidad, and elsewhere until Anagyrus kamali could be successfully established. Kairo et  al. (2000) summarized the program for biological control of pink hibiscus mealybug in the Caribbean region, and they attribute successful biological control to a combination of A. kamali and Cryptolaemus montrouzieri Mulsant (Coccinellidae). Roltsch et al. (2006) reported on the introduction of three parasitoids to California: A. kamali, Gyranusoidea indica Shafee, Alam, and Agarwal (Encyrtodae), and Allotropa sp. nr. mecrida (Walker) (Platygastridae). Not surprisingly, A. kamali was the predominant parasitoid, but G. indica was effective during the winter months, and Allotropa sp. did not appear to establish. Many parasitoids and predators attacking other mealybugs can attack pink hibiscus mealybug as well. Life Cycle and Description. In subtropical climates a generation is completed in about 30–40 days and about 10 generations occur annually. Complete development from egg to the adult stage normally occurs in 25–26 days, but it is temperature dependent. When reared at constant temperatures, the optimal temperature for development (fastest development) was reported to be 27°C, allowing completion of development in about 29 days, or about 300 degree days (Chong et  al., 2008). Amarasekare et  al. (2008) reported about the same developmental responses, and also noted that 80%–90% of eggs hatched between 20°C and 30°C and that the highest fecundity occurred at 25°C. In Egypt, reproduction is usually parthenogenetic, but males are sometimes produced, and both sexual and parthenogenetic reproduction occurs in many populations. In the Caribbean region, biparental reproduction apparently occurs exclusively (Williams, 1996). Hibiscus mealybug survives during cold weather in all stages, but the egg stage is particularly hard. This insect is characterized by the presence of waxy white cotton-like secretions, so infested plants have a white fuzzy appearance. If the cottony material is removed, however, the eggs, nymphs, and adults are revealed to be pink.

Egg. The female produces a cottony egg sac, which is attached to the host plant. The oval egg sac is about twice as long as wide and consists of loose fibers and eggs internally and matted fibers externally. Each egg sac contains 80–650 eggs, which turn pink before hatching. The oval eggs measure about 0.35 mm long and 0.20 mm wide. Eggs hatch in 3–9 days. Nymph. The nymphs are elongate oval. Initially they are orange but then later turn pink. Newly hatched nymphs (crawlers) are mobile, but soon settle and begin feeding. There are three instars in females. Mean duration of the female instars is about 6.7, 6.5, and 7.9 days for instars 1–3, respectively. In males, there are two nymphal stages followed by two “pupal” stages. Mean duration of the male instars is 6.6, 6.5, 1.0, and 5.6 days, respectively. Three pairs of long legs and moderately long six-segmented antennae are evident in nymphs, and the anal region bears a pair of stout hairs. The piercing-sucking mouthparts are narrow and difficult to observe. Occasionally, waxy secretion is found in the posterior region. Pupa. In males, the third and fourth instars are nonfeeding stages in which the nymph transforms into a winged adult. The third instar (puparium) is somewhat elongate and resides in a loose mass of fine white filaments. It measures 1.1–1.5 mm long and 0.35–0.45 mm wide. The fourth instar (pupa) is brownish and shows evidence of wing formation. The antennae are directed posteriorly, and closely resemble the adult, though lacking the terminal abdominal filaments of the adult male. It measures about 1.25 mm long and 0.4 mm wide.

FIG. 8.63  Adult male of Madeira mealybug. (Photo by L. Buss.)

Adult. The adult female is elongated oval, bears three pairs of relatively small legs, and short but apparent with nine-segmented antennae. Occasionally, waxy secretion is

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found in the posterior region. The female is pink and measures 2.0–3.0 mm long and 0.9–2.0 mm wide. It bears stout hairs at the posterior end of the body and is wingless. The adult female may disperse away from the terminal growth of the plant if it is withering and unsuitable for feeding. The preoviposition period varies from 0.5 to 6 days, followed by an ovipositional period of 4–8 days. Oviposition may occur at the terminal portion of the plant, but during cool weather more sheltered locations are sought for oviposition. The adult male is slightly smaller than the female in size. The male is pinkish, elongate, and narrow in body form. It bears one pair of wings and three pairs of moderately long legs. The antennae are prominent and 10-segmented. The tip of the abdomen bears a pair of long, stout filaments that are white. A sex pheromone has been identified and can be used for population monitoring (Zhang and Amalin, 2005). A detailed synopsis of pink hibiscus mealybug biology was given by Mani (1989) and Williams (1986, 1996). Ezzat (1958) and Williams (1996) provided a technical description of the species, and the latter author also presented a key to the species of Maconellicoccus. Miller (1999) presented descriptions and keys to the instars, and comparisons with other common pest species. Culture techniques for pink hibiscus mealybug are provided by Serrano and Lapointe (2002).

determine its presence. If natural enemies are established in an area, particularly Anagyrus kamali, the mealybug should not be a severe pest. In the absence of natural enemies, or if natural enemy activity is disrupted by pesticides, this mealybug can be damaging. Insecticides can provide some control, particularly if systemic insecticides are applied. The application of granular formulations may be necessary for protection of crops with susceptible below-ground product, such as potato. The waxy secretions produced by the mealybugs greatly reduce the effectiveness of contact insecticides. Ants attend pink hibiscus mealybug, so elimination of ants can favor suppression of mealybug by natural enemies. Some natural enemies of pink hibiscus mealybug, including the lady beetle Cryptolaemus montrouzieri Mulsant (Coleoptera: Coccinellidae), are available from commercial insectaries and can be released into affected areas to supplement naturally occurring biological control.

Damage

Distribution. This is a tropical species, and is found in the Caribbean region. However, there are two subspecies that inhabit different areas. The subspecies P. suturellus suturellus occurs in Alabama, Georgia, South Carolina, Puerto Rico, and many other islands in the Caribbean region, though it is most abundant in southern Florida and Cuba. The subspecies P. suturellus capitatus is known from Central America, extending from southern Mexico south to Panama. Host Plants. As suggested by its common name, cotton is an important host plant. However, it is also reported to feed on okra pods, eggplant, tangerine and orange fruit, guava, papaya, and avocado. Other plants fed upon include roselle, Hibiscus sabdariffa (Malvaceae); rose of Sharon, Hibiscus syriacus (Malvaceae); Turk’s cap, Malaviscus arboreus (Malvaceae); rose buds and blossoms, Rosa spp. (Rosaceae); oleander, Nerium oleander (Apocynoideae); Spanish needle, Bidens pilosa (Asteraceae); Portia tree, Thespesia populnea (Malvaceae); Caesarweed, Urena lobata (Malvaceae); black nightshade; Solanum nigrum, (Solanaceae); and teaweed, Sida sp. (Malvaceae). So far as is known, the normal host plants all are in the families Malvaceae, Sterculiaceae, Bombacaceae, and Tiliaceae, which are closely related. The adults and late instar nymphs feed on the ripe or ripening protein-rich seeds of their host plants. As these are available at different times of year, the

These mealybugs may infest any portion of plants, including the below-ground portions. However, the stems and terminal shoot tissues are favored. They secrete a toxic saliva that causes various symptoms in the host plant. Typical symptoms are severe malformation of shoots and leaves, including twisting and crinkling of the foliage. Growth is stunted and tip growth may be bushy rather than elongate. Infested flowers drop and fruit is not produced, or they are small and malformed. Honeydew may accumulate and attract ants, and sooty mold may develop. In hibiscus, one of the most preferred hosts, galls are produced on terminal growth. Plants with cracks and crevices in the bark, as are typically found on woody plants, are often susceptible to infestation. These cracks and crevices provide microrefugia where the mealybugs can escape predation. With more damage-tolerant plants, damage may initially be confined to the new growth, but as the plant dies back the mealybugs will move down the plant to find food.

Management As hibiscus is the preferred host, these plants should be monitored for the presence of pink hibiscus mealybug in an area. Traps baited with sex pheromone can also be used to

PYRRHOCORIDAE—COTTON STAINERS Cotton Stainer

Dysdercus suturellus (Herrich-Schaeffer) (Hemiptera: Pyrrhocoridae)

Natural History

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winged adults disperse among different plant species. The adults have also been observed feeding on caterpillars and pupae of moths, and on plant nectar and honeydew. Natural Enemies. The natural enemies have not been critically assessed. There are many observations of birds and predatory insects feeding occasionally on cotton stainer, but their effects are unknown. The only natural enemies that seem to occur regularly and tachinid flies (Tachninidae). As is the case with many bugs (Hemiptera), the adults tachinids attach eggs to the bug nymphs, the fly larvae burrow into the body of the bug, and the adult bug is eventually killed by emergence of the parasitic fly from the bug. Life Cycle and Description. There are several generations per year. Duration of the life cycle varies from 1 to 3.5  months, depending on temperature. van Doesburg (1968) suggests that members of this genus engage in prolonged copulation of the host plant, interrupted by periods of oviposition; they copulate again between the production of batches of eggs. This species is included in a treatment of economically important Heteroptera by Schaefer and Panizzi (2000). Egg. The eggs are ovoid, being more pointed at one end. They are yellow, and scattered singly or in small loose clusters on the soil or organic debris, or on the soil. The female deposits about 100 eggs. The eggs hatch in about a week.

FIG. 8.65  Mating adults of cotton stainer. (Photo by L. Buss.)

Adult. The adult of D. suturellis suturellis measures 10– 15 mm long. The head, pronotum, and scutellum are bright red, as is the abdomen, though the abdomen is somewhat covered by the dark gray to black wings. The edges of the front wings are bordered with white producing a large X on its dorsum when the wings are folded. The pronotal collar and margins of the abdominal sternites are whitish. The legs and antennae are the same color as in the nymphal stage. The other subspecies, D. suturellis capitata, differs slightly in color. Specifically, the head is black and the abdomen is yellowish white.

Damage

FIG. 8.64  Nymph of cotton stainer. (Photo by J. Castner.)

Nymph. There are five nymphal instars. The first instar is reportedly spent underground. The duration of all except the last instar is about 5 days each under warm conditions, though the terminal instar requires twice along. Thus, the total duration of the nymphal stage is about 21–35 days. The color of the nymphs is bright red, though the fourth and fifth instars have darker wing pads, and the lines marking the abdominal segments are white, becoming increasingly distinct with maturity. The antennae and legs are mostly black, but red basally. The nymphs are gregarious, even to the extent of different species of Dysdercus mixing in the aggregations.

This is a blossom, fruit, and pod-feeding insect, wherein the bug feeds on the soft, developing seeds. Feeding also allows entry of microorganisms that induce fruit drop. Cotton stainer feeds mostly on plants in the family Malvaceae, but rarely attains great abundance. When it was reportedly abundant (the early 1900s), it was nearly always in association with cotton production. Cotton is no longer grown very often in areas inhabited by cotton stainer, and it is no longer considered to be a serious pest. Cotton stainer derives its name from a yellow staining of the cotton lint that followed feeding by the bug on cotton bolls. The stain is variously attributed to seed exudate following bug feeding, and excrement by the bug, but the true cause is uncertain.

Management Modern insecticides would undoubtedly suppress cotton stainer should this be warranted. Before the availability of reliable chemical insecticides, agriculturalists were advised to engage in sanitation, especially destruction of weeds in the family Malvaceae. Also, it was considered not advisable to grow cotton in the vicinity of citrus because citrus fruits were readily damaged by the bugs.

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FAMILY PSYLLIDAE—PSYLLIDS Potato Psyllid

Bactericera cockerelli (Šulc) (Hemiptera: Psyllidae)

Natural History Distribution. This native insect pest occurs in Mexico, the western United States (west of the Mississippi River), and southern areas of western Canada (Alberta, Saskatchewan). Historically, it overwintered only in southern regions such as Arizona, New Mexico, Texas, and northern Mexico, then migrated north during the spring months, traveling both east and west of the Rocky Mountains, infesting susceptible crops throughout the area, and potentially attaining Alberta and Saskatchewan. During the 20th century, most damage occurred in Utah, Colorado, Wyoming, Idaho, Montana, Alberta, and Saskatchewan. However, it occasionally had been reported as far west as California and British Columbia and as far east as Minnesota and Quebec, but these were infrequent occurrences with no long-term establishment. Occurrence in northern regions was considered to be largely a function of long distance, wind-borne dispersal. The absence of suitable host plants in northern climates precluded the possibility of successful overwintering in these cold areas. Then, in 2011, a very damaging outbreak of potato psyllid occurred in Washington, Oregon, and Idaho potato fields. Suddenly, potato psyllid became a more important component of the potato fauna in the Pacific Northwest. Eventually, it was determined that potato psyllid had diverged into several distinct genetic types, or haplotypes. The central and southwestern haplotypes originate and mostly infest crops that occur east of the Rocky Mountains. A western haplotype originates in Baja California, Mexico, and infests crops west of the Rocky Mountains. Most surprisingly, a northwestern haplotype resides and infests crops in the Pacific Northwest, and was the source of the 2011 outbreak. This initially seemed impossible because it was thought that there were no suitable hosts that would allow overwintering. However, a perennial semiwoody weed from Europe called bittersweet nightshade (Solanum dulcamara L.) had invaded the area during the 20th century, and now provides a suitable host plant for overwintering, resulting in an enlarged economic impact (Horton et al., 2015). Host Plants. Potato psyllid is also sometimes known as “tomato psyllid,” and these common names accurately indicate the most preferred hosts. However, in addition to potato and tomato, this psyllid can develop on other ­solanaceous vegetables, particularly pepper and eggplant. Solanaceous weeds such as ground-cherry, Physalis spp.; black nightshade, Solanum nigrum; buffalo bur, Solanum rostratum; matrimony vine, Lycium halimifolium; and

­bittersweet nightshade, Solanum dulcamara L. also support reproduction. Matrimony vine is believed to be an important overwintering host in southern Arizona, as is bittersweet nightshade in the Pacific Northwest. Adults have been collected from numerous plants, including conifer trees, but this is not an indication of the true host range. With the possible exception of field bindweed, Convolvulus arvensis, nonsolanaceous plants seem to be relatively unimportant, and even solanaceous weeds are not as important as crop plants. Natural Enemies. Several predators and parasites of potato psyllid are known, though there is little documentation on their effectiveness. Under rather artificial conditions the larvae and adults of lady beetles (Coleoptera: Coccinellidae), lacewings (Neuroptera: Chrysopidae), flower flies (Diptera: Syrphidae), bigeyed bugs (Hemiptera: Lygaeidae), minute pirate bugs (Hemiptera: Anthocoridae), and damsel bugs (Hemiptera: Nabidae) consume psyllid nymphs and adults (Pletsch, 1947). The wasps Metaphycus psyllidus Compere (Hymenoptera: Encyrtidae) and Tamarixia (Tetrastichus) triozae Burks (Hymenoptera: Eulophidae) parasitize potato psyllid. The latter species feeds externally on nymphs and attacks other psyllid species as well, and though sometimes abundant, is often absent from psyllid populations. Liu et al. (2012a,b) identified insecticides that were compatible with parasitic activity by T. triozae. Weather. Weather is an important element of biology and damage potential. These psyllids seem to be adapted for warm, but not hot weather. List (1939a), for example, demonstrated how prolonged exposure to 32°C depressed egg production and egg hatching. Temperatures of about 21– 27°C are considered optimal. Dispersal northward occurs when higher temperature is attained; temperatures of about 32°C and higher are deleterious to psyllids. Heavy precipitation may be detrimental, but this is less certain. Because psyllids disperse passively over long distances, wind patterns are also critical in determining their occurrence, and account for much of the variability in damage. Once invading psyllids have established, they multiply and cause greatest damage at moderate temperature, and populations fail to develop to damaging levels if the temperature is high. Life Cycle and Description. A generation can be completed in 20–30 days, depending on temperature. The number of generations varies considerably among regions. In Montana, only three generations can be completed before frost kills the host plants. In northern Utah, 3–4 generations are known, and in Colorado there are 4–7 generations. Once psyllids invade an area, however, prolonged oviposition by adults causes the generations to overlap, so it is difficult to distinguish generations. The central and southwestern haplotypes of potato psyllid overwinters in southern Arizona, New Mexico, Texas, and northern Mexico, where populations build to high densities in early spring. By early summer, the psyllids

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d­ isperse from the spring breeding areas northward to Utah and Colorado and shortly thereafter to Nebraska and eastern Wyoming. In July, as the temperature becomes hot in these mid-summer breeding areas, psyllids disperse northward to the northern states and Canadian provinces. Psyllids reappear in the overwintering areas between October and November, presumably having migrated south from northern locations. The western haplotype follows the same sequence of migration, but instead along the west coast, even attaining the Pacific Northwest. However, relatively few of the western strain make the northward trip successfully, and the infestation in the Northwest is attributable to the continued residence of the northwestern haplotype. Egg. The eggs are deposited principally on the lower surface of leaves, usually near the leaf edge, but some eggs can be found everywhere on suitable host plants. The eggs are oval and borne on thin stalks which connect one end of the egg to the leaf. They initially are light yellow, and become dark yellow or orange with time. The egg measures about 0.32–0.34 mm long, 0.13–0.15 mm wide, and with a stalk of 0.48–0.51 mm. Eggs hatch 3–6  days after deposition. Nymph. At hatching, the young nymph quickly escapes from the egg, then crawls down the egg stalk and searches for a place to feed. Nymphs are found mostly on the lower surface of leaves, sometimes on the upper surface, and almost never on the stems. Nymphs, and also adults, produce large quantities of whitish particulate excrement which may adhere to the foliage and fruit. Nymphs are elliptical when viewed from above, but very flattened in profile, appearing almost scale like. There are five nymphal instars. Nymphal body widths were given by Pletsch (1947) as 0.23, 0.35– 0.41, 0.53–0.58, 0.70–0.82, and 1.27–1.32 mm for instars 1–5, respectively, but body lengths were not provided. The body width measurements given by Rowe and Knowlton (1935) tend toward the lower end of the aforementioned size range; they also give nymphal lengths as 0.36, 0.50, 0.70, 1.10, and 1.60 mm, respectively. Initially, the nymphs are orange, but become yellowish green and then green as they mature. The compound eyes are reddish and quite prominent. During the third instar, the wings become evident in the form of pads that remain light in color and become more pronounced with each molt. A short fringe of wax filaments is present along the lateral margins of the body. Nymphs tend to remain essentially sedentary during their development, and prefer sheltered, shaded locations. Mean duration (range) of the instars was reported by Knowlton and Janes (1931) to be 2.8 (1–5), 2.4 (1–5), 2.5 (1–4), 2.7 (1–5), and 4.9 (3–9) days, respectively, for instars 1–5. Thus, nymphal development time required 12–21  days, though the average was about 15 days. These data were confirmed by Yang et al. (2010a,b,c), who documented mean nymphal develop-

ment times of 15.9 and 15.2 days under field and laboratory conditions, respectively. Yang and Liu (2009) determined the development on eggplant and bell pepper. Not surprisingly, nymphal development time was longer on these less favored hosts, requiring a mean of about 19 and 20  days, respectively.

FIG. 8.66  Potato psyllid. (Drawing by USDA.)

Adult. The adults are quite small, and measure only about 2.5–2.75 mm long. In general body form they resemble miniature cicadas, largely because they hold their wings angled and roof like over their body. They have two pairs of transparent wings. The front wings are considerably larger than the hind wings. The antennae are moderately long, about the length of the thorax. Body color ranges from pale green at emergence, to dark green or brown within 2–3 days, and gray or black thereafter. White or yellow lines are found on the head and thorax, and whitish bands on the first and terminal abdominal segments. Adults are active and easily disturbed, a distinct contrast to the nymphal stage. Adults disperse readily if disturbed, usually moving downwind (Henne et al., 2010). The preoviposition period of adults is normally about 10 days, with oviposition continuing for about 20 days. Total adult longevity is normally 25–35 days. Pletsch (1947) reported mean longevity of females to be 20, 23, 29, and 14  days when reared on tomato, potato, eggplant, and pepper, respectively. He also observed mean fecundity of 67, 258, 187, and 53 eggs on these same hosts. Knowlton and Janes (1931), working with potato foliage, reported mean fecundity in various trials of 300–400 eggs, with some individuals producing 1100–1350 eggs. Yang et  al. (2010a,b,c) observed a similar fecundity under laboratory conditions, averaging nearly 400 eggs per female on potato. In contrast, Yang and Liu (2009) reported 338 and 403 eggs per female on eggplant and bell pepper, respectively. The biology of potato psyllid was described by many authors including Knowlton and Janes (1931), List (1939a,b), and Wallis (1946, 1955). However, Pletsch (1947) provided the most complete summary and Cranshaw (1993) published an annotated bibliography. Munyaneza (2012) and Horton et al. (2015) provide an excellent recent summaries of this insect and its impact. Keys to the North American Psyllidae were given by Tuthill (1943).

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Damage Feeding by nymphs of potato psyllid causes a disease known as “psyllid yellows” (Eyer, 1937; Arslan et al., 1985). Psyllids apparently produce a toxin or growth regulator that disrupts plant growth; a plant pathogen is not involved in the inducement of psyllid yellows. Even a few psyllids can affect growth, but large populations are necessary for extensive disruption of plant growth. Symptoms of the disease include upward rolling of the young leaves, yellowing along the midrib and leaf margins, and sometimes induction of a purple color. Stem elongation is suppressed, plants are dwarfed, exceptionally numerous but small tubers and fruits are formed, and tubers sprout prematurely. In some areas, yield losses of 20%–50% are not uncommon in the absence of treatment, and complete losses have been reported in potato. This insect is considered to be the principal pest of tomatoes in some Rocky Mountain areas. Greenhouse-grown tomatoes are also affected. Elimination of psyllids from infested plants results in partial recovery of plants if they are suffering from psyllid yellows. Other psyllid species are occasionally reported from potatoes, but only B. cockerelli occurs in large numbers and causes damage. Beet leafhopper, Circulifer tenellus (Baker), induces similar symptoms in tomato, but this disease (called “curly top”) is caused by a plant virus. In addition to psyllid yellows, potato psyllid is vector of a bacterium known as Candidatus Liberibacter solanacearum, the causal agent of zebra chip disease of potato. In the recent years, this has been a cause for major concern by potato growers in North and Central America, and in New Zealand. Zebra chip-infected potato plants display yellow to purple discoloration of the foliage, shortened internodes, aerial tubers, and other abnormal growth. The tubers are reduced in size and become discolored by stripes and blotches of brown after they are sliced, particularly after they are fried. Zebra chip disease is a more serious problem than psyllid yellows, and is now well established throughout the major western United States potato-growing areas. It affects tomato, tomatillo, eggplant, and pepper in addition to potato. In recent years, damage to tomato, pepper, and eggplant has increased along the lower Rio Grande Valley of Texas, and potato in the Pacific Northwest. Unlike the case of psyllid yellows, the disease does not dissipate after the psyllids are killed. The disease pathogen also develops in solanaceous weeds such as silverleaf nightshade, Solanum elaeagnifolium Cav. that are fed upon by potato psyllid (Thinakaran et al., 2015).

Management Sampling. The adult populations are commonly sampled with the assistance of a sweep net, but egg and nymphal numbers are assessed by visual examination of foliage. The adults also can be sampled with yellow sticky or water-pan traps. Sweep net-sampling gives a better ­ assessment of

p­ syllid density, but yellow traps are adequate for determination of the presence. As is the case with many insects that are not strong fliers, psyllid populations typically are highest at field edges initially, but if not controlled they eventually spread throughout the crop. Insecticides. Traditionally, insecticides were usually applied only when psyllids were found in a field, with applications continuing at least until mid-season. However, with the onset of zebra chip disease, growers are more concerned about the presence of psyllids, and preventative actions (planting time or early treatments) are more common. Good coverage or use of systemic insecticides is important because psyllids are commonly found on the underside of foliage. Once plants are mature, and bear abundant foliage, they are less susceptible to injury, but the zebra chip pathogen begins to have developmental effects on potato within 3 weeks of inoculation, so it is important to prevent pathogen transmission as long as possible. For some other small insects such as aphids and whiteflies, the application of insecticidal soap or dilute dishwashing detergent can suppress populations, but adequate protection from psyllids using these products is infrequent. Even with conventional insecticides, this insect tends to be difficult to manage. Cultural Practices. Early-planted crops are more susceptible to injury than crops planted at mid-season or late, principally because only early crops are present when psyllids disperse into areas in the spring. However, part of the reason early-season crops are more injured is that psyllids do not thrive under the hot weather conditions found later in the summer, and large plants from early-season plantings protect the psyllids from the sun and heat (Wallis, 1955). Sanitation also is important because if discarded potatoes are allowed to sprout they can serve as temporary hosts. Weed management is useful for elimination of solanaceous weeds, which can serve as hosts for both psyllids and the bacterium causing zebra chip disease. Demirel and Cranshaw (2006e) reported that aluminum and white plastic mulch caused significant reductions in psyllid abundance relative to black plastic mulch or bare soil.

FAMILY THYRECORIDAE—EBONY BUGS Little Ebony Bug

Corimelaena pulicaria (Germar) (Hemiptera: Thyreocoridae)

Natural History Distribution. This common native insect is found throughout the United States and southern Canada, except possibly the Maritime Provinces of Canada. Its distribution also includes Central America. In the western United States

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a related and similar-appearing species, C. extensa Uhler, occasionally feeds on crops. Host Plants. This insect has been observed feeding on many plants. Among vegetables attacked are celery, potato, and sweet potato. It has also been associated with field crops such as chufa, clover, soybean, and sugarbeet. It is probably best known for damage to small fruits such as blackberry, grape, raspberry, and strawberry. Weeds are common hosts, including beggarstick, Bidens spp.; ragweed, Ambrosia spp.; redroot pigweed, Amarnathus retroflexus; wild carrot, Daucus carota; wild chervil, Cryptotaenia canadensis; plantain, Plantago lanceolata; common mullein, Verbascum thapsus, and others. Weeds appear to be preferred hosts. Natural Enemies. Natural enemies seem to be unknown. Life Cycle and Description. There is a single generation per year in the northern United States. The adult is the overwintering stage. Adults seek shelter under leaf litter and boards during the winter months. The adults are active early in the year, and eggs are deposited in May and June. Nymphs are found throughout the summer months. Egg. The eggs are deposited singly. They are oval in shape and orange to red in color and measures about 0.6 mm long and 0.4 mm wide. Incubation is about 10–14 days. Nymph. The young nymphs greatly resemble the adults, but the abdomen is colored red. There are five instars. Development time for the nymphs is about 30 days. Because the overwintered adults live through much of the summer and continue to oviposit, overlapping stages of development are found during the summer.

mouthparts of the negro bug are easily observed. The scutellum of these insects, and other members of the ­ family, are greatly enlarged and bluntly rounded, covering nearly the entire abdomen. This effectively hides all but the leading edge of the wings. Where the wings are exposed, along the perimeter of the abdomen, there is a whitish stripe. As its name suggests, the insect is almost entirely black, the principal exception being the aforementioned edges of the wings. This bug is quite small, and measures only 2.2–3.5 mm long. The antennae are five segmented, and the mouthparts consist principally of a four-segmented labium. The biology of this insect is poorly documented. Partial life histories were provided by Riley (1870a) and Davis (1893). McPherson (1982) gave a good summary, especially of host-plant records, and keys for identification. A key to distinguish stink bugs (including little ebony bug) commonly affecting vegetables also is found in Appendix A.

Damage This generally is a minor pest, but it does reach high numbers on occasion. On celery, the bugs are known to aggregate into small clusters and feed at the joint where the leaf petioles meet the stem. This causes the terminal leaves to wilt and die. The bugs then move progressively lower on the plant until even the youngest tissue is destroyed. In addition to its feeding injury, it is reputed to have a foul odor, and by walking on fruit, to impart a disagreeable taste (Riley, 1870a).

Management These insect are infrequent pests, and are readily controlled with foliar applications of insecticides. Because little ebony bug feeds predominantly on weeds, it is a good idea to keep weeds under control, or at least well separated from vegetable and berry crops.

FAMILY TINGIDAE—LACE BUGS Eggplant Lace Bug

Gargaphia solani Heidemann (Hemiptera: Tingidae)

Natural History

FIG. 8.67  Adult of ebony bug. (Photo by L. Buss.)

Adult. The adults are oval in overall form, and greatly resemble beetles. Examination of the mouthparts quickly separates the two groups, however, as the ­piercing-­sucking

Distribution. Eggplant lace bug occurs widely in the eastern United States north of New Hampshire and Iowa, and west to Kansas and Arizona, but it is not common in the Gulf Coast area. There are occasional reports from outside this range, including from Canada, but it is infrequently abundant in such areas. The range of eggplant lace bug also extends into Mexico. This appears to be a native insect.

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Host Plants. As its common name suggests, this insect affects principally eggplant, though it sometimes affects potato and perhaps tomato. Not surprisingly, it is commonly found on weeds in the plant family Solanaceae such as horsenettle, Solanum carolinense, and silverleaf nightshade, S. elaeagnifolium. Eggplant lace bug occasionally is reported from hosts other than Solanaceae, including Compositae, Leguminosae, and Malvaceae. Natural Enemies. Several common predators have been observed to attack eggplant lace bug, among them the lady beetles Hippodamia convergens Guerin-Meneville and Coleomegilla maculata De Geer (Coleoptera: Coccinellidae); the spined soldier bug, Podisus ­maculiventris (Hemiptera: Pentatomidae); the insidious flower bug, Orius insidiosus (Say) (Hemiptera: Anthocoridae); and various spiders. The fairyfly Erythmelus klopomor Triapsitsyn (Hymenoptera: Mymaridae) is an egg parasitoid of this species and other Hemiptera (Puttler et al., 2014). Life Cycle and Description. In Virginia, the complete life cycle takes about 20 days, and these lace bugs are active from May to November. Adults overwinter, often in clumps of grass but sometimes under bark. The number of generations is estimated at 6–8 per year, but overlap so completely that they are hard to distinguish. Egg. The eggs are deposited on the lower surface of leaves in clusters containing over 100 eggs. The eggs are attached on end, and are not erect, but rather tilted at different angles. They are about 0.37 mm long and 0.18 mm wide. They are brown basally and greenish apically. The tips of the eggs are recessed and surrounded by a delicate, lace-like border. The female may require 4–5 days to complete the egg cluster, depositing 15–60 eggs at each oviposition. At completion of egg laying, a sticky secretion is spread over the cluster, and the female tends the eggs, leaving only to feed for brief intervals. The incubation period is 5–8 days. Nymph. Nymphs remain clustered as they feed and grow. There are five instars, each with a duration of approximately 2  days. The first instar is light yellow, with pink eyes, and antennae are about as long as the body. This instar measures 0.3–0.4 mm long. At the molt to the second instar, which measures about 0.8 mm long, the nymph acquires large spines laterally on the thorax and abdomen. The third instar is about the same length as the preceding stage, but is wider, bears spines dorsally, and displays evidence of wing pads. The fourth instar measures about 1.5 mm long, the thorax is greatly expanded laterally, the entire body is covered with spines, and the wing pads extend to about the second abdominal segment. The fifth instar is about 2.2 mm long, and like the preceding instars, yellowish in color. This stage is also adorned with spiny processes, most of which are brown in color.

FIG. 8.68  Eggplant lace bug. (Drawing by J. Capinera.)

Adult. The adult eggplant lace bug is rather flat in general form, and mostly dark brown, though the legs are yellow. The front wings are distinctively lacy, with the basal cells more densely packed. The thorax is pronounced, flaring laterally but also extending posteriorly. The antennae are nearly as long as the body, and thickened apically. The adult measures about 4 mm long and 2 mm wide. Adults have a relatively brief period of oviposition, usually 4–5 days, after which they turn their attention to brood care. Eggplant lace bug adults display complex social interactions, including maternal behavior. For example, the female remains with the eggs and nymphs as they develop. As nymphs deplete the food and become restless, the adult leads them to a new feeding site. Nymphs respond to an alarm pheromone that is released when the bugs are crushed by dispersing quickly, only to reassemble at another location, accompanied by the adult. The alarm pheromone was described by Aldrich et  al. (1991a). The adults use wing fanning to deter predation, and also to communicate with nymphs. There is some evidence for an aggregation pheromone (Kearns and Yamamoto, 1981). Interestingly, females frequently deposit their eggs within egg clusters of other females, a behavior called “egg dumping.” This relegates the recipient female to surrogate mother status, allowing the donor female to continue to produce more eggs without the responsibility for care of her offspring (Tallamy, 1985). An excellent treatment of eggplant lace bug biology was given by Fink (1915), with supplementary information provided by Bailey (1951). Eggplant lace bug was included in keys to the Hemiptera of Missouri (Froeschner, 1944) and the lace bugs of North Carolina (Horn et al., 1979). Loeb (2003) discusses the evolution of egg dumping.

Damage The adults and nymphs suck sap from the leaves of host plants. The first symptom of injury is large discolored blotches or patches, usually circular in shape. This reflects the feeding by the adult, and perhaps the very young nymphs,

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in the vicinity of the egg cluster. Later, as the nymphs disperse away from the egg cluster, the d­ iscoloration spreads until the entire leaf is involved, turning yellow and dry. Often every leaf of an infested plant is discolored as the insects move from leaf to leaf. The adults disperse from plant to plant.

Management Although lace bugs are not often serious pests, they occasionally reach damaging levels. They are readily suppressed with insecticides applied to the foliage, but care must be taken to attain thorough coverage of the plant because these insects are found on the lower surfaces of the leaves.

Chapter 9

Order Hymenoptera—Ants and Sawflies

FAMILY ARGIDAE—SAWFLIES Sweetpotato Sawfly

Sterictiphora cellularis (Say) (Hymenoptera: Argidae)

Natural History Distribution. This native species occurs throughout the eastern United States, which covers as far west as eastern Nebraska and Texas. It has been known as a serious pest only in the Chesapeake Bay region, specifically Maryland, Virginia, and North Carolina and near the mouth of the Mississippi River, in Louisiana and Mississippi. Host Plants. Sweetpotato sawfly is mainly observed from Ipomoea spp., and rarely collected from other than sweet potato plants. Natural Enemies. A larval parasitoid, Schizocerophaga leibyi Townsend (Diptera: Tachinidae), is reported to be an effective biological control agent, having been reared from 60% to 70% of sweetpotato sawfly cocoons. Chapman and Gould (1929) attributed the small size of the autumn generation principally to the actions of this fly. A wasp parasitoid, Boethus schizoceri (Riley and Howard) (Hymenoptera: Ichneumonidae), is also observed from sweetpotato sawfly. Life Cycle and Description. Apparently, three generations occur annually in Virginia, with larvae present in July, August, and September. All three generations can be completed within 28 days. This insect is presumed to overwinter in the pupal stage. Egg. The eggs are inserted within the leaf tissue in rows along the principal veins on the lower surface of the leaf. They cause the leaf tissue to bubble or blister, and each blister contains a single egg. Rows of 6–12 eggs are common, and a single leaf may contain over 200 eggs. The egg is whitish and has a somewhat flattened oval shape. The egg measures about 1.6 mm long during hatching, but sawfly eggs enlarge before hatching. Eggs hatch in 6–7 days. Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00009-1 © 2020 Elsevier Inc. All rights reserved.

FIG. 9.1  Sweetpotato sawfly larva. (Drawing by J. Capinera.)

Larva. The larva is yellowish green and adorned with numerous black, blunt spines. The larva bears seven pairs of abdominal prolegs, but they are short and obscure, the terminal prolegs especially cryptic. Apparently, there are five instars. The duration of the larval stage is about 10 days. At maturity, the larva measures about 12 mm long. Pupa. When the larva completes its growth and is ready to pupate, it drops to the soil and constructs a brownish silken cocoon on, or near, the soil surface. The cocoon is an oval shape and measures about 14 mm long and 5 mm wide. The cocoon frequently has soil and leaf debris attached to the outside. The pupal stage apparently has not been described. The adult emerges from the cocoon about 9–12 days after cocoon formation.

FIG. 9.2  Sweetpotato sawfly adult. (Drawing by J. Capinera.)

Adult. The adults are reported to be short-lived and weak fliers. The body of adult males is shiny black, and the wings dusky. The female is similar in color except that her abdomen is orange-red. In both sexes, the legs are dark basally but lighter or colorless distally. Females measure 383

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about 7 mm long, with a wingspan of about 12 mm. The male is slightly smaller in size. The antenna of the male is branched whereas that of the female is unbranched. The biology of sweetpotato sawfly was described by Chapman and Gould (1929).

Damage The larvae are defoliators of sweet potato, consuming leaf tissue except stems and basal portion of the principal veins. As adults are not very dispersive, they tend to deposit many eggs in small areas. This leads to severe defoliation in very localized sections of fields, and sometimes the death of numerous larvae occurs as they exhaust the food supply. Repeated defoliation by sawfly may reduce the yield by 50%.

Management This species is infrequent and apparently has caused no large-scale damage since the early 1900s. Observations in the early 1900s revealed that damage was limited to the sweet potato cultivar “Big-Stem Jersey,” and changes in cultivar selection by sweet potato growers may have accounted for the demise of this insect as a pest. Foliar insecticides are effective, and crop rotation is recommended due to the poor dispersive ability of the adults.

FAMILY FORMICIDAE—ANTS Red Imported Fire Ant

Solenopsis invicta Buren (Hymenoptera: Formicidae)

Natural History Distribution. Red imported fire ant was accidentally introduced into the United States at Mobile, Alabama between 1933 and 1941. It gradually spreads over most of the southeast states, including Georgia, Alabama, Mississippi, and Louisiana in 1957, then adjacent states from North Carolina to Texas in 1985, and finally California in 1998. Portions of New Mexico and Arizona are also affected. This species seems destined to occupy nearly the southern half of the United States except very arid areas. Red imported fire ant also has successfully invaded Puerto Rico. Activity decreases on cold weather, and this species seemed ill adapted for cold-weather conditions, which is limiting northward spread. In the United States, the northern limits now are southern Tennessee and southern Oklahoma. The origin of Solenopsis invicta is South America, where it is most abundant in southwestern Brazil and in Paraguay. It has successfully invaded China and Australia. Red imported fire ant is not the only fire ant in North America, but it is the only species that commonly affect vegetable crops. Two native species, Solenopsis ­geminata

(Fabricius) or tropical fire ant and S. xyloni McCook or southern fire ant, also occur in the southeastern states, with S. xyloni also extending to California. Another immigrant species, S. richteri Forel or black imported fire ant, was also introduced from South America, but likely from areas south of the homeland of the red imported fire ant. Black imported fire ant is quite similar to red imported fire ant, though darker in color, and so initially these two species were confused. Black imported fire ant has a very restricted distribution and is limited to northern Alabama and Mississippi and western Tennessee. Red imported fire ant tends to replace most other species of ants when it invades a new area, including the aforementioned Solenopsis spp. Host Plants. These ants are omnivorous and feed on a mixture of plant and animal matter. They are effective predators of insects, spiders, earthworms, and other small invertebrates. Plant feeding is limited and often occurs when ants are deprived of other food. Nevertheless, fire ants are known to feed on vegetables such as bean, cabbage, corn, cucumber, eggplant, okra, potato, and sweet potato and on other crops such as young citrus trees, peanut, sorghum, soybean, and sunflower. Okra fruit is particularly at risk. Sweet plant exudates and honeydew from homopterous insects are readily consumed. Natural Enemies. Many natural enemies of red imported fire ant are known, but few seem to be effective under either South American or North American conditions. Competition from other ants is believed to be one of the most important factors limiting abundance and dispersal in their native land, but North American ants seem to be ineffective competitors of fire ants, at least within disturbed habitats. Colony survival, particularly of incipient colonies, is also affected by competition among fire ant nests. Minim workers from new colonies often raid brood from nearby nests, boosting the size of receiving colonies and allowing new colonies to achieve maturity sooner. Although raided colonies disappear, workers and queens from the losing colonies may be amalgamated into the receiving colonies (Tschinkel, 1992). Among the promising pathogens of fire ant are the fungi Kneallhazia (Thelohania) solenopsae and Beauveria bassiana, and several viruses. None has demonstrated consistent effectiveness in North America, though there is some evidence that Kneallhazia significantly suppresses reproduction of the ants under some conditions (Oi and Williams, 2002, 2003). Together, the pathogens and phorid parasitoids seem to have had suppressive effects in some areas. South American parasitoids, Pseudacteon spp. (Diptera: Phoridae), which decapitate ants during the final stages of development, have been introduced to the southern states of USA, and related species are being introduced. An unusual South American parasitic ant, Solenopsis daguerrei (Santschi) (Hymenoptera: Formicidae), invades fire ant colonies and takes over control of the fire ant colony, possibly through food diversion. This and other parasites are being

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investigated for the potential to achieve fire ant suppression. Natural enemies have been discussed by Jouvenaz (1983, 1986), Banks et al. (1985), and Wojcik (1986a). Life Cycle and Description. Ants are colonial insects, with the number of ants per colony depending on colony age and availability of food. Activities of the colony members are highly structured, with some members (castes) specialized for reproduction, tending of the young, and obtaining food. The seasonal reproductive cycle usually begins in March, and eggs from mature colonies give rise to both worker and reproductive castes. Workers predominate and the maximum proportion of sexual forms (about 10%) produced in June. A life cycle can be completed in about 30 days. The longevity of colonies is uncertain, though individual queens can live 6–7 years. Egg. Colonies may be initiated by a single-mated queen or several queens. Mating flights may occur at any time, but peak abundance is from May to August. Flights are preceded by rain, normally within 1–2 days, and at least 80% relative humidity. After the mating flight, the queen usually breaks off her wings and begins excavation of a burrow within 4 h of her mating flight. Initially, the burrow consists of a vertical tunnel 6–12 cm deep into the soil and small cells. The queen lays 15–20 eggs within 2–3 days and produces 20–125 eggs by the time the first larvae hatch. The newly deposited egg is whitish, oval in shape, and measures about 0.2 mm wide and 0.3 mm long. As the egg approaches maturity, it acquires the larval form. Only about half of the eggs are fertile, the remaining serve as food for the young larvae. Mean duration of the egg is 8.4, 5.2, 5.0, and 5.8 days when cultured at 25°C, 30°C, 32°C, and 35°C, respectively; the optimal development of eggs and other stages occurred at about 32°C (O’Neal and Markin, 1975b). Once the colony is firmly established, the queen can produce up to 2000 eggs per day. Larva. There are four instars, and all stages are whitish, C-shaped, and lack distinct appendages. As described by Petralia and Vinson (1979), first instars are hairless, and measure about 0.27–0.42 mm long. Head capsule width is 0.14–0.16 mm. Second instars have only a few simple hairs, measure about 0.42–0.57 mm long, and have a head capsule width of 0.16–0.19 mm. Third instars have moderately numerous short hairs of various types, measure about 0.59–0.91 mm long, and have a head width of 0.20–0.25 mm. Fourth instars are similar to third instars in general appearance, though the body hairs are slightly longer. The body length of fourth instars is 0.8–1.8 mm, and the head width is 0.26–0.32 mm. Head capsule widths increase slightly during both the third and fourth instars. Duration of instars 1–4 was 1, 1, 2, and 3 days, respectively, when cultured at 32°C (O’Neal and Markin, 1975b). Porter (1988) reported that the duration of the larval stage decreased from 28 to 11 days as temperature increased from 24°C to 35°C. The castes are quite similar in appearance during the larval stage; the principle difference

is the size. Mean size is greatest among larvae destined to be sexual forms, followed by major, minor, and minim workers. Development time is proportional to size, with the larger reproductives requiring nearly twice as much as the minim workers, and the other castes intermediate. Pupa. Pupal development is often reported to require about 6–8 days, though Porter (1988) indicated that the duration of the pupal stage decreased from 28 days when cultured at 21°C to less than 7 days at 35°C. The pupa greatly resembles the adult in form, though it is whitish. Size varies considerably, depending on the caste of the adult form. The pupae, along with the eggs and larvae, comprise the ant brood and are moved about within the colony by workers according to environmental conditions.

FIG.  9.3  Red imported fire ants, including males and workers. (Photo by L. Buss.)

Adult. A total of 20–30 days is normally required for egg d­ eposition and emergence of the first mature worker ants, which is called minim workers. Within 90 days of colony founding, the colony may consist of 200 minims and a few minor workers. Porter and Tschinkel (1986) described the importance of minims in colony founding and success. Within 5  months, there may be 1000 minor and a few major workers. Within 7 months, the colony may consist of 6000–14,000 workers and may contain about 3% major workers. The number of ants per colony is estimated at 11,000, 30,000, and 60,000 workers at 1, 1.5, and 3 years, respectively. After 3 years, the colony is considered to be matured, and the number of workers is about 230,000 per colony. Abundance typically decreases on each winter, however, and reproduction ceases at northern latitudes and decreases on more southern locations. Colonies may begin to produce reproductives at 5–7 months, but normally it is older colonies that produce most reproductives. Adult worker ants vary considerably in size, ranging from about 1.6–6 mm long. They lack wings and vary in color from light reddish brown to dark brown. The gaster is sometimes marked with an orange spot. The assignment of workers to

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castes is based on arbitrary head width categories, not on distinct morphological or behavioral differences. Adult reproductives bear two pairs of wings, with the front wings longer than the hind wings. The females resemble the workers in color, but the males are black except for the pale antennae.

FIG. 9.4  Red imported fire ant nest. (Photo by L. Buss.)

The physical structure of the nest changes markedly over time. As noted earlier, initially it is a small burrow with chambers. As workers are produced, however, the tunnels are enlarged, and additional chambers are produced. Within 90  days of colony founding, a soil surface mound is produced, typically 5–7 cm in height and 3–7 cm in diameter. At this time, there are 5–15 vertical tunnels extending a meter or more into the soil down to the water table. Mounds in areas with sandy soil tend to be relatively flat, whereas in clay soil they can be 0.5–1.0 m in height. Large c­ olonies may construct several mounds. They may continue to use the same mound for several years or may construct new ones. The mound enables the colony to optimize environmental conditions for the brood. In cool weather, the brood is moved to the sunny side of the mound near the surface, while in hot weather, the brood is found deep underground tunnels.

Food for the colony is collected by foraging workers, which often forage for considerable distances from the colony, with the distance dependent on colony food requirements. Maximum foraging occurs between the soil temperature (5 cm depth) of 21–35°C, but some foraging can occur between 10°C and 37°C. Foraging is aided by the construction of tunnels leading back to the nest; these often extend 15–25 m from the mound. Foraging ants search randomly for food, but once the food is located, ants returning to the tunnels deposit trail pheromones that serve to recruit other ants. Soon there is a stream of ants leading to the food. Food obtained by the foraging workers is quickly distributed throughout the colony. Food typically passes from foragers to nurses, and then to larvae and queens. Colony organization and coordination are maintained by secretion of pheromones. Several pheromones associated with fire ants have been identified, including the brood, trail, queen tending, and nestmate recognition pheromones. There are two types of colonies: single queen (monogyne) and multiple queen (polygyne). The monogyne colonies contain about 100,000–240,000 workers, fight with other colonies, and tend to be situated farther apart, with mound densities of 100–350 per hectare. In contrast, polygyne colonies contain 20–60 queens and 100,000–500,000 workers, do not fight with other polygyne colonies and have densities of 500–2000 mounds per hectare. Polygyne colonies tend to erect lower mounds than monogyne colonies and certain smaller workers which are lighter in color. Polygyne colonies are becoming more common in the southern United States. The fire ants were described by Buren (1972). Hung et  al. (1977) provided keys to the common species of fire ants. Creighton (1950) presented a comprehensive treatment of North American ants, though red imported fire ant was not distinguished in this treatment. Wheeler and Wheeler (1990) provided a good key to the North American genera. Biology of fire ant was reviewed by Lofgren et al. (1975), Vinson and Greenberg (1986), and Vinson (1997). Vander Meer (1988) discusses the caste structure. A laboratory rearing method was described by Williams et  al. (1980). Bibliographies were published by Banks et  al. (1978), Wojcik and Lofgren (1982), and Wojcik (1986b). Zoogeography was summarized by Buren et al. (1974).

Damage

FIG. 9.5  Red imported fire ants feeding on eggplant. (Photo by J. Castner.)

Red imported fire ant thrives in disturbed habitats such as cropland and pastureland. Thus, agricultural environments are particularly likely to be infested, but similar sites such as roadsides, irrigation ditches, and athletic fields also support high densities. Red imported fire ant does not survive well in climax plant communities, particularly if the dense shade is provided by trees and shrubs.

Order Hymenoptera—Ants and Sawflies Chapter | 9  387

An interesting feature of these ants is that they are unable to ingest solid food. They place solid food near the mouth of fourth instar larvae, which then secret digestive enzymes. The liquefied food is then passed around the colony. Nevertheless, fire ants readily collect solid materials, including plant tissue. Young seedlings, in particular, can be killed by fire ants. For example, Adams (1983) reported the destruction of over 50% of eggplant seedlings in Florida. The damage was in the form of stem girdling and destruction of the growing point of the young plants. Seedling injury is the most frequent form of damage, but ants also feed belowground on seeds, roots, and tubers; they remove bark from young citrus trees and cause abortion of okra flowers by feeding at the base. Root tissue seems to be damaged more commonly than is generally recognized. Plant injury occurs most commonly when ants are deprived of food, as when land is first cultivated. Fire ants are not normally considered to be primary plant pests except for okra, where the damage can be severe and frequent. However, they affect crop production indirectly because their mounds interfere with equipment operations, and laborers are reluctant to harvest heavily infested crops due to the toxic sting of fire ants. Fire ants are well known for their venomous sting, which induces a burning sensation in victims—in fact, this is the basis of their common name. Although the individual sting is much less toxic than that of bees and many other stinging insects, the abundance of the ants and the tendency of victims to receive multiple stings represent a real threat to human health. Humans are not the only animals at risk. Reptiles, ground-nesting birds, young mammals, and even fish have been reported to be killed by fire ants. Animals reproducing during the warmest periods of the year are at great risk because ant foraging is reduced in cool weather. Rates of reproduction of several forms of wildlife reportedly increase the following area-wide suppression of fire ants. This is also of interest to livestock producers because newborn livestock is at risk; blinding of young calves is the most common form of livestock injury. The impact of fire ants on the plant, animal, and human health was reviewed by Adams (1986), Lofgren (1986), Jemal and Hugh-Jones (1993), and Allen et al. (1994, 2004). Damage is somewhat offset by the beneficial predatory behavior of red imported fire ant. Predation of fire ants on sugarcane borer, Diatraea saccharalis (Fabricius) (Lepidoptera: Pyralidae), boll weevil, Anthonomus grandis Boheman (Coleoptera: Curculionidae), and lone star tick, Amblyomma americanum (Linnaeus) (Acari: Ixodidae), is particularly well documented. However, predation occurs on many species of insects, particularly caterpillar larvae on plants and fly larvae breeding in animal manure, and red imported fire ant is usually considered to be a beneficial insect within the context of cotton and sugarcane ­production systems. Reagan (1986) reviewed the beneficial aspects of fire ants.

Management Sampling. Fire ants and their mounds are easily detected, and the visual examination of fields is adequate for most purposes. Density is usually expressed as number of mounds per unit of land area, though colony size is an important variable. Insecticides. Insecticides are effective for ant suppression, but reinvasion of treated areas from untreated areas can occur, sometimes giving the impression that insecticides were not effective. Ants are often treated by applying toxin-treated bait. The attractive component of the bait is normally soybean oil, and the granular carrier can be any of several relatively inert substances, often corncob grit. However, the acceptance by fire ants of baits varies seasonally, with high protein baits preferred in the warm months and high carbohydrate baits accepted more readily in the cool months (Stein et al., 1990). The toxin added to the bait formulation is usually a slow-acting insecticide or an insect growth regulator. Slow induction of mortality is advantageous because it allows the toxin to be transported through the colony to the brood and perhaps even to the queens. Even just treatment of field perimeters can be beneficial, as foraging ants may collect bait deposited in perimeter areas and return it to colonies located within crop fields. An alternative method of suppression is the application of granular, dust, or liquid insecticide to mounds, or to tilled soil, but mortality is often less complete than with bait formulations. Ants from mounds treated with insecticide may relocate following treatment, necessitating one or more reapplications of insecticide before the colony is eliminated. Insecticide applications can be integrated with biological control agents. Oi et al. (2008) reported that when sites receiving insecticide treatment were accompanied by the release of biological control agents [the fungus Kneallhazia (Thelohania) solenopsae and the parasitoid Pseudacteon tricuspis Borgmeier], suppression of fire ants was greatly enhanced Cultural Practices. There are few effective nonchemical approaches to managing fire ants. Suppression of other insects is helpful because it deprives the principal food supply of ant. Individual mounds can be treated with boiling water, but at least 10 L of water are necessary to penetrate most mounds, and its effectiveness is limited. Boiling water also kills nearby plants. Physical destruction of mounds by digging or tilling and shoveling two mounds together in hopes of stimulating a fatal fight are of little value. Large individual plants can be protected from foraging ants by placing a wide barrier of adhesive around the base of the plant. This approach is often best preceded by wrapping the trunk or stem with tape or foil to prevent the adhesive from disfiguring the plant or damaging the plant tissue. Biological Control. The diseases and parasites discussed under “natural enemies” are being inoculated for the permanent establishment and are not generally available

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from commercial sources, but other organisms have been suggested for biological suppression if applied regularly. In particular, entomopathogenic nematodes (Nematoda: Steinernematidae and Heterorhabditidae) and the straw itch mite, Pyemotes tritici (Lagrexe-Fossat and Montane) have been suggested for biological suppression. However, the performance of these biotic agents has been disappointed

under field conditions. When fire ant moves to a new area, its natural enemies, particularly the microbial pathogens, tend to get left behind. The absence of these microbes may indicate strong selective pressure against founders due to fitness costs associated with harboring the microbes (Yang et al., 2010a, b, c). Thus, it may be necessary to inoculate new areas of invasion with the microbial pathogens.

Chapter 10

Order Lepidoptera—Caterpillars, Moths, and Butterflies

FAMILY ACROLEPIIDAE—FALSE DIAMONDBACK MOTHS Leek Moth

Acrolepiopsis assectella (Zeller) (Lepidoptera: Acrolepiidae)

Natural History Distribution. Leek moth is found throughout western Europe and parts of Asia, and was first observed in North America in Ontario, Canada in 1993. It has since spread to most of southeastern Canada, and to New York, but is expected to spread eventually to most of the eastern United States, the coastal regions of the Pacific Northwest and southern British Columbia, and north-central Mexico (Mason et al., 2011). Host Plants. The principal crop hosts are garlic, leek, and onion (all Alliaceae), though chives and shallots also are injured. In Canada, the greatest damage occurs to leek and onion. Allison et al. (2007) studied susceptibility of cultivated Allium spp. as well as some native and cultivated plants related to Allium. Wild leek, A. tricoccum Ait., was as attractive to ovipositing flies as was cultivated leek, A. ampeloprasum Linnaeus, and garlic, A. sativum Linnaeus. Nodding onion, A. cernuum Roth (Liliaceae), and prairie onion, A. stellatum Fraser (Liliaceae), were nearly attractive. Some eggs were also deposited on daffodil, Narcissum pseudonarcissus (Linnaeus) (Amaryllidaceae), and iris, Iris versicolor Linnaeus (Iridaceae), which are related to Allium, but are in different families. However, larvae survived to the third instar only on Allium spp. Overall, preference and survival were higher on plants closely related to Allium. However, larval survival was variable even within the Allium spp., with nodding onion displaying significantly lower survival than the other Allium spp. Oviposition behavior is somewhat variable, with females more accepting Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00010-8 © 2020 Elsevier Inc. All rights reserved.

of less-preferred plants in the absence of preferred hosts. There are many potential wild hosts, but most have not been evaluated for susceptibility. Natural Enemies. A number of generalist wasp (Hymenoptera) parasitoids have been found to be associated with leek moth in Canada: Itoplectis conquistador (Say), Scambus hispae (Harris), S. pterophori (Ashmead) (all Ichneumonidae), Bracon furtivus Fyles (Braconidae), and Conura albifrons (Walsh) (Chalcididae). However, they have not been very effective. In Switzerland, however, high levels of mortality were observed in leek moth populations, especially among newly hatched larvae (Jenner et al., 2010). However, the authors concluded that the pupa might be the best target for biological control because the mortality of this stage was most variable. Efforts are being undertaken to release a more effective parasitoid from Europe, namely the pupal parasitoid Diadromus pulchellus Wesmael (Hymenoptera: Ichneumonidae) (Mason et  al., 2010). Garcia del Pino and Morton (2007) evaluated the nematode Steinernema feltiae (Steinernematidae) and the bacterium Bacillus thuringiensis for suppression of leek moth larvae under field conditions and reported that the nematode was quite effective, and considerably more effective than B. thuringiensis. Life Cycle and Description. In northern areas of Europe there are one to two generations per year, but the number increases in warmer climates, from three to five in France, and five to six in southern Italy. A similar pattern is expected in North America, and in Ontario there are three flights of adults and two or three generations per year. Should this insect spread throughout North America, six or more generations are predicted annually in southern areas. The flight periods in Ontario are April-May for the overwintered population of moths, June-July (first complete generation), and July-August (second complete generation) (Mason et  al., 2011). Egg deposition by the 389

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moths peaks toward the end of these periods. Thus, the first complete generation in southern Canada and northern United States occur principally from May to June, and the second from July to August. Duration of the life cycle is 3–6  weeks in Canada. Adults undergo reproductive diapause when the developing larvae experience day lengths shorter than 15 h. Adults overwinter in plant debris, either in or near host crop fields. Mason et al. (2010) estimated the developmental threshold at 6–7°C, and the degreeday requirement for the egg to adult stage to be 445 daydegrees above the developmental threshold. Similarly, Åsman (2001) estimated the developmental threshold at 6°C and the degree-day requirement for a complete generation to be 630  day-degrees above the developmental threshold. Egg. The small, translucent eggs of this insect are deposited on the underside of leaves. They are oval in shape, and iridescent white in color. Eggs are about 0.5 mm long and 0.2 mm wide. Eggs are deposited 10–14  days after emergence of the moths, which is also 2–6 days after mating. Females deposit about 100 eggs singly at the base of host plants. Hatching occurs in 5–8 days and larvae enter the leaves within 24 hours. Larva. The larva is creamy white in color, measuring about 13–14 mm in length at maturity. The raised structures on the body are brown. The prothoracic and anal plates are also brown. The head is yellowish brown, sometimes marked with a network of reddish-brown lines. The larval period is 15–20 days, and there are five instars. The head capsule widths are 0.16, 0.24, 0.36, 0.52–0.56, and 0.68–0.84 mm in instars 1–5, respectively (Mason et al., 2010). These same authors also provided development rates at several temperatures, and reported that at 25°C the mean (± SE) development period was 6.0 ± 0.07, 2.8 ± 0.12, 2.6 ± 0.13, 2.9 ± 0.14, 3.2 ± 0.14, and 3.6 ± 0.31 days for the egg and larval instars 1–5 to complete development. Also, 3.7 ± 0.19, 7.6 ± 0.22, and 24.9 ± 0.48 days was required for the prepupal, pupal, and egg to adult periods. Pupa. The reddish-brown pupa is enclosed in a loose, white, net-like structure. It is about 7 mm long. It is normally found attached to leaves or other structures near the host plant. Adult. The front wings of the moth are mostly brown but speckled with darker brown, black, and white. At about the midpoint of the trailing edge of the forewing is a larger, triangular white spot. The hind wings are grayish brown. A fringe of hairs is found on the front and hind wings. The head and thorax are dark brown, the abdomen grayish brown. With the wings folded, it measures only about 1 cm long, and the wingspan is about 1.5 cm. The zigzag flight

activity of leek moth is limited to evening periods, though mating occurs in the morning. Adults normally live for up to 2 months, though during the winter they persist for several months.

Damage This leaf-mining insect feeds on both the leaves and the bulb. They also attack the flower stalks and inflorescences of hosts. The larva initially is a leafminer, feeding inside the hollow leaves of Allium and creating a 2–5 mm gallery in the leaf. Feeding inside the leaf, larvae often leave the outer epidermis in place, creating a ‘window-pane’ effect. Larger larvae also create holes in the leaves. Young leaves are preferred and eventually, the larva works its way to the center of the plant to feed on the inner (youngest) leaves. Host plants that are harvested with the leaf attached, such as leeks and chives, are particularly susceptible to injury. Feeding reduces the growth potential of the plant. Although this insect is not a vector of plant pathogens, feeding damage may allow entry of plant pathogens into the bulb. Damage tends to be greater at the edges of crop fields, and in later generations.

Management Sampling. Females produce a sex pheromone to attract males. The pheromone is known (Minks et al., 1994), and commercially available. Due to the small size and nocturnal nature of the moth, pheromone sampling is preferred. Traps should be placed at about 1 m high at the edges of crop fields. Insecticides. Because this moth can be very damaging, growers in areas where A. assectella occur often apply insecticides. In Germany, good control has been accomplished by commencing insecticide application when 10% of the plants have become infested. Alternatively, insecticide can be applied about 3 weeks after the beginning of the egg deposition period. The latter approach resulted in slightly less damage than when the 10% action threshold was used for timing of insecticide application (Richter and Hommes, 2003). Although a number of chemical and biologically based insecticides can provide suppression, Bacillus thuringiensis and azadirachtin products seem not to be effective (Olmstead and Shelton, 2012). Cultural Practices. A net covering can be used to prevent moths from ovipositing on susceptible Allium crops (Richter and Hommes, 2003). However, this technology is labor-intensive, and therefore expensive. Intercropping with red clover or cabbage did not affect egg deposition by leek moth (Åsman, 2001). Leek moth did not favor either leek or chives for oviposition, but larger Allium plants are more preferred than small and can function as a trap crop by diverting eggs from the main crop (Åsman, 2002).

Order Lepidoptera—Caterpillars, Moths, and Butterflies Chapter | 10  391

FAMILY BEDELLIDAE—BEDELLID MOTHS Morningglory Leafminer

Bedellia somnulentella (Zeller)

Sweetpotato Leafminer

Bedellia orchilella Walsingham (Lepidoptera: Lyonetiidae)

Natural History Distribution. Morningglory leafminer apparently is a native to Eurasia, but now it is widely distributed in the United States. Morningglory leafminer likely occurs in southern Canada but it is not recorded as a pest there. It is found in most areas of the world, including both tropical and temperate climates. Sweetpotato leafminer is associated with sweet potato in Hawaii, but its origin is uncertain, and it does not occur elsewhere in the United States or Canada. Host Plants. Larvae of both species feed on plants in the family Convolvulaceae. Among vegetables, only sweet potato is attacked. Wild hosts include bindweed, Convolvulus spp., and morningglory, Ipomoea spp. Natural Enemies. Morningglory leafminer is frequently parasitized by Apanteles bedelliae Viereck (Hymenoptera: Braconidae). Also reared from morningglory leafminer is Spilochalcis albifrons Walsh (Hymenoptera: Chalcididae), but there is some disagreement whether this wasp is a primary parasite or hyperparasite. A parasitoid identified as Omphale metallicus Ashmead (Hymenoptera: Eulophidae) attacks sweetpotato leafminer. In Hawaii this parasitoid is thought to be a significant mortality factor; nevertheless, leafminer can be a serious problem in the absence of insecticides. Apanteles bedelliae was introduced to Hawaii to aid in the suppression of sweetpotato leafminer, and is reported to be effective (Zimmerman, 1978). Life Cycle and Description. Although these species are similar, they are treated separately because there are some differences in their biology. In morningglory leafminer, total development time, from the egg to the adult stage, requires about 30 days at 18°C, but only 16–17  days at 29°C. Two generations are reported annually in New York. Egg. The eggs of morningglory leafminer are described as translucent white and a flattened sphere in shape. They measure about 0.3 mm long and 0.2 mm wide. Eggs generally are deposited on the lower surface of leaves, and usually adjacent to leaf veins. They are placed singly or in small clusters of 2–3 eggs. Embryonic development requires about 4.5 days at 27°C.

FIG.  10.1  Morningglory leafminer larva burrowing from leaf tissue. (Photo by L. Buss.)

Larva. Upon hatching, larvae of morningglory leafminer burrow directly into the leaf tissue. They are yellowish gray though they turn greenish with age. Larvae develop dorsal pink spots during the third instar which disappear in the fourth instar and are replaced by reddish tubercles in the fourth and fifth instars. White tubercles are added during the fifth instar. First instar larvae appear to be legless, but both thoracic legs and prolegs are evident in subsequent instars. Head capsule widths are 0.09, 0.15, 0.22, 0.31, and 0.45 mm in instars 1–5, respectively. Larvae are somewhat flattened in appearance and the anal prolegs project posteriorly, forming a fork. The first two instars form serpentine mines, but young third instars leave the mine, construct a loose webbing of silk on the lower surface of the leaf, and reenter the leaf tissue to form a blotch-like mine. The remaining instars also function as blotch miners. An interesting feature of the feeding behavior is that except while molting the larvae use the silk as a point of attachment while feeding, insert the anterior portion of their body into the mine but leave the posterior portion protruding from the mine. At this point, the larva voids its fecal material externally. The larva usually forms several blotch mines during its development. Fecal material is voided from the blotch and usually hangs down in a continuously webbed chain. Larval development time is about 11 days at 27°C.

FIG. 10.2  Morningglory leafminer pupa. (Photo by L. Buss.)

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Pupa. Newly formed pupae are greenish with a mottledred pattern but the red gradually fades and the pupae become greenish or brownish. The posterior end of the pupa is anchored to silk webbing. The anterior end is also supported by silk threads, however, so the pupa is positioned parallel to the leaf rather than hanging from the substrate by its anal end. Duration of the pupal stage is about 4.5 days at 27°C. Adult. Adults of morningglory leafminer are grayish brown and with fringed wings. The wingspan is about 11.5 mm. Females begin ovipositing soon after mating and ­deposit about 170 eggs (range 80–324 eggs). Most eggs are laid about 1–8 days after oviposition commences, at about 20 eggs per day. The longevity of adults is about 25 days. Sweetpotato leafminer is a relatively unimportant pest on a minor crop. Thus, it is perhaps not surprising that little about this insect is known, and that phenology is undocumented. As is the case with morningglory leafminer, a complete generation requires about 30 days. In Hawaii, it likely occurs whenever its host is available. Egg. The eggs of sweetpotato leafminer are laid singly in a crevice along a vein, usually on the lower surface of a leaf. The eggs are oval and flattened, and measure 0.3 × 0.2 mm in size. The eggs are whitish to reddish in color and highly iridescent. Duration of the egg stage is about 8 days. Larva. The larval stage of sweetpotato leafminer is small, growing from about 0.4 mm to 7.0 mm long as it matures. The larva tunnels between the upper and lower leaf surfaces as it feeds. The tunnels are quite long and narrow, unlike morningglory leafminer where the mine becomes blotch-like as the larva reaches maturity. As the food supply is exhausted the larva leaves its tunnel for a new feeding position and creates a new tunnel. When not in the tunnel the larva moves with a looping motion. The caterpillar is greenish throughout its development, and unlike many Lepidoptera, the head, prothoracic, and anal shield are not darkly pigmented, though they may have a brownish tint. Moderately sized hairs are sparsely distributed on the body. Duration of the larval stage is about 10 days. Pupa. When the sweetpotato leafminer larva matures it leaves the mine and spins a few silken strands which are used to anchor the pupa to the outside of the leaf. The pupa is dark-green when first formed, assuming a brown color with black spots as it matures. The head bears a dark angular projection, and the eyes are black. The pupa measures about 3.5 mm long. Duration of the pupal stage is about 6 days. Adult. The moth of sweetpotato leafminer is grayish brown and lacks distinctive markings. The wings, however, are unusual in form. The forewing tapers to a point distally,

and the trailing edge bears a long fringe. The hind wing is very narrow but bears a fringe of long hairs on all sides. The wingspan of the moth is about 7.5 mm. Morningglory leafminer was described by Shorey and Anderson (1960) and Parrella and Kok (1977). Biology of this species in Egypt seems to be nearly identical (Tawfik et  al., 1976). Clemens (1862) gave a good description of the larvae. Biology of sweetpotato leafminer was described by Fullaway (1911); Zimmerman (1978) added useful observations.

Damage The larvae of both species mine the foliage of sweet potato, leaving long-winding tunnels containing fecal material. As noted earlier, the larva of morningglory leafminer changes its behavior and forms blotch mines later in life. If infestations are heavy the leaves acquire a withered or seared appearance. Sweet potato is quite tolerant of foliar injury, so leafminers are not usually considered to be a serious pest. Parrella and Kok (1977) suggested that hot weather was unfavorable for survival of morningglory leafminer, which might explain why it rarely was reported to be damaging.

Management Foliar insecticides, particularly systemic materials, can be applied for leafminer suppression. Except perhaps in Hawaii, these species are rarely abundant, and insecticide applications are used sparingly so as not to disrupt naturally occurring parasitoids.

FAMILY DEPRESSARIIDAE— DEPRESSARIID MOTHS Parsnip Webworm

Depressaria radiella (Goeze) (Lepidoptera: Depressariidae)

Natural History Distribution. Parsnip webworm was originally known from Europe and apparently was introduced to North America some time before 1869. Its distribution now includes southern Canada from Nova Scotia to British Columbia and the northern United States south to Maryland and Arizona. Host Plants. In addition to feeding on parsnip, this insect feeds on several umbelliferous weeds, including cow parsnip, Heracleum lanatum; and angelica, Angelica spp. Records of wild carrot, Daucus carota, are doubtful. Natural Enemies. Parasitism varied from 0% to 100% in Iowa (Gorder and Mertins, 1984), but was limited to the larval stage. Apanteles depressariae Muesebeck (Hymenoptera: Braconidae) is the principal parasitoid, though others have been collected on occasion.

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Life Cycle and Description. There seems to be only a single generation per year, though in Iowa a few eggs are deposited in late summer, suggesting the possibility of a partial second generation. Egg to adult development times is reported to be 38 days in Iowa and 62 days in Nova Scotia.

Pupa. Pupation occurs within a silken sheath, generally within the stalk of the food plant. The pupa is brown. Duration of the pupal stage is about 13 days.

Egg. The eggs are deposited principally on foliage, and to a lesser extent on flower stalks. They are elliptical but slightly rectangular. They are white and ribbed longitudinally. Egg length averages about 0.56 mm (range 0.36–1.14 mm). Duration of the egg stage is about 4 days. FIG. 10.4  Parsnip webworm drawing. (Drawing by USDA.)

Adult. The adults fly at night but are not attracted to lights. The moths are fairly large for this group of moths, with wing lengths of 9.5–13.0 mm. The front wings are yellowish brown, marked with gray, whereas the hind wings are grayish. Overwintering occurs in the adult stage, with the adults in reproductive diapause. Egg production under laboratory conditions averages 470 eggs per female (range 170–830 eggs). Females oviposit on umbels or leaf surfaces in May and June. Females deposit more eggs on larger plants (Zangerl and Berenbaum, 1992). The most complete account of parsnip webworm biology was given by Gorder and Mertins (1984). Methods for rearing were developed by Nitao and Berenbaum (1988), who also provided data on developmental biology. Brittain and Gooderham (1916) gave a good morphological description of this insect, except that some of the measurements are incorrect. FIG. 10.3  Parsnip webworm larva. (Drawing by USDA.)

Larva. Upon hatching the young larvae bore into the blossom to feed. They prefer unexpanded flowers and generally distribute themselves so that there is only one insect per flower head (Thompson and Price, 1977). The larval color is generally greenish yellow dorsally with yellow laterally and ventrally but sometimes tends toward blue gray. The head is black, and the length of the body is well-marked with rows of raised black spots. There are six larval instars. Head capsule widths are about 0.1, 0.3, 0.5, 0.7, 1.0, and 1.6 mm, respectively for instars 1–6. Larval development requires about 21 days, with the duration of the instars about 3, 2, 3, 4, 3, and 8 days, respectively. The larva, at maturity, attains a length of 16–18 mm. Larvae rest within the flower or other sheltered locations on the plant, surrounded by a tunnel webbed from silk. When disturbed, the larvae retreat within the tunnel, and if pursued wriggle violently and drop to the soil. The ability of larvae to detoxify furanocoumarins, which prevents many insects from attacking parsnip, is a characteristic of this herbivore. However, larvae feed preferentially on plants low in furanocoumarins (Zangerl and Berenbaum, 1993).

Damage Parsnip webworm feeds primarily on the flowers and seeds of parsnip, and is of concern only where seed production is desired. The larvae web together with the flower heads and feed within, including the seeds, leaving the flower structure a mass of webbing and fecal material.

Management This insect should rarely warrant control efforts, but foliar insecticides applied before the opening of blossoms can prevent larval feeding.

FAMILY CRAMBIDAE—BORERS, BUDWORMS, LEAFTIERS, WEBWORMS, AND SNOUT MOTHS Alfalfa Webworm

Loxostege cereralis (Zeller) (Lepidoptera: Crambidae)

Natural History Distribution. This native insect is primarily western in distribution, occurring from British Columbia to Quebec in

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the north and from California to western Texas in the south. It is highly dispersive, however, and sometimes attains the northeastern states. As a pest, its range is limited to the western Great Plains and Rocky Mountain region. Host Plants. Alfalfa webworm has a broad host range, very similar to that of the better-known beet webworm, Loxostege sticticalis (Linnaeus), with which it is sometimes confused. Its common name is misleading because though females oviposit in alfalfa and larvae consume alfalfa foliage, it is not routinely associated with this crop. In general, plants in the family Chenopodiaceae are preferred. However, when this insect is abundant and the favored plants are no longer available, a wide range of plants can be injured. Among vegetables susceptible to injury are bean, beet, cabbage, cantaloupe, carrot, eggplant, lettuce, parsnip, pea, pepper, spinach, and likely many others. Grasses normally are avoided, but corn can be fed upon if nothing else is available. Alfalfa and sugarbeet are the field crops most often damaged during periods of great abundance. Among the weeds fed upon by alfalfa webworm, at least during periods of abundance, are dock, Rumex sp.; lambsquarters, Chenopodium album; common mallow, Malva neglecta; ragweed, Ambrosia sp.; redroot pigweed, Amaranthus retroflexus; saltbush, Atriplex sp.; Russian thistle, Salsola kali; sunflower, Helianthus annuus; and sweetclover, Melilotus sp. (Hoerner, 1933). In moth oviposition and larval food preference tests, Capinera et  al. (1981b) reported that kochia, Kochia scoparia, and Russian thistle were favored for oviposition, though some eggs were deposited on all weeds tested, whereas only lambsquarters, redroot pigweed, and shepherdspurse, Capsella bursa-pastoris, were fed upon. Maxson (1948) reported finding eggs on dandelion, Taraxacum officinale; field bindweed, Convolvulus arvensis; kochia; lambsquarters; plantain, Plantago major; purslane, Portulaca oleracea; nightshade, Solanum sp.; ragweed, Ambrosia sp.; redroot pigweed; and saltbush. Adults feed on nectar from dandelion and field bindweed. Natural Enemies. Several natural enemies are described, though none are known to be particularly effective. In Colorado, for example, Hoerner (1933) reported that only 3% of the larvae were parasitized. Among the parasitoids are Cremnops vulgaris (Cresson), Meteorus campestris Viereck (both Hymenoptera: Braconidae), Aplomya caesar (Aldrich), A. trisetosa (Coquillett), Lespesia archippivora (Riley), and Phryxe vulgaris (Fallén) (all Diptera: Tachinidae). Life Cycle and Description. In Canada, two to three generations are reported annually, but in the United States three generations seem to be normal. A generation can be completed in about 40  days under optimal conditions. As with beet webworm, mature larvae overwinter and the proportion of each generation that diapauses increases as the summer progresses. In Colorado, the first flight of moths occurs in May, followed by additional flights in June-July

and September. However, Capinera et al. (1981a) suggested that the first two flights might both result from protracted emergence of overwintering larvae. Egg. Moths begin oviposition within a few days of emergence and continue to oviposit for about 2 weeks. The flattened, oval eggs are about 1 mm long and 0.7 mm wide. They are white initially, but eventually, become yellow. They usually are deposited in clusters of 2–20 overlapping eggs on the underside of foliage. Although individual eggs greatly resemble those of beet webworm, the clustering arrangement of alfalfa webworm serves to distinguish them from the single-row oviposition pattern of beet webworm. Eggs of alfalfa webworm hatch in 4–6  days during warm weather.

FIG. 10.5  Alfafa webworm larva. (Photo by J. Capinera.)

Larva. The larval stage has six instars. Larvae increase in length from about 3 mm at hatching to about 25 mm at maturity. Mean head capsule widths are 0.24, 0.41, 0.64, 1.01, 1.36, and 1.75 mm during instars 1–6, respectively. Development time of larvae fed sugarbeet foliage and reared at 27°C was 2.7, 1.8, 1.9, 2.3, 2.4, and 5.5 days, respectively, for instars 1–6. The optimal temperature for larval development is about 30°C, and larvae fail to develop at 15°C. Young larvae are pale yellowish green, but they become darker as they mature. The head and thoracic plate are dark during instars 1–3, becoming irregularly colored with light and dark blotches thereafter. During the final instar, the larva bears a broad whitish dorsal line along its body, bordered by broad black dorsolateral stripes, a distinct contrast with the dark dorsal line of beet webworm. Most abdominal segments bear six dark spots with light centers dorsally, with a single dark hair arising from each. Larvae sometimes spin a mat of silk on foliage, where they rest during the daylight hours. They also produce strands of silk which are used to web together foliage and to construct a long thin tube into which they may retreat if disturbed. The silk tube often connects to clods of soil or other protected retreats. Duration of the larval stage is about 17  days at 30°C, increasing to about 25  days at 25°C and 35°C. Duration of

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the larval stage is extended to several months for larvae that enter diapause. During diapause they remain below-ground within the silken pupation cell and then pupate in the spring. Pupa. Larvae enter the soil to a depth of up to 2.5 cm when they are ready to pupate, where they construct a silklined cell. The pupa is yellowish initially, turns dark brown with age, and measures about 25–30 mm long. The posterior, pointed end of the pupa bears eight small spoon-shaped appendages. This character serves to distinguish alfalfa webworm from beet webworm, which bears eight spines instead. Duration of the pupal stage is often 14–21 days under field conditions but requires only 8–9 days when reared at a constant temperature of 25–35°C.

FIG. 10.6  Alfalfa webworm adult. (Photo by J. Capinera.)

Adult. The moth is fairly small, measuring only about 2.5–3.0 cm in wingspan. It is grayish brown in general color, though containing some irregular black marks and a transverse cream-colored band on the distal margin of the forewings. Adults are attracted to light, and during periods of abundance tremendous numbers aggregate around light sources. Many observers have described the appearance of the moth flights as being equivalent to a blinding snowstorm. Alfalfa webworm moths are often confused with adults of beet webworm, Loxostege sticticalis (Linnaeus). However, they are easily differentiated by viewing the underside of the wings. Both species have a narrow dark line along the distal edge of the wings, but the line is complete in beet webworm, whereas it is broken in alfalfa webworm. The biology of alfalfa webworm was given by Hoerner (1933), Maxson (1948), and Capinera et al. (1981a, 1981b). Keys to the moths were included by Munroe (1976) and Capinera and Schaefer (1983). The larva was included in the key to the genus, Losostege, by Allyson (1976), and in the field key by Capinera (1986); Capinera et  al. (1981a) pictured the instars.

Damage Larvae injure plants by consuming foliage. Young larvae skeletonize leaves, but older larvae consume leaf tissue completely, often leaving only the stems and large veins. They often web together leaves and feed within the clustered foliage. Damage potential varies greatly among crops. For example, larvae consume over 50 cm2 of sugarbeet foliage during their development, but only about 13 cm2 of alfalfa foliage (Capinera et al., 1981b).

Management Sampling. The eggs and larvae can be found by visual examination of plants, but both of these life stages are difficult to detect. Moths are attracted to light and can be captured in light traps. Adults also can be flushed during the day, especially toward evening. Insecticides. Alfalfa webworm populations are usually suppressed by application of insecticide to foliage. Bacillus thuringiensis affords some control. Populations are infrequently damaging, so insecticides should not be applied unless high densities of adults or larvae are observed. Cultural Practices. Several practices can alleviate webworm damage. Tillage can disrupt and destroy overwintering larvae within their silken tubes in the soil. Destruction of preferred weeds before adult oviposition flights can minimize the deposition of eggs within crops. Destruction of weeds after eggs hatching, however, tends to drive larvae to nearby crop plants. Construction of deep furrows with steep or slippery sides has sometimes been recommended to stop the advance of dispersing larvae.

Beet Webworm

Loxostege sticticalis (Linnaeus) (Lepidoptera: Crambidae)

Natural History Distribution. Beet webworm is found in the northern regions of Europe and Asia, and apparently is an immigrant from Europe. In the United States and Canada, beet webworm is present from coast to coast but is a pest principally in western sugarbeet-growing areas from Alberta and Manitoba in the north to Utah and Kansas in the south (Pepper, 1938). Host Plants. Known principally as a pest of sugarbeet, this insect also feeds readily on table beet and chard. During periods of abundance over 80 species of plants are damaged, but infestation is normally limited to Beta vulgaris and to certain weeds. Among the vegetables that are occasionally injured are cabbage, cantaloupe, carrot, cucumber, garlic, lettuce, mustard, onion, pea, potato, pumpkin, rhubarb, spinach, and turnip. Grasses, including corn, are rarely eaten. Weeds readily consumed include l­ambsquarters, Chenopodium album; redroot pigweed, Amaranthus

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­retroflexus; and Russian thistle, Salsola kali. These weeds commonly serve as the preferred oviposition site of moths, with larvae dispersing to other, less preferred plants when the weeds perish or are consumed. Larvae have been reared successfully on such diverse flora as alfalfa; onion; lambsquarters; sagebrush, Artemesia sp.; sunflower, Helianthus annuus; and Canada thistle, Cirsium arvense (Pepper and Hastings, 1941). Natural Enemies. Several parasitoids are known from beet webworm in North America. Numerous other species have been identified in Europe and Asia, but none have been imported. Wasps seem to be the most important mortality agent, though this has been little studied. Among the fly, parasitoids are Aplomya caesar (Aldrich), Euphorcera omissa (Reinhard), Lespesia archippivora (Riley), L. ciliata (Macquart), and Stomatomyia parvipalpis (Wulp) (all Diptera: Tachinidae). A fungal disease, Tubulinosema loxostegi (Microsporidia: Tubulinosematidae) has only recently been described from Siberia (Malysh et al., 2013). Predators also are detrimental to webworm survival. Among the insect predators are potter wasps (Hymenoptera: Vespidae), digger wasps (Hymenoptera: Sphecidae), robber flies (Diptera: Asilidae), and damsel bugs (Hemiptera: Nabidae). Numerous species of birds, particularly blackbirds (Icteridae), have been cited as contributing to webworm mortality, but no assessments of impact are available for North America. Weather. The weather has been implicated repeatedly in the development of outbreak populations of beet webworm, and subsequent extensive damage. Damaging populations are generally limited to the Great Plains and Rocky Mountain region by too much precipitation to the east and too little precipitation to the west. Populations survive well in areas with 2.5–6.5 cm of precipitation monthly during the growing season. Also, the mean temperature greater than 13°C is deleterious, which limits the southern occurrence of webworms (Pepper, 1938). The condition of prepupae as they enter the winter is considered critical. High temperature during autumn hastens larval development, shortens the feeding period, and reduces larval weight. Thus, cool weather during autumn promotes development of large larvae that have high fecundity during the following spring (Bykova, 1984). In recent years, the migration of beet webworm has been studied extensively in Russia and China. Both longand short-distance migrations occur, often at high altitudes and assisted by the wind. The moth flights were oriented to the northeast in the spring, but less so in the summer months (Feng et al., 2004). The larvae enter diapause under long daylength conditions (14–15 h), and presumably cannot persist in the northern areas of their geographic distribution without annual infusion of moths from southern areas (Akhanaev et  al., 2013). Long-distance migration ­synchronizes oviposition by moths, resulting in large

i­ nsect populations and increased likelihood of plant damage (Cheng et al., 2012). Life Cycle and Description. One generation requires 30–40  days, and three to four generations occur annually. Moths first appear in June; thereafter, all stages of development are present until cold weather. Mature larvae overwinter in the soil, and though few larvae from the first generation diapause, an increasing proportion of larvae from each generation enters diapause as the season progresses. Egg. Beet webworm eggs are flattened, oval in shape and measure about 1 mm long and 0.7 mm wide. They are occasionally deposited singly, but more often in small clusters, usually in a single row, with individual eggs overlapping slightly. They are usually found on the underside of leaves, though females sometimes deposit eggs near succulent plants on dry twigs and clods of soil. Each female deposits 200–300 eggs, and oviposit over a broad temperature range of about 20–32°C. Duration of the egg stage under field conditions is normally 3–5 days.

FIG. 10.7  Beet webworm larva. (Photo by J. Capinera.)

Larva. Larvae are mostly green or yellowish green, but sometimes darker. A pronounced dark stripe is located dorsally, and a broken dark stripe on each side; each dark stripe is bordered on each side by a white stripe. Despite the presence of the stripes, perhaps the most striking feature is the numerous white circular spots on the body segments. Each circular marking consists of a dark spot from which protrudes a hair and is surrounded by a white ring. Larvae display five instars and grow about 4 mm long at hatching to 20 mm at maturity. Total larval development time is 12, 18, and 29 days when reared at a constant temperature of 32°C, 26°C, and 22°C, respectively. Under normal field conditions, the duration of the larval stage is about 17–20 days. Larval feeding occurs over a temperature range of 15–44°C, but the optimal temperature is about 30°C. Larvae often construct a silken tube leading from a protected area on the host plant or in the soil, to the feeding site, and sometimes

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web together leaves. This webbing behavior is the basis for the common name. Pupa. At larval maturity, insects enter the soil and construct a silk-lined cell that varies from 2.5 to 5.0 cm long. The cell is oriented vertically, with the uppermost end within 1 cm of the surface. Pupation occurs within the cell, with pupae changing from yellow to brown as they mature. They measure about 12 mm long. Duration of the pupal period is related to temperature, with development periods of 35, 16, 9, and 6 days when reared at 18°C, 22°C, 27°C, and 32°C. Duration of the pupal stage under normal field conditions is about 11 days. Not all larvae that enter the soil proceed to immediate pupation, as many enter diapause in the prepupal stage. The proportion of each generation entering diapause varies; in Montana the proportion of first-generation larvae that enters diapause varies from 0.5% to 60%. Induction of diapause is related to day length; short day length (less than 13 h of photoperiod) induces diapause (Khomyakova et al., 1986). Pupation of overwintering larvae (prepupae) occurs in the spring. The posterior, pointed end of the pupa bears eight small spines. This character serves to distinguish beet webworm from alfalfa webworm, Loxostege cereralis (Zeller), which bears eight small spoon-shaped appendages instead.

larval populations because infertility is common among females. Adults are commonly seen collecting nectar. A female-produced sex pheromone was identified by Struble and Lilly (1977). Beet webworm moths are often confused with adults of alfalfa webworm. However, they are easily differentiated by viewing the underside of the wings. Both species have a narrow dark line along the distal edge of the wings, but the line is complete in beet webworm whereas it is broken in alfalfa webworm. Beet webworm was described by Gillette (1905), Marsh (1912c), Paddock (1912), Pepper and Hastings (1941), and Maxson (1948). Adults were included in keys by Munroe (1976) and Capinera and Schaefer (1983). The larva was included in the field key by Capinera (1986).

Damage The first two instars feed on the underside of foliage, skeletonizing the leaves. Large larvae consume holes in foliage, eventually eating all except the principal veins and stems. At high densities, fourth instar or older larvae may disperse long distances in dense aggregations, a behavior typical of “armyworms.” It is under this high density and dispersing conditions that so many plants are destroyed. Damage to crop plants also results when preferred weed species are exhausted and larvae are forced to seek alternate food sources.

Management

FIG. 10.8  Beet webworm. (Photo by J. Capinera.)

Adult. The emergence of moths from the overwintering population occurs in May-July in Montana. In some years, large synchronous emergences follow protracted periods of warm weather. Other years, brief periods of favorable weather interspersed with unfavorable weather result in protracted emergence. The moths are grayish brown in general color, with irregular dark and light markings crossing the forewings. Most prominent of the markings is a dark border distally on the front wing bordered by a cream-colored band. When at rest, the wings are folded back to give the triangular form typically found in the family Pyralidae. The wingspan of beet webworm moths is about 21–22 mm. The moths may disperse in great aggregations and are attracted to lights. Heavy flights do not necessarily precede high

Sampling. The eggs and larvae can be found by visual examination of plants, but both of these life stages are difficult to detect. Moths are attracted to light and can be captured in light traps. Adults also can be flushed during the day, especially toward evening. Thus, the adult population census is an important element of population monitoring. However, sterility is common in adults, and their presence does not necessarily indicate impending damage; rather, it should serve as a stimulus to initiate careful monitoring of eggs and larvae. Insecticides. Beet webworm populations are usually suppressed by application of insecticide to foliage. Bacillus thuringiensis provides some control. Populations are infrequently damaging, so insecticides should not be applied unless high population densities of larvae are observed. Cultural Practices. Several practices can alleviate webworm damage. Tillage can disrupt and destroy overwintering larvae within their silken tubes in the soil. Destruction of preferred weeds before adult oviposition flights can minimize the deposition of eggs within crops. Destruction of weeds after egg hatching, however, tends to drive larvae to nearby crop plants. Crops planted into land immediately after alfalfa are at greater risk because not only it is a suitable host but in the later stages of its growth cycle, it is often interspersed with numerous weeds.

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Cabbage Budworm

Hellula phidilealis (Walker) (Lepidoptera: Crambidae)

Natural History Distribution. Cabbage budworm is a tropical insect. It is a common pest of cabbages in the Caribbean and in Central and South America, and also occurs in West Africa. In the United States, it occurs in Florida, Texas, Arizona, and Hawaii. Host Plants. Budworm attacks several crucifer crops, including cabbage, cauliflower, Chinese cabbage, radish, and turnip. Natural Enemies. This insect has not been well studied, and few natural enemies are known. The parasitic fly Nemorilla maculosa Meigen (Diptera: Tachinidae) is reported to parasitize budworm. However, in the Caribbean region, Alam (1992) reported seven species of parasitoids, including four species of Braconidae, and one each of Ichneumonidae and Benthylidae (all Hymenoptera), plus one species of Tachinidae (Diptera). Life Cycle and Description. The total generation time of cabbage budworm is about 30  days, and it breeds continuously in tropical areas, including southern Florida. Egg. The eggs are deposited singly or in small clusters, often along the leaf midrib. The oval eggs are white when first deposited, but soon turn brown, and hatch in about 3 days. They measure about 0.5 mm in diameter and bear a longitudinal ridge. The female produces 3–4 egg clusters, producing eggs at about 16 per day over a period of about 5 days. Total egg production averages about 65 per female, but may reach 160 per female. Larva. Newly emerged larvae feed initially on the lower surface of the leaf and then tunnel into a petiole or leaf vein. Frass is ejected from the entrance hole as the larvae feed. Feeding tunnels are lined with silk threads. The larval developmental period averages about 16  days. There are usually six instars, with average development times of 4.2, 2.2, 2.9, 2.1, 2.3, and 5.6 days, respectively. Larvae are creamy white with three reddish-brown longitudinal stripes dorsally. At maturity, they measure about 10–14 mm long. The head capsule is black. About 2 days before pupation the larvae cease feeding and spin a whitish silken cocoon, and pupate within. Pupa. Pupation may occur on the plant, near the exit hole, or nearby in the soil. The pupa is about 9 mm long, yellow brown in color, and covered with a gray waxy secretion. The pupal period is about 10 days.

Adult. Adults are dimorphic. The males are dark brown with an undulating wavy fringe on the forewings, whereas the females are light brown and lack the fringe. The hind wings are whitish, becoming darker at the margins. The moths are quite similar in appearance to cabbage webworm, Hellula rogatalis (Hulst), but they lack the yellowish tint on the forewings. Adult longevity averages 10  days, and the moths are nocturnal. The biology of cabbage budworm was given by Cadogan (1983) and King and Saunders (1984).

Damage Larvae bore into leaf stalks, stems, and the growing points of plants. Destruction of the growing point causes the plant to produce several small heads rather than one large one; these small heads have no commercial value. Damage to stems can result in stunted plants. It is considered to be a serious pest in parts of the Caribbean, but elsewhere it is only occasionally important. It is a pest of small plots and gardens, particularly at low altitude.

Management Effective insecticidal control, including the use of Bacillus thuringiensis, requires good penetration into the foliage canopy. As insects quickly bore into plant tissue, it is important to treat before the infestation becomes severe. Sanitation also is very important. Many problems stem from the presence of crop residues or volunteer plants, especially Chinese cabbage.

Cabbage Webworm

Hellula rogatalis (Hulst)

Oriental Cabbage Webworm

Hellula undalis (Fabricius) (Lepidoptera: Crambidae)

Natural History Distribution. Cabbage webworm, Hellula rogatalis, apparently is an American insect and is found in the southern states from Virginia and Florida to California. Occasionally it is reported from a more northern location, such as Nova Scotia, but is not a pest in northern climates. When first studied in Georgia in the late 1800s, it was confused with the closely related H. undalis. This latter species, sometimes called the oriental cabbage webworm, is found in tropical and subtropical areas of Europe, Africa, and Asia. It is also found in Hawaii and other islands in the Pacific Ocean. Host Plants. These insects feed on several crucifer crops, including broccoli, cabbage, collards, kale, kohlrabi,

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mustard, radish, rutabaga, and turnip. Studies by Latheef and Irwin (1983) failed to demonstrate significant preference among collards, kale, mustard, and turnip. Weed hosts include shepherdspurse, Capsella bursa-pastoris; and purslane, Portulaca oleracea. Natural Enemies. Despite a report by Chittenden and Marsh (1912) of several fly (Diptera: Tachinidae) and wasp (Hymenoptera: Braconidae and Ichneumonidae) parasitoids, more recent studies have failed to identify natural enemies of cabbage webworm (Ru and Workman, 1979; Kok and McAvoy, 1989). Life Cycle and Description. Cabbage webworm generation time is 43, 34, and 23 days at 26°C, 30°C, and 35°C, respectively. They breed continuously in southern Florida and Hawaii, but in more temperate areas such as Virginia, they become numerous enough to cause damage only during the autumn months.

leaf, but large larvae also feed on the leaf midrib. There are five instars. The head capsule widths are 0.2, 0.3, 0.5, 0.8, and 1.2 mm, respectively. Average body length for the five instars is 0.9, 2.4, 4.8, 6.4, and 9.9 mm, respectively. Development time of the larval instars at 30°C is 2.8, 2.4, 1.9, 2.4, and 4.3 days, respectively. The mature larva is yellowish gray, with five brownish-purple longitudinal bands running the length of the insect, and attains a maximum length of 15 mm. The head capsule is black. The body is sparsely covered with moderately long yellow- or light-brown hairs and tapers at both the anterior and posterior ends.

Egg. The eggs are gray or yellowish green when first deposited, but turn pink as embryonic development proceeds. The flattened eggs, which are often marked with a distinct nipple at one end, measure about 0.3 mm wide and 0.5 mm long. Oviposition does not occur at 15°C, though eggs incubated at this temperature can hatch. Temperatures of 20–30°C are suitable both for oviposition and egg hatching. Eggs hatch after about 3 days. They are normally deposited singly or in small masses on the terminal leaves.

Adult. The moth consists of yellowish-brown front wings which are marked with white bands and a dark kidney-shaped spot. The hind wings are grayish white, ­ though the margin is darker. The wingspan of the moth is about 18–21 mm. Oviposition begins about 3–5 days after moths emerge. About 150–300 eggs are produced by each female. Adult longevity is normally 7–14 days. The biology of cabbage webworm was presented by Chittenden and Marsh (1912) and McAvoy and Kok (1992). A larval key that includes cabbage webworm is Sparks and Liu (2001). Keys for the identification of Hellula spp. larvae were provided by Allyson (1981). The oriental cabbage webworm, H. undalis, is very similar in appearance to H. rogatalis (Munroe, 1972). The genitalia are used to distinguish these species. The biology of H. undalis was given by Youssef et al. (1973) and Sivapragasam and Abdul Aziz (1992) and is virtually identical to that of H. rogatalis.

Pupa. Pupation occurs in the soil, in a webbed cocoon comprising grains of soil. The duration is 5.0–5.5 days at 30°C. The pupa is yellowish brown, and measures about 7–9 mm long.

Damage

FIG.  10.9  Cabbage webworm larva webbing leaf tissue. (Photo by J. Capinera.)

Larva. The first and second instars feed singly as a leafminer between the upper and lower epidermis. Initially, the larva is yellowish gray, but soon takes on an appearance resembling the mature larva. At about the third instar, larvae begin to web and fold the foliage. Much of the damage occurs on the lower surface of the

Larvae initially mine the leaves, eventually webbing and rolling the foliage. They may cause enough damage to destroy the growing tip of plants. Thus, the problem is most severe with young plants (Smith and Brubaker, 1938). Webworms also sometimes burrow into veins, causing the death of the leaf beyond the point of feeding. Cabbage webworm is normally a minor component of the lepidopterous defoliator complex of crucifers. However, Latheef and Irwin (1983) suggested that it had the potential to become one of the most serious pests in Virginia, especially of autumn-grown crops. Kok and McAvoy (1989) reported that webworm comprising 43% of the larvae affected broccoli during a 1987 study in Virginia, making it the most abundant defoliator.

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Management Insecticidal control can be difficult owing to the cryptic feeding behavior of the larvae, and the tendency to feed on the rapidly expanding terminal growth. To keep the terminal tissue protected with chemical insecticides or Bacillus thuringiensis, frequent applications are required, at least once in a week. Evaluation of trap crops in India (Srinivasan and Krishna Moorthy, 1992), using early planted mustard to attract insects and thereby to reduce damage to cabbage, was effective for both diamondback moth, Plutella xylostella (Linnaeus) and oriental cabbage webworm, H. undalis; this is likely effective for H. rogatalis, also. In Virginia, late-maturing cultivars tend to be more heavily infested than early maturing varieties (Vail et al., 1991).

Celery Leaftier

Udea rubigalis (Guenée)

False Celery Leaftier

Udea profundalis (Packard) (Lepidoptera: Crambidae)

Natural History Distribution. Celery leaftier, also known as “greenhouse leaftier,” is found throughout the United States. Its distribution is favored by its adaptability to both indoor- and outdoor-plant cultivation. As a celery pest, it has proved to be numerous, and destructive, in all major celery-growing regions including California, Florida, Michigan, and New York. It is known to cause damage in the field in southern Ontario, but in most of Canada, it is known only as a greenhouse pest. This insect also occurs in Central and South America. False celery leaftier, which occurs only on the Pacific Coast from British Columbia to California, is a much less important pest and is poorly known. It closely resembles celery leaftier. Host Plants. In the field, celery leaftier is principally a pest of celery, but has also damaged sugarbeet and lettuce, and feeds on bean, beet, cabbage, cauliflower, kale, parsley, and probably other crops. In the greenhouse, its host range is quite large and includes ageratum, anemone, calendula, carnation, cineraria, cucumber, dahlia, daisy, geranium, lettuce, sweetpea, snapdragon, rose, and violet, but chrysanthemum is most important. Plants in the family Compositae seem preferred. A long list of greenhouse hosts was given by Weigel et al. (1924). However, neither the flowers nor the vegetables are usually attacked under field conditions, and Ball et al. (1935) suggested that the luxuriance of the forced, greenhouse-grown plants was the most important factor in allowing larvae to develop on these plants.

Weeds are important hosts. In Florida, key elements in celery leaftier biology are redroot pigweed, Amaranthus retroflexus, and spiny amaranth, A. spinosus. These species and to a lesser extent several other weed hosts are ­important in maintaining the leaftiers through the celery-free summer period. Other known hosts are plantain, Plantago spp.; wild lettuce, Lactuca spp.; cowslip, Caltha palustris; ­tickweed, Verbesina virginica; and water hemp, Acnida sp. In California, sugarbeet similarly provides a suitable host during the celery-free summer months. Essig (1934) reported that false celery leaftier fed extensively on hawksbeard, Crepis sp. Natural Enemies. Studies in Florida documented the importance of several insect parasitoids of celery leaftier (Ball et al., 1935), though the most important was an egg parasite, Trichogramma sp. (Hymenoptera: Trichogrammatidae). It could account for 70%–90% mortality but was only effective during warm weather. During the summer months, the egg parasites also use Amaranthus-infesting webworms as hosts. Of lesser importance were the larval parasitoids, Casanaria infesta Cresson (Hymenoptera: Ichneumonidae) and Cotesia marginiventris (Cresson) (Hymenoptera: Braconidae). These normally accounted for less than 5% mortality. Ball et  al. (1935) provided an unusually good documentation on the importance of birds as predators of celery leaftier in Florida. They contended that during the spring northward migration, birds faced difficulty in finding adequate food supplies. Thus, celery leaftier and celery looper, Anagrapha falcifera, which can be abundant in celery during the spring months, are a prime food source. Small fields were often maintained free of caterpillars by birds, principally palm warbler, Dendroica palmarum, and tree swallow, Tachycineta bicolor. Life Cycle and Description. Under ideal conditions, the celery leaftier may complete its life cycle in a month, and this occurs under greenhouse conditions. Under field conditions, development is delayed, depending on the weather. In California, a generation requires 1  month in the summer, 2 months in the spring and autumn, and over 3  months during the winter. Thus, five to six generations occur annually, with three to four during the period of June-December and two during the remainder of the year (Campbell, 1927). In Illinois, four generations occur under field conditions. Egg. Celery leaftier eggs are about 0.8 mm long and 0.6 mm wide, spherical, and slightly flattened. The eggs are shiny, and whitish initially, growing darker as they mature. The eggs are deposited on the underside of leaves singly or in small groups of up to 12. As is common with pyralids, the eggs overlap one another. The incubation period is usually about 6  days (range 4–10  days). Most eggs are deposited just below the outer layer of the plant canopy.

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FIG. 10.10  Celery leaftier larva. (Drawing by USDA.)

Larva. The larvae are pale yellowish white and measure only about 1 mm long initially, but soon become pale green, with a single dark green line dorsally and bordered on each side by a broad whitish band. This striped pattern persists throughout the larval period. The color of the ventral surface of the larva tends toward yellow. The head, thoracic shield, and thoracic legs are pale, though the shield bears a dark, oval spot on each side. The larvae are sparsely covered with long hairs. Larvae feed on the lower surface of leaves, often rolling leaves or webbing them together. In most respects, larvae behave like other webworms, including the tendency to retreat into their webbed shelter or to wriggle violently and spin down from the plant on a strand of silk if disturbed. At maturity, the larvae measure only about 17–19 mm long. There are five larval instars, each requiring 2–5 days for completion. The total development time of the larval stage averages about 21 days (range 15–30 days). Head capsule widths for the instars are about 0.20, 0.27, 0.40, 0.67, and 1.05 mm, respectively. Pupa. Pupation normally takes place in a thin, whitish silk cocoon amongst folded leaves. The pupa is smoky brown and measures 8.5–9.0 mm long. Duration of the pupal stage averages 10 days (range 6–16 days).

Adult. The moth of celery leaftier is small, the wingspan measuring only 15–21 mm. The front wings are light brown, suffused with reddish brown, and are irregularly marked with black lines. The hind wings are grayish, becoming brown distally. The outer margin of the fore and hind wings are marked with a row of small, dark spots. Adults live for several days; males tend to survive about 4–5 days and females 9–10 days. They are nocturnal and remain hidden during the daylight hours. Mating occurs almost immediately upon adult emergence, and oviposition begins within 24 h of emergence. Eggs, which tend to be deposited principally during the first few days of adulthood, are deposited at night. Adults of both sexes can be attracted to traps containing the floral attractant phenyacetaldehyde, which can be made more attractive with the addition of linalool and beta-myrcene (Landolt et al., 2004). The eggs, larvae, pupae, and moths of false celery leaftier are not easily differentiated from celery leaftier and in nearly all respects the biology is the same. Munroe (1966) gave characters to distinguish the moths. False celery leaftier averages slightly larger than celery leaftier, the wingspan ranging from 19 to 25 mm. The larvae are also slightly larger. Mean head capsule widths for the five instars are 0.23, 0.34, 0.54, 0.82, and 1.23 mm, respectively. The field ecology of celery leaftier was best described by Ball et al. (1935); developmental biology was provided by Fletcher and Gibson (1901) and Weigel et  al. (1924). False celery leaftier is poorly studied; Tamaki and Butt (1977) gave the only substantive biological information on this insect.

Damage Larvae feed mostly on the underside of foliage. Depending on the thickness of the plant tissue they may eat the entire leaf, only the lower surface, or leave the foliage skeletonized. They also cover the leaves with a thin layer of silk. The silk webbing may be used to draw portions of the leaf together. Celery leaftier larvae may work their own way down to the youngest tissue, the “heart” of celery, and feed on both leaves and petioles (Stone et al., 1932). Jones and Granett (1982) indicated that false celery leaftier also damaged the heart of celery in California.

Management

FIG. 10.11  Celery leaftier adult. (Drawing by USDA.)

Moths of celery leaftier are attracted to phenylacetaldehyde, and this chemical may be useful with some types of traps for population monitoring (Cantelo et al., 1982). They also come to light traps. Foliar insecticides are applied for leaftier suppression, and growers are encouraged to target the young larvae for control because they are easier to kill. This insect is only occasionally threatening, however.

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Cross-Striped Cabbage Worm Evergestis rimosalis (Guenée) (Lepidoptera: Crambidae)

Natural History Distribution. Cross-striped cabbage worm is recorded principally from North America to the east of the Rocky Mountains but is infrequent in the northernmost states and eastern Canada. This insect is known as a common pest only in the southeastern states and in Central America, and even in the southeastern states it usually is a minor component of the defoliator complex. It is likely a native insect, though its close relatives are native to Europe. Cross-striped cabbage worm was accidentally introduced to Australia and Jamaica. Host Plants. Broccoli, Brussels sprouts, cabbage, collards, kale, rutabaga, and turnip are among the vegetable crops known as hosts. Brussels sprouts and collards tend to support relatively large numbers of cabbage worms, and cabbage and kale relatively few. Other crucifers such as rape, and many weeds, likely support this insect. Natural Enemies. Several parasitoids are known, including Cotesia congregata Say, C. xylina (Say), and C. orobenae Forbes (all Hymenoptera: Braconidae). Except for C. orobenae, the importance of the parasitoids has not been thoroughly assessed, and predators and diseases are undocumented. Mays and Kok (1997) reported on C. orobenae, and showed that the occurrence of this parasitoid closely tracked the abundance of its host. Kok and AcostaMartinez (2001) documented differential susceptibility of C. orobenae to insecticides, with B. thuringiensis not affecting the parasitoids. Life Cycle and Description. Development time, from egg to adult stages, ranges from 61  days at 20°C to 18  days at 35°C. The number of annual generations is three in Illinois, and four in Maryland. Cross-striped cabbage worm tends to be most abundant during the autumn generation in the north. In the south, however, it can be quite abundant during the winter- and spring-cropping period. Egg. The eggs are laid on the underside of leaves in small masses, usually from 3 to 5 to about 25. They are yellow, flattened, and overlap slightly. They are oval, measure about 1.2 mm long and 0.9 mm wide. They hatch in about 12.4 days when reared at 20°C and 1.8 days at 35°C, hatching most readily at 20–30°C.

FIG. 10.12  Cross-striped cabbageworm larvae. (Photo by L. Buss.)

Larva. Four instars are reported for this species, with mean head capsule widths (range) of 0.34 (0.32–0.38), 0.56 (0.52–0.60), 1.04 (0.98–1.10), and 1.66 (1.58–1.71) mm, respectively, for instars 1–4. Mean duration of the instars, when cultured at 25°C, is 3.0, 2.0, 2.0, and 5.8  days, respectively. Small larvae are gray, with black tubercles bearing stout hairs. Larger larvae are bluish-gray dorsally, and with numerous transverse black bands. Transverse lines are ­relatively rare among caterpillars and serve as a key diagnostic feature for this insect. The prominent tubercles are gray, marked with black. A yellow line occurs along each side of the caterpillar. The underside of the caterpillar is green, mottled with yellow. The mature larva measures about 15–17 mm long. Larval development requires 2–3 weeks. Pupa. Transformation into a pupa occurs in the soil, near the surface, in a small cocoon covered with particles of sand. The yellowish-brown pupa measures about 10–12 mm long. Duration of this stage is 9–11 days when reared at 30–20°C.

FIG. 10.13  Cross-striped cabbage worm. (Photo by L. Buss.)

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Adult. The adult has a wingspan of about 25 mm. The front wings are straw colored, but are also marked with olive or purplish brown, and crossed by narrow transverse lines. The hind wings are transparent and whitish, with a darker band at the margin. The preoviposition period is 3–6 days, and the oviposition period is 6–14  days, when reared at 20–30°C. Females oviposit readily at 20–30°C, but oviposition frequency decreases at higher and lower temperatures. Adults live for over 20 days if held under cool conditions but survive for only 5–11 days at 30°C or above. The biology of cross-striped cabbage worm was given by Chittenden (1902) and Mays and Kok (1997). A key including the adult stage of cross-striped cabbage worm was contained in Munroe (1973). Cross-striped cabbage worm is included in the key to “cabbageworms” in Appendix A.

Damage Larvae feed on foliage, creating small holes. They prefer the terminal buds. If disturbed, they have a tendency to drop from the foliage on a silken thread. Larvae also may burrow into the center of developing heads.

Management Cross-striped cabbage worm, along with several other lepidopterous defoliators, exhibits a positive response to nitrogen fertilization of host plants. However, indications are that unlike the case with piercing-sucking insects, it is not the increased nitrogen levels per se, but the increased foliar biomass that favors the abundance of the cabbage worms (Jansson et  al., 1991b). In general, methods used for management of imported cabbage worm, Pieris rapae (Linnaeus), are appropriate for this insect.

European Corn Borer

Ostrinia nubilalis (Hübner) (Lepidoptera: Crambidae)

Natural History Distribution. First found in North America near Boston, Massachusetts in 1917, European corn borer quickly spread to the Great Lakes region. By 1948 it was established throughout the midwestern corngrowing region and eastern Canada. It has now spread as far west as the Rocky Mountains in both Canada and the United States, and south to the Gulf Coast states. European corn borer is thought to have originated in Europe, where it is widespread. It also occurs in northern Africa. The North American European corn borer population is thought to have resulted from multiple introductions from more than one area of Europe.

Thus, there are at least two, and possibly more, strains present. The presence of an eastern or New York strain, and a midwestern or Iowa strain is evident because different pheromone blends are required to capture moths from each population. Both strains sometimes occur in the same area (Eckenrode et al., 1983). Host Plants. European corn borer has a very wide host range, attacking practically all robust herbaceous plants with a stem large enough for the larvae to enter. However, the eastern strain accounts for most of the wide host range, with the western strain feeding primarily on corn. Among vegetable crops injured are beet, broccoli, celery, corn, cowpea, eggplant, lima bean, pepper, potato, rhubarb, snap bean, spinach, Swiss chard, and tomato. Vegetables other than corn tend to be infested if they are abundant before corn is available, or late in the season when senescent corn becomes unattractive for oviposition; snap and lima beans, pepper, and potato are especially damaged. In North Carolina, for example, potato is more attractive than corn at peak emergence of the first moth flight, and more heavily damaged (Anderson et  al., 1984). Other crops sometimes attacked include buckwheat, grain corn, hop, oat, millet, and soybean, and such flowers as aster, cosmos, dahlia, gladiolus, hollyhock, and zinnia. Corn is the most preferred host, but many thickstemmed weeds and grasses also support European corn borer, especially if they are growing amongst, or adjacent to, corn. Some of the common weeds infested include barnyardgrass, Echinochoa crusgalli; beggarticks, Bidens spp.; cocklebur, Xanthium spp.; dock, Rumex spp.; jimsonweed, Datura spp.; panic grass, Panicum spp.; pigweed, Amaranthus spp.; smartweed, Polygonum spp.; and others. A good list of host plants was given by Caffrey and Worthley (1927). Natural Enemies. Native predators and parasites exert some effect on European corn borer populations, but imported parasitoids seem to be more important. Among the native predators that affect the eggs and young larvae are the insidious flower bug, Orius insidious (Say) (Hemiptera: Anthocoridae); green lacewings, Chrysoperla spp. (Neuroptera: Chrysopidae); and several lady beetles (Coleoptera: Coccinellidae) (Jarvis and Guthrie, 1987; Andow, 1990). Insect predators often eliminate 10%–20% of corn borer eggs. Avian predators such as downy woodpecker, Dendrocopos pubescent (Linnaeus); hairy woodpecker, D. villosus (Linnaeus); and yellow-shafted flicker, Colaptes auratus (Linnaeus) have been known to eliminate 20%–30% of overwintering larvae. Native parasitoids include Bracon caulicola (Gahan), B. gelechiae Ashmead, B. mellitor Say, Chelonus annulipes Wesmael, Macrocentrus delicatus Cresson, and Meteorus

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campestris Viereck (all Hymenoptera: Braconidae); Gambrus ultimus (Cresson), G. bituminosus (Cushman), Itoplectis conquisitor (Say), Campoletis flavicincta (Ashmead), Nepiera oblonga (Viereck), Rubicundiella perturbatrix Heindrich, Vulgichneumon brevicinctor (Say) (all Hymenoptera: Ichneumonidae); Dibrachys carus (Walker) and Eupteromalus tachinae Gahan (both Hymenoptera: Pteromalidae); Syntomosphyrum clisiocampe (Ashmead) (Hymenoptera: Eulophidae); Scambus pterophori (Ashmead) (Hymenoptera: Hybrizontidae); Trichogramma nubilale Ertle and Davis and T. minutum Riley (both Hymenoptera: Trichogrammatidae); and Archytas marmoratus (Townsend) and Lixophaga sp. (both Diptera: Tachinidae). Although many species of native parasitoids are known, native parasitoids rarely cause high levels of corn borer mortality. Exotic parasitoids numbering about 24 species have been imported and released to augment native parasitoids. About six species have successfully established. Among the potentially important species is Lydella thompsoni Herting (Diptera: Tachinidae), which may kill up to 30% of second-generation borers in some areas, but has disappeared or gone into periods of low abundance in other areas. Other exotic parasitoids that sometimes account for more than trivial levels of parasitism are Eriborus terebrans Gravenhorst (Hymenoptera: Ichneumonidae), Simpiesis viridula (Hymenoptera: Eulophidae), and Macrocentris grandii Goidanich (Hymenoptera: Braconidae) (Burbutis et al., 1981; Andreadis, 1982a; Losey et al., 1992). A comprehensive review of biological control agents imported in the first half of the 1900s was published by Baker et al. (1949). Several microbial disease agents are known from corn borer populations. The common fungi Beauveria bassiana and Metarhizium anisopliae are sometimes observed, especially in overwintering larvae. The most important pathogen seems to be the fungus (microsporidian), Nosema pyrausta, which often attains 30% infection of larvae and sometimes 80%–95%. It creates chronic, debilitating infections that decrease longevity and fecundity of adults, and decreases survival of larvae that are under environmental stress (Hill and Gary, 1979; Andreadis, 1984; Lewis et  al., 2009). Unfortunately, N. pyrausta also infects the parasitoid M. grandii (Andreadis, 1982b). Life table studies conducted on corn borer populations in Quebec with a single annual generation perhaps provide insight into the relative importance of mortality factors (Hudon and LeRoux, 1986c). These workers demonstrated that egg mortality (about 15%) was low, stable and due mostly to predators and parasites. Similarly, the mortality of young larvae, due principally to dispersal, dislodgement, and plant resistance to feeding was fairly low (about

15%) but more variable. Mortality of large larvae during the autumn (about 22%) and following spring (about 42%) was due to a number of factors including frost, disease and parasitoids, but parasitism levels were low. Pupal mortality (about 10%) was low and stable among generations. The factor that best accounted for population trends was considered the survival of adults. Dispersal and disruption of moth emergence by heavy rainfall are thought to account for high and variable mortality (68%–98%, with a mean of 95%), which largely determines population size of the subsequent generation. Overall generation mortality levels were high, averaging 98.7%. Weather. There are many reports that weather influences European corn borer survival. Heavy precipitation during egg hatch, for example, is sometimes given as an important mortality factor (Jarvis and Guthrie, 1987). Low humidity, low nighttime temperature, heavy rain, and wind are detrimental to moth survival and oviposition. However, during a 10-year, three-state study, Sparks et  al. (1967) reported no consistent relationship between weather and survival. Life Cycle and Description. The number of generations varies from one to four, with only one generation occurring in northern New England and Minnesota and in northern areas of Canada, three to four generations in Virginia and other southern locations, and usually two generations in the northern United States and southern Canada. In many areas, generation number varies depending on weather, and there is a considerable adaptation for local climate conditions even within strains. For example, though the developmental rates of single-generation strains are lower than multiplegeneration strains, at northern locations such as Prince Edward Island the single-generation strain develops quickly (Dornan and Stewart, 1995). European corn borer overwinters in the larval stage, with pupation and emergence of adults in early spring. Diapause apparently is induced by exposure of last instar larvae to long days, but there is also a genetic component. Moth flights and oviposition usually occur during June-July and August-September in areas with one to two generations annually. In southern locations with three generations, moth flights and oviposition typically occur in May, late June, and August. In locations with four generations, adults are active in April, June, July, and August-September. Egg. The eggs are deposited in irregular clusters of about 15–20 (range 5–50 eggs). They are oval, flattened, and creamy white, usually with an iridescent appearance. They darken to a beige or orangish tan color with age. They are normally deposited on the underside of leaves and overlap like shingles on a roof or fish scales. The eggs measure about 1.0 mm long and 0.75 mm wide. The developmental threshold for eggs is about 15°C. Eggs hatch in 4–9 days.

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FIG. 10.14  European corn borer larva. (Photo by J. Capinera.)

Larva. Larvae tend to be light brown or pinkish-gray dorsally, with a brown to black head capsule and a yellowishbrown thoracic plate. The body is marked with round dark spots on each body segment. The developmental threshold for larvae is about 11°C. Larvae normally display six instars, but 4–7 instars have been observed. Head capsule widths are about 0.30, 0.46, 0.68, 1.03, 1.66, and 2.19 mm in instars 1–6, respectively. Mean body lengths during the six instars are about 1.6, 2.6, 4.7, 12.5, 14.5, and 19.9 mm, respectively. For populations with only five instars, mean head capsule widths are 0.29, 0.44, 0.80, 1.27, and 2.00 mm, respectively. Young larvae tend to feed initially within the whorl, especially on the tassel. When the tassel emerges from the whorl, larvae disperse downward where they burrow into the stalk and the ear. Mortality tends to be high during the first few days of life, but once larvae establish a feeding site within the plant, survival rates improve. Larvae in the final instar overwinter within a tunnel in the stalk of corn, or in the stem of another suitable host. Duration of the instars varies with temperature. Under field conditions in New York, development time was estimated at 9.0, 7.8, 6.0, 8.8, 8.5, and 12.3 days for instars 1–6, respectively, for a mean total development period of about 50 days. In contrast, during the next year development time at the same site was 4.4, 4.3, 4.6, 5.8, 8.5, and 9.0 days for the six instars, for a mean total larval development period of about 35 days (Caffrey and Worthley, 1927). Pupa. Pupae usually occur in April or May, and then later in the year if more than one generation occurs. The pupa is normally yellowish brown. The pupa measures 13–14 mm long and 2–2.5 mm wide in males and 16–17 mm long and 3.5–4 mm wide in females. The tip of the abdomen bears 5–8 recurved spines that are used to anchor the pupa to its cocoon. The pupa is ordinarily, but not always, enveloped in a thin cocoon formed within the larval tunnel. Duration of the pupal stage under field conditions is usually about 12 days. The developmental threshold for pupae is about 13°C.

FIG. 10.15  European corn borer, adult female. (Photo by J. Capinera.)

FIG. 10.16  European corn borer, adult male. (Photo by J. Capinera.)

Adult. The moths are fairly small, with males measuring 20–26 mm in wingspan, and females 25–34 mm. Female moths are pale yellow to light brown, with both the forewing and hind wing crossed by dark zigzag lines and bearing pale, often yellowish, patches. The male is darker, usually pale brown or grayish brown, but also with dark zigzag lines and yellowish patches. Secondary host plants and adjacent grassy areas play a significant role in the mating behavior of adults, as adults rest and mating takes place in such areas of dense vegetation, called “action sites.” Retention of droplets from rainfall and dew in this dense vegetation stimulates the sexual activity of females. Moths are most active during the first 3–5 h of darkness. The sex pheromone has been identified as 11-tetradecenyl acetate, but eastern and western strains differ in the production of Z and E isomers. The western strain produces a blend that approximates 97:3 Z:E, whereas the eastern strain uses a blend of 3:97 Z:E. The preoviposition period averages about 3.5  days. Duration of oviposition is about 14  days, with oviposition averaging 20–50 eggs per day. The female often deposits 400–600 eggs during her life span, though there are also estimates of

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mean fecundity of about 150 eggs in some locations. Total adult longevity is normally 18–24 days. Brindley and Dicke (1963), Brindley et  al. (1975), Hudon and LeRoux (1986a, b), and Hudon et  al. (1989) published reviews of the biology and management of European corn borer. Detailed biology was presented in Vinal and Caffrey (1919), and Caffrey and Worthley (1927). Sex pheromone blends were identified (Klun and Robinson, 1971; Kochansky et  al., 1975; Showers et  al., 1974). Beck (1987) reviewed corn borer seasonal biology and offered interesting insight into the pheromonal races or strains. Rearing procedures were given by Reed et al. (1972). European corn borer was included in the larval key by Capinera (1986) and the moth key by Capinera and Schaefer (1983). A key to stalk borers associated with corn in southern states was presented by Dekle (1976); this publication also included pictures of the adults. A key to pyralid borers was also included by Stehr (1987). A key to common stalk boring caterpillars is included in Appendix A.

Damage This is a very serious pest of both sweet and grain corn, and before the availability of modern insecticides, this insect caused very marked reductions in corn production. Young larvae feed on tassels, whorl, and leaf sheath tissue; they also mine midribs and eat pollen that collects behind the leaf sheath. Sometimes they feed on silk, kernels, and cobs, or enter the stalk. Older larvae tend to burrow into the stalk and sometimes the base of the corn ear, or into the ear cob or kernels. Feeding by older larvae is usually considered to be most damaging, but tunneling by even young larvae can result in broken tassels. The presence of 1–2 larvae within a corn stalk is tolerable, but the presence of any larvae within the ear of sweet corn is considered intolerable by commercial growers and this is their major concern. European corn borer is considered to be the most important sweet corn pest in northern production areas, and second-generation borers are the principal source of ear damage. Heavily tunneled stalks of grain corn suffer from lodging, reducing the capacity for machine harvesting. Lodging is not a serious threat to sweet corn. In addition, corn borer feeding makes corn susceptible to infection by Fusarium spp., resulting in ear fusariosis and Fusarium toxin production. Fusarium toxin can be poisonous to livestock fed grain corn. In crops other than corn, the pattern of damage is variable. European corn borer larvae damage both the stem and fruit of beans, pepper, and cowpea. The temporal occurrence of fruit affects susceptibility to injury, of course; in Wisconsin, snap beans 14–30 days from harvest were susceptible to damage by larvae, but young plants and fruit near harvest suffered little damage (Sanborn et  al., 1982b). In celery, potato, rhubarb, Swiss chard, and

t­omato, it is usually the stem tissue that is damaged. In beet, spinach, and rhubarb, leaf tissue may be injured. The entry of borers into plant tissue facilitates entry of plant pathogens. The incidence of potato blackleg caused by the bacterium Erwinia carotovora atroseptica, for example, is higher in potato fields with stems heavily infested by corn borers. Direct damage by corn borers to potato vines, however, results in negligible yield loss (Nault and Kennedy, 1996c).

Management Sampling. Moths can be sampled with blacklight and pheromone traps, and catches by these traps are correlated (Legg and Chiang, 1984; Welty, 1995). Pheromones attract only males, whereas both sexes are captured in traps with a blacklight. Blacklight traps tend to be more reliable, but light traps can capture many other insects, necessitating a great amount of sorting. Pheromone-baited water pan traps seem to be the most efficient method of adult monitoring (Thompson et  al., 1987; Stewart, 1994). Trap catches are usually used to initiate intensive in-field scouting for egg masses, as moth catches are only roughly correlated with density. Plant phenology can be used to predict corn borer development. In New York, for example, peak flight of the first brood of moths corresponds to bloom of elderberry, Sambucus canadensis, and peak second brood flight corresponds to peak bloom of hydrangea, Hydrangea paniulata grandiflora (Straub and Huth, 1976). Thermal summations are also highly predictive (Jarvis and Brindley, 1965). Moths seek shelter during the daylight hours in dense grass and weeds near cornfields. Flushing moths from such habitats gives an estimate of population densities (Sappington and Showers, 1983). Eggs can be sampled by visual examination, but this is a very time-consuming effort. Similarly, larval populations can be estimated from visual examinations, particularly of whorls during the first generation. A sequential sampling protocol for larvae was developed for potato (Nault and Kennedy, 1996b), and Hoffmann et  al. (1996b) described a sequential sampling plan based on infested plants. Insecticides. Liquid formulations of insecticide are commonly applied to protect against damage to sweet corn, particularly from the period of early tassel formation until the corn silks are dry. Recommendations vary from a single application before silking, to weekly applications (Ferro and Fletcher-Howell, 1985). Liquid applications are usually made to coincide with egg hatching in an effort to prevent infestation. If corn borers are present in a field, however, the critical treatment time is just before the tassels emerge, or at tassel emergence from the whorl. This plant growth period is significant because the larvae are active at this time and are more likely to contact insecticide. A popular alternative to liquid insecticides is the use of granular formulations, which can be dropped into the whorl

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for effective control of first-generation larvae because this is where young larvae tend to congregate. The insecticide is more persistent when applied in a granular formulation (Straub, 1983). Botanical insecticides such as rotenone and ryania are moderately effective against young corn borers but must be applied frequently (Turner, 1945). In grain corn, insecticide applications for suppression of secondgeneration corn borers can be made outside the cornfields in areas of thick grass, or action sites, where adults tend to aggregate (Showers et al., 1980). This approach has not been assessed for sweet corn. For borer suppression on potato, a single application of insecticide timed to coincide with the presence of first instar larvae provides an optimal yield (Nault and Kennedy, 1996a). Cultural Practices. Destruction of stalks, the overwintering site of larvae, has long been recognized as an important element of corn borer management. Disking is not adequate; plowing to a depth of 20 cm is necessary for the destruction of larvae. Mowing of stalks close to the soil surface eliminates more than 75% of larvae and is especially effective when combined with plowing (Schaafsma et  al., 1996). Minimum tillage procedures, which leave considerable crop residue on the surface, enhance borer survival. Diversified cropping is detrimental to corn borer population survival. Intercropping with red clover, for example, resulted in lower borer density (Lambert et al., 1987). Early planted corn is taller and attractive to ovipositing female moths, so late planting has been recommended, but this is useful mostly in areas of only a single generation per year. If a second generation occurs, such late-planted corn is heavily damaged. Planting border rows of a highly attractive variety of corn to surround a less attractive variety has been investigated in France (Derridj et  al., 1988). The attractive variety, especially if it is an early flowering cultivar, receives most of the eggs of moths dispersing into the field. If treated with insecticide or destroyed, this border row trap could provide protection for the main corn crop. Soil conditions can affect corn borer oviposition patterns. Research conducted in Ohio demonstrated that corn grown in rich organic soils were not as attractive to moths as low-protein plants grown in conventionally fertilized soil (Phelan et al., 1996). Host-Plant Resistance. Extensive breeding research has been conducted, and resistance has been incorporated into grain corn, especially against first-generation borers. A principal factor in seedling resistance to young larvae is a chemical known as DIMBOA, which functions as a repellent and feeding deterrent (Klun et  al., 1967). It has proven difficult to incorporate the known resistance factors into sweet corn without degradation of quality. However, some progress has been made in producing commercially acceptable resistant cultivars, especially when hostplant resistance is complemented by the use of other suppressive

tactics such as application of Bacillus thuringiensis (Bolin et al., 1996). Transgenic corn that incorporates genetic material from a toxin produced by Bacillus thuringiensis var. kurstacki (called Bt corn) has become popular in recent years. Expression of the genetic material makes the plant toxic to corn borers and related lepidopteran insects, but not to other animals. Widespread planting of Bt corn has greatly reduced the abundance of European corn borer (Burkness et  al., 2001; Hutchinson et  al., 2010). Commercial varieties of both sweet corn and grain corn are available. Pepper cultivars differ in their susceptibility to corn borer. Hot pepper cultivars are most resistant, and most green bell peppers are susceptible. Biological Control. Biological control has been attempted repeatedly in sweet corn and other vegetables susceptible to European corn borer attack. Bacillus thuringiensis products can be as effective as many chemical insecticides but often prove to be less effective than some (Bartels and Hutchison, 1995). Most single-­factor approaches, with the exception of newer formulations of Bacillus thuringiensis have proven to be erratic. Release of native Trichogramma spp. (Hymenoptera: Trichogrammatidae), for example, provides variable and moderate levels of suppression (Andow et  al., 1995). In Massachusetts, an egg parasitoid normally associated with a related Ostrinia species in China was released. This new parasitoid, Trichogramma ostriniae (Hymenoptera: Trichogrammatidae), may prove useful for augmentative biological control programs but seems susceptible to disruption by adverse weather (Wang et  al., 1997a, b). The effects of egg parasitoids are enhanced by the application of Bacillus thuringiensis (Losey et al., 1995; Mertz et al., 1995). Application of pathogens such as Nosema pyrausta and Vairimorpha necatrix (Microsporida: Nosematidae) has been proven to have benefited under experimental conditions (Lewis et al., 1982), but a commercial product has not been developed.

European Pepper Moth

Duponchelia fovealis (Zeller) (Lepidoptera: Crambidae)

Natural History Distribution. This insect has long occurred in the Mediterranean region and Canary Islands, but more recently has spread more extensively in Europe, and also to the Middle East, western Asia, Africa, and North and South America. European pepper moth has been detected widely in the United States and Canada, but in North America, these detections are mostly based on movement of plant material and are not indicative of breeding populations. However,

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it seems likely to establish permanent populations in the southeastern and west coast regions of the United States. In cooler climates it likely would remain as a greenhouse pest only. Host Plants. This species feeds on a broad range of plants, including aquatic, ornamental and some vegetable crops in addition to certain weeds. Among the vegetable crops injured are celery, beet, tomato, pepper, squash, ­cucumber, lettuce, and corn. Other food crops include basil, thyme, strawberry, blackberry, fig, pomegranate, and blackberry. Among the numerous ornamental plants affected are lisianthus, poinsettia, coleus, gerbera daisies, impatiens, kalanchoe, geranium, azalea, chrysanthemum, geranium, croton, and rose. Weeds known to support European pepper moth include lambsquarters, Chenopodium album Linnaeus (Amaranthaceae); common purslane, Portulaca oleraceae Linnaeus (Portulacaceae); common wood sorrel, Oxalis acetosella L. (Oxalidaceae); narrowleaf plantain, Plantago lanceolata L. (Plantaginaceae); bindweed, Convolvulus sp. (Convolvulaceae); pigweed, Amaranthus sp. (Amaranthaceae), and others. Importantly, the larvae can also develop on decaying plant material, so they can survive in the absence of actively growing plants (Stocks and Hodges, 2013). Suitability of an artificial diet for laboratory culture has been demonstrated (Zawadneak et al., 2017). Natural Enemies. Several natural enemies are available for European pepper moth suppression, though they are usually used on high-value crops under greenhouse conditions. The principal beneficials used to suppress European pepper moth are entomopathogenic nematodes, predatory mites, and trichogrammatid egg parasitoids (Block and Messelink, 2009; Stocks and Hodges, 2013). Life Cycle and Description. This species displays from two generations per year in cooler portions of its range, and up to 8–9 per year under greenhouse conditions. In Brazil, when cultured on a strawberry leaf diet at 25°C, the mean egg to adult development time (± SE) was 36.6 ± 0.7 days (Zawadneak et al., 2017). Oviposition commences after 8.2 (± 0.3) days. Females and males live an average (± SE) of 16.5 ± 2.3 and 15.5 (± 2.0) days, respectively. Egg. Like many crambids, the eggs are oval and flattened. They measure about 0.7 mm long and 0.5 mm wide, so they are quite small. They may be deposited singly or in small groups; in the latter case, they usually overlap, like shingles on a roof. They are usually deposited on the abaxial (lower) leaf surface, but also elsewhere including the soil surface. When first laid, the eggs are whitish or yellowish, but turn red and then brown as the embryo develops. The female moth produces about 200–300 eggs over the course of her life. When held at 25°C, eggs hatch in an average of 5.4 (± 0.2) days.

FIG. 10.17  European pepper moth larva. (Photo by L. Buss.)

Larva. Newly hatched larvae are about 1.5 mm long. At this stage, the body is salmon-pink in color, and the head and thoracic plate are black. Each body segment contains several raised structures (pinacula) that appear to be dark spots from which a hair or hairs originate. As the larva grows, the body color changes to whitish or brownish, and the dark spots are less pronounced. Also, the color of the larva is influenced by the food consumed. On foliage, they tend to be lighter, but when feeding on decaying matter they are darker. The larva attains a length of 17–30 mm prior to pupation. The spots disappear as the larva prepares to pupate. There are five larval instars. The mean (± SE) head capsule widths when fed strawberry leaves were 0.23 ± 0.01, 0.33 ± 0.01, 0.51 ± 0.02, 0.80 ± 0.01, and 1.17 ± 0.02 mm for instars 1–5, respectively. When fed an artificial diet the head capsules were quite similar (Zawadneak et  al., 2017). These authors also determined the mean (± SE) development times at 25°C, which was 5.3 ± 0.2, 3.9 ± 0.2, 3.2 ± 3.3, 4.1 ± 0.2, and 8.7 ± 0.5 days for instars 1–5 when fed strawberry leaves, and quite similar when fed artificial diet. In addition, the prepupa required an additional 2.0 ± 0.0 days prior to pupation when fed strawberry. Larvae reportedly prefer cryptic locations, and they often seek dark and sheltered locations such as near the main stem of the plant, beneath pots, or between the pot and the soil. There they feed on the stem and roots. Also, they produce webbed silk tunnels through which they move. When vegetation is dense, they move up into the crown of the plant, and sometimes into the fruit (Zawadneak et al., 2016). Pupa. Pupation occurs in a cocoon constructed from silk, fecal matter, and soil particles. The cocoon is 15–19 mm long. Initially yellow, the pupa becomes darker as the developing moth gets closer to emergence. The pupa is 10– 12 mm long, and female and male pupae weigh about 25 and 22 mg, respectively. At 25°C, the mean (± SE) pupal period was 7.8 ± 0.3 days.

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The larvae also prefer dense vegetation and accompanying moist environments, so removal of weeds, thinning of crop plants, and reduction of irrigation can reduce the suitability of crop habitat to larvae. Greenhouse vents should be netted to eliminate entry by moths.

Garden Webworm

Achyra rantalis (Guenée) (Lepidoptera: Crambidae)

Natural History FIG. 10.18  European pepper moth. (Photo by L. Buss.)

Adult. The adults are brown moths that measure about 9–12 mm long and have a wingspan of 19–21 mm. The forewings also have gray patches, and each bears two wavy, yellowish-white lines that cross the wings. The apical (distant from the body) line bears a finger-like projection that points away from the moth’s body. The proximal (close to the body) line is less pronounced and less wavy. The hind wings are more uniform gray brown, and lack distinctive lines. The abdominal segments are marked by alternating cream-colored and dark-brown bands. The abdomen is rather thin and elongate, especially in the male. When at rest, the moths tend to curve the tip of the abdomen upward.

Damage Larvae chew holes in roots, leaves, flowers, buds, and fruit, including submerged vegetation. Often the stems are girdled. This is a serious problem for greenhouse production in Europe, but in the United States, it remains mostly a threat.

Management Sampling. Visual inspection is recommended to detect eggs, larvae, pupae, and adults. However, the silken tunnels are often easiest to locate. It can be difficult to detect eggs and larvae, but moths readily use traps baited with sex pheromone. Used in conjunction with pheromones, water pan traps are most effective, but sticky traps and funnel traps also capture moths. Light traps can capture the moths, but this is a nonselective technique. Insecticides. Various contact insecticides are efficacious if contact can be made, but this can be difficult due to the cryptic and belowground habits of larvae. Systemic insecticides are useful once larvae have burrowed into plants. In greenhouse environments, fumigation is possible. Young larvae are susceptible to the bacterial insecticide Bacillus thuringiensis. Cultural Practices. Removal of decaying plant material is recommended because larvae will feed on such detritus.

Distribution. This webworm is found throughout the eastern United States, west to the Rocky Mountains, and also in California. Despite its wide distribution, it rarely damages except in the southern Great Plains region. Garden webworm also occurs in eastern Canada, in most of Mexico, and throughout the Caribbean. It is native to North America. Host Plants. Garden webworm is quite similar to alfalfa webworm, Loxostege cereralis (Zeller), and beet webworm, L. sticticalis (Linnaeus), in most aspects of its biology, including host range. Though its biology is not well documented, it is known to attack such vegetables as bean, beet, cabbage, cantaloupe, cucumber, chard, corn, cowpea, eggplant, lettuce, onion, potato, pumpkin, spinach, squash, sweet potato, and tomato. Field crops including alfalfa, clover, pea, soybean, and sugarbeet are injured. If allowed to become abundant in alfalfa, larvae may disperse when the alfalfa is harvested and may damage nearby crops, including such crops as corn and cotton, which are not normally eaten. Grain crops generally are avoided. Weed hosts include dock, Rumex spp.; lambsquarters, Chenopodium album; Parthenium sp.; pigweed, Amaranthus spp.; ragweed, Ambrosia spp.; saltbush, Atriplex patula; smartweed, Polygonum spp.; and sunflower, Helianthus spp. Pigweed and lambsquarters are regarded as favorite weed hosts. Natural Enemies. Several parasitoids of garden webworm are known, but there is little information on their relative importance. Among the wasps are Cremnops vulgaris (Cresson), C. haematodes (Brulle), Apanteles conanchetorum Viereck, A. pyraloides Muesebeck, Cardiochiles explorator (Say) (all Hymenoptera: Braconidae), and Phytodietus rufipes (Cresson) and Diadegma pattoni (Ashmead) (both Hymenoptera: Ichneumonidae). Among the fly parasitoids are Eusisyropa blanda (Osten Sacken), E. boarmiae (Coquillet), Hyphantrophaga hyphantriae (Townsend), Lespesia archippivora (Riley), Lixophaga variablis (Coquillet), Nemorilla pyste (Walker), Patelloa leucaniae (Coquillet), Pseudoperichaeta erecta (Coquillet), Stomatomyia parvipalpis (Wulp), and Winthemia quadripustulata (Fabricius) (all Diptera: Tachinidae). Life Cycle and Description. A complete life cycle requires about 40 days. The number of annual generations is thought to be 3–4 throughout its range. In Texas there are

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four flights of adults: they occur in May, late June-early July, early August, and mid-September. However, because of overlapping generations, it is difficult to discern separate flights. Egg. The oval eggs initially are nearly transparent, becoming cream or yellowish in color. They measure about 0.64 mm wide and 1.1 mm long. The flattened, overlapping eggs are deposited in clusters of 8–20, usually on the underside of foliage. Total fecundity is estimated at 300–400 eggs. Duration of the egg stage is 2–5  days but averages 2.8 days. Larva. The larva generally is pale-green dorsally and yellowish-green ventrally. It measures 21–24 mm long at maturity. The head and thoracic plate are yellowish but marked with dark spots. The thoracic plate markings are in the form of two dark bars on each side. The body is well-marked with raised, dark spots enclosing a paler area from which emerges a stout black hair. This is especially evident in the lateral spots below the spiracles, which appear almost ring-like. There are six such spots on each abdominal segment. A light mid-dorsal stripe occurs dorsally along the center of the abdomen. Mean head capsule widths (range) are 0.25 (0.20–0.26), 0.37 (0.32–0.40), 0.58 (0.55–0.70), 0.92 (0.80–1.00), 1.10 (1.05–1.20), and 1.34 (1.25–1.50) mm, respectively, for instars 1–6. Mean overall body lengths (range) are 2.1 (1.6–2.5), 3.7 (2.2–4.3), 6.1 (4.0–9.3), 9.5 (6.6–13), 11.0 (9–14), and 13.4 (9–20) mm, respectively. During the first instar, larvae normally are gregarious but disperse afterwards. If disturbed, larvae wriggle violently, drop from the plant on a strand of silk, or rapidly retreat within a silken tube, if one is present. Duration of the larval stage requires 14–28 days, averaging 16 days and is followed by a prepupal period of 1–3 days. Pupa. The mature larva pupates in the soil within a silklined cell, usually under debris or near the soil surface. The pupal case measures about 12.5 mm long and 3.5 mm wide and is encrusted with soil. It is closed at the bottom but open at the tip, allowing easy escape of the moth. The pupa is light to dark brown and measures about 8.0–9.5 mm long. The tip of the abdomen bears three stout spines. Duration of the pupal stage requires 4–13 days, averaging 8.7 days. Adult. This small moth has a wingspan of about 17–43 mm, males averaging smaller than females. It is yellowish brown or reddish brown but bears lighter and darker markings. The forewing and hind wing usually are crossed by a light irregular band, and sometimes a dark band. The hind wing is yellowish. The moth is quite variable in coloration, which may account for the many times it has been described as a new species. Adults feed on nectar from various flowers and commence oviposition about 3–6 days after emergence.

Elements of garden webworm biology were given by Sanderson (1906), Sanborn (1916), Kelly and Wilson (1918), Poos (1951), and Smith and Franklin (1954). Capps (1967) provided a description of several stages. Keys to some webworm larvae, including garden webworm, were published by Allyson (1976) and Sparks and Liu (2001). A key to the adults was provided by Munroe (1976).

Damage Larvae feed only on the leaf epidermis during the first two instars, skeletonizing the tissue. Thereafter they consume the entire leaf. Eventually, they defoliate plants, consuming all except the stems and major veins. Webworms usually wrap young leaves in a loose web and feed within the protection of the web. During periods of abundance, entire plants are shrouded in webbing. This webbing is not diagnostic, however, because beet webworm and alfalfa webworm also display this behavior.

Management Infestations often result from the presence of weeds in crop fields. Destruction of weeds can deter oviposition. The other major source of webworms is alfalfa fields, as adults disperse at maturity and larvae disperse when the alfalfa is cut. Early harvesting of alfalfa, especially if the larvae are young and incapable of long-distance dispersal in the absence of food, can reduce webworm numbers. Alfalfa can also be treated with insecticide before harvest if it is heavily infested. Insecticidal suppression of webworms is normally accomplished by foliar applications, though this is seldom warranted. The bacterium Bacillus thuringiensis provides some control. Populations of adults can be monitored using blacklight traps. Adult garden webworms can also be captured in traps baited with a blend of phenylacetaldehyde, methyl-2-methoxybenzoate, methyl salicylate, and β-myrcene (Landolt et al., 2011b).

Hawaiian Beet Webworm

Spoladea recurvalis (Fabricius)

Spotted Beet Webworm

Hymenia perspectalis (Hübner)

Southern Beet Webworm

Herpetogramma bipunctalis (Fabricius) (Lepidoptera: Crambidae)

Natural History Distribution. Hawaiian beet webworm, Spoladea recurvalis (Fabricius), is found throughout the world in tropical and subtropical regions. In North America, it is found in the southern states from Florida west to California and

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Hawaii, and also in Puerto Rico. It sometimes causes damage as far north as Virginia, but it cannot overwinter there and must reinvade annually; thus it inflicts injury only late in the season. Its origin is uncertain, but it is not native to North America. Spotted beet webworm, Hymenia perspectalis (Hübner), and southern beet webworm, Herpetogramma bipunctalis (Fabricius), are closely related to Hawaiian beet webworm. They are not well known, but also are tropical pests with a wide geographic range. In the United States, they similarly are southern insects but can occasionally cause injury as far north as Illinois and Virginia. Host Plants. Hawaiian beet webworm is largely restricted to plants in the family Chenopodiaceae. Among vegetable crops, beet, chard, spinach, Indian spinach, and New Zealand spinach are normally injured. Sugarbeet is readily attacked, but this crop is rarely grown in the warm environments favored by Hawaiian beet webworm. When they face starvation, this insect may feed on other crops, but it is much less damaging than the better-known webworms—beet webworm, Loxostege sticticalis (Linnaeus); alfalfa webworm, L. cereralis (Zeller); and garden webworm, Achyra rantalis (Guenée). Weed hosts are pigweed, Amaranthus spp., lambsquarters, Chenopodium album, and purslane, Portulaca oleracea. Pigweed is preferred even over cultivated hosts. Spotted beet webworm and southern beet webworm display the same dietary preferences as does Hawaiian beet webworm. Tingle et al. (1978) indicated that southern beet webworm was the dominant caterpillar on Amaranthus hybridus in Florida cornfields during the late summer months. Natural Enemies. Hawaiian beet webworm is known to be parasitized by Cotesia marginiventris (Cresson) (Hymenoptera: Braconidae), Venturia infesta (Cresson) (Hymenoptera: Ichneumonidae), Argyrophylax albincisa (Weidemann), Chaetogaedia monticola (Bigot), Eucelatoria armigera (Coquillett), and Nemorilla pyste (Walker) (all Diptera: Tachinidae). Not surprisingly, the parasitoid complex attacking spotted and southern beet webworms are similar to Hawaiian beet webworm (Tingle et  al., 1978). Venturia infesta (Cresson) (Hymenoptera: Ichneumonidae) has been reared from all three webworms. Also, Apanteles mimoristae (Muesebeck) (Hymenoptera: Braconidae) is associated with spotted beet webworm, whereas Gambrus ultimus (Cresson), Temelucha sp. (both Hymenoptera: Ichneumonidae) and C. marginiventris (Hymenoptera: Braconidae) attack southern beet webworm. Argyrophylax albincisa (Weidemann) is associated with spotted beet webworm and Nemorilla pyste (Walker) with southern beet webworm (both Diptera: Tachinidae). Life Cycle and Description. In Hawaii, Hawaiian beet webworm is active throughout the year and about 10 generations occur annually. During warm weather, one generation

is completed in about 30 days. The biologies of spotted and southern beet webworms are less certain but seem to be basically the same as Hawaiian beet webworm except as noted below. Dispersal behavior of Hawaiian beet webworm has been studied extensively in Asia, where adults disperse long distances from favorable (warm winter) overwintering areas to unfavorable (cold winter) areas annually. Moths fly readily during the first few days of the adult stage. Feeding is not necessary before flight commences, but longevity, flight duration, and flight velocity increase if they have access to honey. Mating status and reproductive status do not affect flight (Shirai, 2006). Egg. The egg is elliptical, iridescent white, and flattened, and may be deposited singly or in rows of up to several eggs. The egg measures 0.6 mm long, 0.4 mm wide, and 0.25 mm in height. They most often are laid on the lower surface of leaves adjacent to large veins. Duration of the egg stage is about 4 days.

FIG. 10.19  Hawaiian beet webworm larva. (Photo by L. Buss.)

FIG. 10.20  Southern beet webworm larva. (Photo by L. Buss.)

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Larva. Young larvae of Hawaiian beet webworm feed beneath the leaves, and occasionally spin light webs in which they rest. The larvae initially are pale green, though a few dark spots are found on the head and thoracic plate. As larvae attain maturity they develop broad, irregular, whitish lateral stripes that contrast strongly with a dark stripe dorsally. The larvae also become uniformly pinkish or rust colored as they prepare to pupate. The head capsule increases in width from 0.25 to 1.2 mm as the larva grows. The body bears numerous stout hairs over the length of its body but lacks the dark spots or rings found with such hairs on many webworms. The larval development period is normally 9–13 days. Larvae of spotted beet webworm also are green, but as its common name suggests, bear numerous spots. The thorax and abdomen are equipped with raised-dark spots from which arise dark hairs. The head bears purplish dots, though the center is unmarked. The thoracic plate has black borders. Larvae of southern beet webworm are dark-green with a dark, nearly black, head. The thoracic plate is similarly dark except for the central area. The body bears numerous large, dark, raised spots from which arise black hairs. The spots are light in color centrally, however, resulting in a ring-like appearance and causing this insect to resemble garden webworm, Achyra rantalis (Guenée).

are marked with a white transverse band that nearly crosses the wings. The forewing also bears one elongate and two small white spots distally. The margin of the front wings is alternating dark and light. Narrow light bands are found on the abdomen. The wingspan measures 17–23 mm.

FIG. 10.22  Adult of spotted beet webworm. (Drawing by USDA.)

Spotted beet webworm is similar to Hawaiian beet webworm in general appearance. The spotted species is lighter brown, however, with a reddish tint, and the white wing markings are less discrete and pronounced. The wingspan is about 20 mm.

Pupa. Mature larvae drop to the soil to pupate. They burrow slightly beneath the soil surface and form firm, compact, elliptical cocoons of silk covered with grains of soil. The pupa is light brown and the posterior end is equipped with terminal spines bearing hooked tips. The pupa measures about 10 mm long and 2.5 mm wide. Duration of the pupal stage is 7–14 days. FIG. 10.23  Adult of southern beet webworm. (Drawing by USDA.)

Southern beet webworm moths are light yellowish gray, sometimes with an iridescent purplish cast. The front wings bear three dark spots along the leading edge, but the markings are not very distinct. The hind wings are not distinctly marked. The abdomen is darker. The wingspan is 22–26 mm. The biology of Hawaiian beet webworm was given by Marsh (1911) and Chittenden (1911b) and Walker and Anderson (1939). Spotted beet webworm was described by Davis (1912) and Chittenden (1913a). Southern beet webworm was described by Chittenden (1911a). Sparks and Liu (2001) include Hawaiian beet webworm in their key to ­vegetable-infesting larvae. FIG. 10.21  Adult of Hawaiian beet webworm. (Photo by L. Buss.)

Damage Adult. The moth of Hawaiian beet webworm is dark brown, and often tinted purple. The front and hind wings

Young larvae remain on the lower surface of foliage, not eating entirely through the leaves. When nearly mature,

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however, they consume the entire leaf. Unlike some other webworm species, webbing is not very pronounced.

Management Infestations often result from the presence of weeds in crop fields. Destruction of weeds can deter oviposition. Insecticidal suppression of webworms is normally accomplished by foliar applications, though this is seldom warranted except, perhaps, for Hawaiian beet webworm in Hawaii. The bacterium Bacillus thuringiensis provides some control.

Melonworm

Diaphania hyalinata Linnaeus

Cucumber Moth

Diaphania indica (Saunders) (Lepidoptera: Crambidae)

Natural History Distribution. Melonworm is most abundant in tropical climates, where cucurbits can grow during the winter months. It occurs throughout most of Central and South America and the Caribbean. The United States is the northern limit of its permanent range, and its wintertime occurrence is generally limited to south Florida and perhaps south Texas. Melonworm disperses northward annually. Its distribution during the summer months is principally the southeastern states, extending west to Texas and, sometimes, north to New England and the Great Lakes region. Melonworm has been reported in North Carolina as early as July (Smith, 1911). In recent years it has also been reported to be a serious pest in Sudan, Africa (Mohaned et al., 2013), although this may be a misidentification; it is more likely that this is cucumber moth, Diaphania indica (Saunders). Cucumber moth is found widely in warm areas of Asia and Africa, including northern Australia, where it is also known as pumpkin caterpillar, melon borer, cotton caterpillar, and other names. Its scientific name is occasionally given as Palpita indica (Saunders). Its distribution is also said to include Central America, Florida, Texas, and the Caribbean region (Heppner, 2003; Solis, 2006; Gilligan and Passoa, 2014). However, based on extraneous morphology, cucumber moth is nearly indistinguishable from melonworm, so these distribution records may not be accurate. Host Plants. Melonworm is restricted to feeding on cucurbits. Summer and the winter squash, cantaloupe, and cucumber (usually Cucumis spp.) are generally satisfactory hosts. Pumpkin (usually Cucurbita pepo) is of variable quality as a host, probably because pumpkins are bred from several Cucurbita species. Watermelon

(Citrullus lanatus) is generally not injured by this insect. Guillaume and Boissot (2001) reported variability in Cucumis genotypes to D. hyalinata. Interestingly, Panthi et al. (2016) compared oviposition, growth, and survival of melonworm on yellow squash, zucchini squash, cucumber, and watermelon. In most respects, the squash plants were better hosts. However, watermelon was curiously suitable for larvae, but not for oviposition. Weed hosts are unknown. Cucumber moths similarly consume cucumber (Cucumis sativus), gourd (Lagenaria siceraria), wax gourd (Benincasa hispida), watermelon (Citrullus lanatus and Ci. vulgaris), oriental melon (Cucumis melo), and other Cucurbitaceae (Ke et al., 1988). In addition, their diet is sometimes said to include noncucurbits such as cotton (Gossypium indicum; Malvaceae), beans (Phaseolus sp.; Fabaceae), and pigeon pea (Cajanus cajan; Fabaceae). However, host records are often based on spurious or casual observations, often made during periods of great insect abundance when insect behavior is not normal. Until further careful research is conducted, D. indica should be considered only as a cucurbit pest. Consistent with this, Pandey (1977) assessed 34 plants from 10 families and reported that only the cucurbits were eaten. Interestingly, there are no records of damage by D. indica to cucurbits in North America, and because it would be exceedingly easy to confuse the two species, it may be that D. hyalinata has been mistaken for D. indica. Natural Enemies. Natural enemies of melonworm are nearly the same as those of pickleworm. All of the parasitoids found by Pena et  al. (1987b) to attack pickleworm also attacked melonworm: Apanteles sp., Hypomicrogaster diaphaniae (Muesebeck), Pristomerus spinator (Fabricius) (all Hymenoptera: Braconidae), Casinaria infesta (Cresson), Temelucha sp. (both Hymenoptera: Ichneumonidae), and undetermined trichogrammatids (Hymenoptera: Trichogrammatidae) (Pena et  al., 1987b; Capinera, 1994). However, additional species parasitize melonworm, including Gambrus ultimus (Cresson), Agathis texana (Cresson) (both Hymenoptera: Ichneumonidae) and an undetermined tachinid fly (Hymenoptera: Tachinidae). The tachinids known from melonworm are Nemorilla pyste (Walker) and Stomatodexia cothurnata (Wiedemann). Studies conducted in Puerto Rico (Medina-Gaud et al., 1989) reported levels of parasitism reaching 24%. Generalist predators such as Calosoma spp. and Harpalus (both Coleoptera: Carabidae), the soldier beetle Chauliognathus pennsylvanicus De Geer (Coleoptera: Cantharidae), and the red imported fire ant Solenopsis invicta Buren (Hymenoptera: Formicidae) have also been reported to be mortality factors. In Brazil, Polybia ignobilis (Haliday) and P. scutellaris (White) (Hymenoptera: Vespidae) were especially

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important predators of melonworm, attacking all larval instars except the first (Santana Jr. et  al., 2012). In addition, these authors reported that other important predators were flower flies (Diptera: Syrphidae), minute pirate bugs (Hemiptera: Anthocoridae), ladybird beetles (Coleoptera: Coccinellidae), and other generalist predators were ­responsible for mortality. In addition, levels of egg mortality reaching 90% were caused by the ant Paratrechina sp. (Hymenoptera: Formicidae), although the key mortality factor was parasitism by tachinid flies (Diptera: Tachinidae) (Gonring et al., 2003). Also in Brazil, the pupal parasitoid Trichospilus diatraeae Cherian (Hymenoptera: Eulophidae) was shown to be quite effective (Da Silva et al., 2015). Naturally occurring diseases of melonworm are not known. Cucumber moths have a number of documented natural enemies. Peter and David (1991b) documented 16 parasitoids, 3 predators, and a fungal (microsporidian) disease agent from one state in India. These authors (Peter and David, 1991a) also indicated that the braconid Apanteles taragamae Viereck was the key mortality factor, but also reported parasitism by Apanteles machaeralis Wilkinson, Phanerotoma hendecasisella Cam. (both Hymenoptera: Braconidae), Goniozus sensorius Gordh (Hymenoptera: Bethylidae), Trathala flavoorbitalis (Cam.) (Hymenoptera: Ichneumonidae), and Elasmus brevicornis Gahan (Hymenoptera: Elasmidae). Visalakshy (2005) noted three additional parasitoids of cucumber moth, namely Trichogramma chilonis Ishii (Hymenoptera: Trichogrammatidae), Dolichogenidea stantoni (Ashmead) (Hymenoptera: Braconidae), Xanthopimpla punctata (Fabricius) (Hymenoptera: Ichneumonidae), and the entomopathogenic fungus Metarhizium (Nomurea) rileyi. Life Cycle and Description. The melonworm can complete its life cycle in about 30  days. It is present throughout the year in southern Florida, where it is limited mostly by availability of host plants. It disperses northward annually, usually arriving in northern Florida and other southeastern states in June or July, where not more than three generations normally occur before cold weather kills the host plants. The biology of D. indica seems to be quite similar to D. hyalinata, though they have never been reared under exactly the same ­conditions so comparisons are difficult. The following insect description is based on melonworm except where noted. Egg. Melonworm moths deposit oval, flattened eggs in small clusters, averaging 2–6 eggs per mass. Apparently, they are deposited at night on buds, stems, and the underside of leaves. Initially, they are white but soon become yellow. They measure about 0.7 mm long and 0.6 mm wide. Hatching occurs after 3–4 days.

FIG. 10.24  Larva of melonworm. (Photo by L. Buss.)

Larva. There are five instars. Total larval development time is about 14  days, with mean (range) duration of the instars about 2.2 (2–3), 2.2 (2–3), 2.0 (1–3), 2.0 (1–3), and 5.0 (3–8) days, respectively. Head capsule widths are about 0.22, 0.37, 0.62, 1.04, and 1.64 mm, respectively (Smith et  al., 1994). Larvae attain lengths of about 1.5, 2.6, 4.5, 10, and 16 mm in instars 1–5, respectively. Neonate larvae are colorless, but by the second instar, larvae assume a pale yellow-green color. They construct a loose silken structure under leaves that serve to shelter them during the daylight hours. In the fifth instar, larvae have two subdorsal white stripes extending the length of the body. The stripes fade or disappear just before pupation, but they are the most distinctive characteristic of the larvae. Smith (1911) provided a complete description of larvae. Pupa. Before pupation, larvae spin a loose cocoon on the host plant, often folding a section of the leaf for added shelter. The melonworm cocoon is much better formed than the cocoon of pickleworm, and the melonworm’s preference for green foliage as a pupation site also serves to differentiate the insects. The pupa is 12–15 mm long, about 3–4 mm wide, and fairly pointed at each end. It is light to dark brown. The pupal stage persists for 9–10 days.

FIG. 10.25  Adult of melonworm. (Photo by L. Buss.)

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Adult. The moth’s wingspan is about 2.5 cm. The wings are pearly white centrally, and slightly iridescent, but are edged with a broadband of dark brown. Moths frequently display brushy hair pencils at the tip of the abdomen when at rest. Melonworm moths remain in the crop during the daylight hours, and though generally inactive during the day, they fly short distances when disturbed. Characters used to distinguish adult melonworm moth from adult cucumber moth are the width of the dark band at the wing margin, and the color of the terminal abdominal segments viewed dorsally. In melonworm, the dark marginal bands of the wings are relatively narrow, and the two terminal segments are dark (all other segments are white). In contrast, in cucumber moth, the wing bands are slightly broader, and although the second to the last abdominal segment is dark, the terminal (and all other except the subterminal segment) are normally white. More definitive determination of the species requires dissection of the abdomen and examination of the genitalia. Smith (1911) provided a detailed account of melonworm biology. Rearing techniques were given by Elsey et al. (1984) and Valles et al. (1991). Accounts of cucumber moth biology are presented by Hosseinzade et al. (2014a, b) and Nagaraje et al. (2018).

Damage Melonworm feeds principally on foliage, especially if foliage of a favored host plant such as summer or winter squash is available. Usually, the leaf veins are left intact, resulting in lace-like plant remains. However, if the available foliage is exhausted, or the plant is a less preferred plant such as cantaloupe, the larva may feed on the stems or the surface of the fruit, or even burrow into the fruit. As is the case with pickleworm, growers sometimes refer to these insects as “rindworms,” because they cause scars on the surface of melons. Cucumber moth larvae feed in the same manner, preferring foliage but feeding on other organs of the plant if necessary. In a study of melonworm damage potential to summer squash conducted in south Florida, melonworm caused a 23% yield loss due to foliage damage (indirect loss) and a 9–10% yield reduction owing to fruit damage (direct loss) (McSorley and Waddill, 1982). Kelsheimer (1949) considered this insect to be the most important pest of cucurbits in Florida. However, it is easy to kill with modern insecticides, so its status as a key cucurbit pest has been supplanted by pickleworm, which feeds internally and therefore is more difficult to reach with most insecticides.

Management Sampling. Pheromone production by female moths peaks at about sunset (Valles and Capinera, 1992). The sex pheromone of melonworm has been identified (Raina et al., 1986),

but is not available commercially. It is also known for cucumber moth (Choi et al., 2009) and is commercially available. Moths are not attracted to light traps. Larval abundance is usually assessed visually, though beating a leaf from the median third of the cucurbit vine is recommended by Bacci et al. (2006) for melonworm. Insecticides. Historically, melonworm was considered to be a very damaging pest, but because it feeds preferentially on foliage it is easy to control with a number of insecticides. In tropical areas, it is often considered more damaging than pickleworm. In temperate areas, and especially in commercial vegetable production areas, it is treated as only a minor pest. In insecticide-free cucurbit production and in home gardens, melonworm can cause serious damage. Although not routinely used, botanical insecticides can have significant effects on melonworm. Silva et al. (2015), working in Brazil, showed that citronella oil and eucalyptus oil were toxic to D. hylinata. Andiroba oil, eucalyptus oil, garlic extract, and citronella oil inhibited feeding by larvae, and andiroba oil, eucalyptus oil, garlic extract, and rotenone inhibited oviposition. The authors concluded that eucalyptus oil and citronella oil were most promising for incorporation into melonworm pest management programs. Pollinators, particularly honeybees, are very important in cucurbit production, and insecticide application can interfere with pollination by killing honeybees. If insecticides are to be applied when blossoms are present, it is advisable to use insecticides with little residual activity and to apply insecticides late in the day, when honeybee activity is minimal. Biological Control. In addition to chemical insecticides, Bacillus thuringiensis is commonly recommended for suppression of melonworm. The entomopathogenic nematode Steinernema carpocapsae provides only moderate suppression because the nematodes do not survive long on the foliage, where larvae are found resting and feeding (Shannag and Capinera, 1995). Soumya et  al. (2017) studied biological suppression of cucumber moth in India and reported that Bacillus thuringiensis, or a combined release of two parasitoids (Trichogramma chilonis and Dolichogenidea stantoni) gave better protection than several other biological suppressants. In these studies, the release of D. stantoni alone, or application of Metarhizium (Nomurea) rileyi, or Beauveria bassiana, gave moderate suppression. Less successful were release of T. chilonis alone or application of Metarhizium anisopliae. Cultural Practices. Row covers can be used effectively to exclude melonworm adults (Webb and Linda, 1992), and presumably cucumber moths. Intercropping of corn and beans with squash was shown to reduce damage by melonworm (Letourneau, 1986). Because melonworm prefers squash to most other cucurbits, trap cropping has been suggested, and of course, destruction of crop residue which

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may contain melonworm pupae is recommended (Smith, 1911). Early plantings, except in tropical areas where melonworm overwinters, often escape serious damage.

Pickleworm

Diaphania nitidalis (Stoll) (Lepidoptera: Crambidae)

Natural History Distribution. Pickleworm is a tropical insect that occurs widely in Central and South America and the Caribbean. In the United States, it routinely survives the winter only in south Florida and perhaps south Texas. Pena et al. (1987a) documented the overwintering biology in south Florida, but overwintering has been observed as far north as Sanford, in central Florida, during mild winters. Pickleworm is highly dispersive and invades much of the southeast each summer. North Carolina and South Carolina regularly experience crop damage by pickleworm, but often this does not occur until August or September. In contrast, northern Florida and southern Georgia are flooded with moths each year in early June as warm, humid tropical summer weather conditions become firmly established. Although it regularly takes 1 or 2 months for the dispersing pickleworms to move north from Florida to the Carolinas, in some years they reach locations as far north as Michigan and Connecticut. Presumably, they are assisted in their northward dispersal by favorable wind patterns. However, there is some evidence that picklworms can also overwinter in particularly mild coastal areas of other southeastern states (M. Jackson, pers. comm.). In Canada, pickleworm has occasionally been found in southern Ontario. In Puerto Rico, it is more common in the mountains than at low elevations and is not found at all in dry areas of the island (Wolcott, 1948). Host Plants. Pickleworm feeds only on cucurbits, but both wild and cultivated species are suitable hosts. Creeping cucumber, Melothria pendula, is considered to be an important wild host. Wild balsam apple, Mormordica chorantia, which has also been reported to be a host, is of questionable significance (Elsey et al., 1985). Summer and the winter squash species are good hosts. Pumpkin is considered of variable quality as a host, probably because pumpkins are bred from several Cucurbita species. The Cucumis species—cucumber, gerkin, and cantaloupe—are attacked but not preferred. Among all cucurbits, summer squash is most preferred and most heavily damaged. Cultivars vary widely in susceptibility to attack, but truly resistant cultivars are unknown (Dilbeck et al., 1974). Cucurbits are intolerant of cold weather. Although diapause is unknown in pickleworm, it is the lack of host plants during the winter months that functionally limits the distribution of pickleworm. Natural Enemies. Pickleworm has several natural enemies in North America, but none reliably suppress damage. Generalist predators such as Calosoma spp. and

Harpalus (both Coleoptera: Carabidae); the soldier beetle Chauliognathus pennsylvanicus De Geer (Coleoptera: Cantharidae); and the red imported fire ant, Solenopsis invicta Buren (Hymenoptera: Formicidae); have been reported to be important mortality factors. Also, several parasitoids are known, including Apanteles sp., Hypomicrogaster diaphaniae (Muesebeck), Pristomerus spinator (Fabricius) (all Hymenoptera: Braconidae); Casinaria infesta (Cresson), Temelucha sp. (both Ichneumonidae); and undetermined trichogrammatids (Pena et al., 1987b; Capinera, 1994). The braconid Cardiochiles diaphaniae Marsh (Hymenoptera: Braconidae) has been imported from Colombia and released into Florida and Puerto Rico in an attempt to obtain higher levels of parasitism (Smith et al., 1994). A study in Brazil found that predation by Paratrechina sp. ants (Hymenoptera: Formicidae) and parasitism by Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae) were the most important factors during the egg stage. During the larval stage, Polybia ignobillis Halliday wasps (Hymenoptera: Vespidae) and rainfall were critical mortality factors. During the pupal stage, Labidus coecus (Latreille) ants (Hymenoptera: Formicidae) were important mortality agents (Gonring et al., 2002). Life Cycle and Description. The pickleworm can complete its life cycle in about 30 days. Over much of its range, multiple and overlapping generations may occur. The number of generations was estimated to be four in Georgia (Dupree et  al., 1955) and two or three in North Carolina (Fulton, 1947). Egg. The eggs are minute, measuring only about 0.4–0.6 mm wide and 0.8 mm long. The shape varies from spherical to flattened. Their color is white initially, but changes to yellow after about 24 h. The eggs are distributed in small clusters, usually 2–7 per cluster. They are deposited principally on the buds, flowers, and other actively growing portions of the plant. Hatching occurs in about 4 days (Smith, 1911). Elsey (1980) estimated egg production to be 300– 400 per female.

FIG. 10.26  Immature larva of pickleworm. (Photo by J. Capinera.)

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FIG. 10.27  Mature larva of pickleworm. (Photo by J. Capinera.)

Larva. There are five instars. Total larval development time averages 14 days. Mean duration (range) of each instar is about 2.5 (2–3), 2 (1–3), 2 (1–3), 2.5 (2–3), and 5 (4–7) days, respectively. Head capsule widths for the five instars are about 0.25, 0.42, 0.75, 1.12, and 1.65 mm, respectively (Smith et al., 1994). Body lengths average 1.6, 2.5, 4.0, 10, and 15 mm during instars 1–5, respectively. Young larvae are nearly white with numerous dark gray or black spots. The dark spots are lost at the molt to the fifth instar. Larval color during the last instar is somewhat variable, depending largely on the insect’s food source. For example, they tend to be orange when feeding on blossoms, green when feeding on stem tissue, and white when feeding on fruit. Before pupation, larvae tend to turn a dark copper color. When mature, larvae often attain a length of 2.5 cm. Smith (1911) provided a good description of the larval instars. Pupa. Pupation usually occurs in a leaf fold; often dead, dry material is used to make the pupal shelter. There is only weak evidence of a cocoon, usually just a few strands of silk. The pupa is elongate, measuring about 13 mm long and 4 mm wide. It is light brown to dark brown and tapers to a point at both ends. Pupation usually lasts about 8–9 days.

The female moth produces a pheromone that attracts males, with peak production occurring at 5–7 h after sunset (Klun et al., 1986; Elsey et al., 1989; Valles et al., 1992). Moths are fairly distinctive in appearance. The central portion of both the front and hind wings is a semitransparent yellow color, with an iridescent purplish reflection. The wings are bordered in dark brown. The wing expanse is about 3 cm. Both sexes often display brushy hair pencils at the tip of the abdomen. Moths are not found in the field during the daylight hours and probably disperse to adjacent wooded or weedy areas during the heat of the day. Moths do not produce eggs until they are several days old. Good sources of information on pickleworm biology were supplied by Dupree et  al. (1955), Fulton (1947), Quaintance (1901), and Smith (1911). Rearing techniques were provided by Elsey et al. (1984), Robinson et al. (1979), and Valles et al. (1991).

Damage Pickleworm may damage summer and winter squash, cucumber, cantaloupe, and pumpkin. Watermelon generally is not a host. The blossom is a favored feeding site, especially for young larvae. In plants with large blossoms, such as summer squash, larvae may complete their development without entering the fruit. They may also move from blossom to blossom, feeding and destroying the plant’s capacity to produce fruit. Very often, however, the larva burrows into the fruit. The larva’s entrance is marked by a small hole, through which frass is extruded. The presence of the insect makes the fruit unmarketable, and fungal or bacterial diseases often develop once entry has occurred. If larvae burrow into fruit just before harvest, their presence is difficult to detect, yet a considerable amount of larval growth and feeding damage may occur. When all blossoms and fruit have been destroyed, larvae attack the vines, especially the apical meristem. Cantaloupe is not a preferred host, and larvae often seem reluctant to burrow into the fruit. Rather, they feed on the surface or “rind,” causing scars. Thus, pickleworm is sometimes referred to as “rindworm.”

Management

FIG. 10.28  Adult of pickleworm. (Photo by J. Capinera.)

Adult. Emerging moths fly during much of the evening hours, but most flight occurs 3–5 h after sundown, with peak flight at approximately midnight (Valles et al., 1991).

Sampling. It is very difficult to scout for this insect and predict its appearance. Moths are not attracted to light traps, and pheromone traps have had limited success (Elsey et al., 1991; Valles et al., 1991). Pheromone lures are not currently available commercially. Brewer and Story (1987) developed sampling plans for pickleworm larvae in squash. They suggested that the most reliable sampling unit was the large green staminate flower bud. However, the small eggs, nightflying behavior, and inability to trap the insect reliably have led most growers to depend on preventative applications of insecticides.

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Insecticides. Cucurbit producers in areas where pickleworm damage is likely to occur usually apply chemical insecticides from the onset of fruiting through harvest. The internal feeding behavior of larvae, which is so difficult to detect at harvest, causes particular emphasis on prevention of damage. In areas that are on the fringe of the normal range, there are many seasons when damage does not occur, but producers apply insecticides as a preventative measure because the prediction of occurrence is so difficult. Botanical insecticides can be effective (Arant, 1942). Pollinators, particularly honeybees, are very important in cucurbit production, and insecticide application can interfere with pollination by killing honeybees. If insecticides are to be applied when blossoms are present, it is advisable to use insecticides with little residual activity and to apply late in the day, when honeybee activity is minimal. Biological Control. The entomopathogenic nematode Steinernema carpocapsae has been shown to effectively suppress pickleworm injury in squash (Shannag et al., 1994; Webb and Capinera, 1995). Nematode survival is quite good in large-blossomed squash, where the nematodes can kill the young pickleworm before it burrows into the fruit. Nematode-based control can rival the use of conventional insecticides (Webb and Capinera, 1995). The nematodes can persist for 4 days on foliage, and survive in both mature and immature flowers. Bacillus thuringiensis also can kill pickleworm, but frequent applications must be made so that the larva ingests the bacterium and its toxin as it chews its way into the bloosom or fruit; once inside the plant, the larva is not likely to be affected. Although no picklewormspecific entomopathogens are known, multiply embedded viruses from velvetbean caterpillar (AgMNPV) and alfalfa looper (AcMNPV) are effective on pickleworm larvae (Jackson et al., 2008). However, presently they are not commercially available. Cultural Practices. It is possible to cover plants with the screen or row covers to prevent moths from depositing eggs on the foliage (Webb and Linda, 1992). However, as the plants must be pollinated, usually by honey bees, some allowance must be made to leave the plants uncovered. Given the night-flying behavior of the moths and the daytime activities of honeybees, this is not a difficult task on a small planting, but cost-prohibitive on large acreage. Some growers are able to prevent plant injury through careful timing of their cropping cycle. By planting early, it is often possible to harvest part of the crop before pickleworms appear. Usually, the crop is eventually infested, so some yield is lost. Plowing under the crop residue is recommended to destroy pupae in the leaf debris (Smith, 1911). The presence of aluminum or polyethylene mulch of various colors was shown by Dupree (1973) not to influence pickleworm damage to squash. Wolfenbarger and Moore (1968) reported that white, black, and aluminum mulches did not reduce pickleworm infestation of squash

as ­compared to unmulched crop, but that white mulch produced lower levels of fruit injury than aluminum mulch. In a comparison of monocultural and polycultural crop production systems conducted by Letourneau (1986), no difference in abundance of pickleworm on squash was observed. In the same study, distribution of melonworm, Diaphania hyalinata (L.) was significantly lower in polycultures, a common response for an insect with a restricted host range. The differential response between the two species is likely due to the more active, dispersive nature of pickleworm. Smith (1911) reported that squash could be used as a trap crop to keep pickleworm from attacking cantaloupe, a less preferred host. He recommended that destruction of squash blossoms, or even the entire plant, can be done periodically to keep pickleworms from exhausting the food supply and then moving onto adjacent cantaloupes. In contrast, Dupree et al. (1955) reported unsatisfactory results with trap cropping. More recently, Leiner and Spafford (2016a) showed that squash was a preferred oviposition site, relative to watermelon and cantaloupe, and suggested that squash could be used as part of a push-pull cropping system, pulling the moths away from less preferred crops. Also, the larvae did not distinguish between the three crops (Leiner and Spafford, 2016b), so it is the female that is making the choice for the larvae, but if the plants are in contact with one another the larvae could easily redistribute and do more damage than might be expected based on female moth choice.

Purplebacked Cabbage Worm Evergestis pallidata (Hufnagel) (Lepidoptera: Crambidae)

Natural History Distribution. This is a decidedly northern species and is found throughout most of Canada except for British Columbia, and as far south as Virginia and Kentucky in the eastern United States, and northern Arizona in the West. It is likely of European origin but has been in North America at least since 1869. It is recorded as a pest principally in Canada’s Maritime Provinces. Host Plants. Purplebacked cabbage worm feeds on a variety of cruciferous plants, including broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, horseradish, kale, kohlrabi, radish, rutabaga, and turnip. Horseradish and turnip seem to be most favored by this insect. Although moths deposit eggs on shepherdspurse, Capsella bursapastoris; and sheep sorrel, Rumex acetosella; larvae do not develop successfully on these plants. Natural Enemies. Few natural enemies are known. The wasps Bracon montrealis Morrison and Meteorus autographae Muesebeck (both Hymenoptera: Braconidae) have been reared from this caterpillar.

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Life Cycle and Description. There is only a single generation annually in Newfoundland, but two generations per year in Virginia. Overwintering occurs as a mature larva (prepupa) in the cocoon. Egg. The female deposits small batches of about 3–12 eggs on the underside of host plant foliage. The bright yellow, oval eggs are about 1.1 mm long and 0.8 mm wide, flattened, and overlap like fish scales. The eggs darken markedly just before hatching. Larvae hatch in 4–8 days. Larva. Newly hatched larvae are whitish green, and measure 1.5–2.0 mm long. The body bears numerous dark tubercles, each bearing one or more hairs. Head capsule widths are about 0.3, 0.5, 0.9, and 1.6 mm for the four instars, respectively. Duration of the instars is reported to be about 9.1, 8.9, 10.3, and 25.7 days, followed by a protracted prepupal period. Larvae are nocturnal and are usually found hiding between leaves during the day. Mature larvae are robust, bristly, and darker in color, normally olive-green to purple-brown, and measure about 20–22 mm long. There is a conspicuous yellow band on each side, with a narrow white band beneath, and the larva is colored ash-gray or greenish below. The body tapers at both the anterior and posterior ends. The larvae feed on the underside of leaves, then drop to the soil to prepare a cocoon. Pupa. The cocoon is oval, about 12–15 mm long, and covered with soil particles. The winter is passed as a prepupa in the cocoon, with pupation occurring in the spring. The pupa is light to dark brown. Adult. The moth escapes through a loosely constructed end of the cocoon. The moth has a wingspan of about 22–28 mm. The front wings are straw yellow, with irregular, narrow dark lines crossing the wing. The hind wings are whitish or pale yellow with darker margins. Morris (1958), Munroe (1973), and Howard et al. (1994) provided the biology of purplebacked cutworm.

Damage Larvae generally eat holes in the leaves, webbing them together, but also attack the crown and even the roots of such crops as rutabaga. Morris (1958) reported that it was a serious pest in Newfoundland, but its abundance varied widely from year to year.

Management Moths can be attracted to traps baited with phenylacetaldehyde, which offers the potential for population monitoring (Cantelo et  al., 1982). Chemical insecticides and Bacillus thuringiensis can be applied against the larvae, but this is usually a minor pest as compared to other crucifer-feeding

caterpillars. Spring tillage can destroy the cocoon and deep tillage can prevent the moths from emerging. Early planted turnip can be used as a trap crop to help protect cabbage and rutabaga.

Sod and Root Webworms

Crambus spp. and others (Lepidoptera: Pyralidae)

Natural History Distribution. There are several native pasture-dwelling webworms that occasionally damage crops. Commonly they are called sod webworms because they are usually associated with pasture and lawn grasses. They occur throughout the United States and southern Canada. Among the species known to cause damage are corn root webworm, Neodactria caliginosellus (Landry); silverstriped sod webworm, Crambus praefectellus (Zincken); larger sod webworm, Pediasia trisectus (Walker); and striped sod webworm, Fissicrambus mutabilis (Clemens). Food Plants. Sod and root webworms feed principally on pasture and sod grasses in the family Gramineae. However, if pasture or sod is tilled and the ground is planted to nongrass crops, they too may be injured by the residual webworm population. In addition to grasses such as bluegrass, corn, orchardgrass, rye, timothy, and wheat, some webworms have been known to attack alfalfa, cabbage, clover, mint, and tobacco. Consumption of the latter hosts is unusual. Weed grasses such as crabgass, Digitaria sanguinalis, and even some broad-leaf weeds such as sheep sorrel, Rumex acetosella, and aster, Aster ericoides, are consumed by larvae. Corn root webworm displays a particular preference for plantain, Plantago lanceolata, and oxeye daisy, Chrysanthemum leucanthemum. Natural Enemies. The natural enemies are not well known, but their impact is thought to be significant. Cockfield and Potter (1984) estimated a 75% reduction in eggs within 48 h due to predation. Among those thought to be important are the mite egg predators Hypoaspis sp., Cosmolaelaps sp. (both Acari: Laelapidae), and Parasitus sp. (Acari: Parasitidae); the ground beetles Anisodactylus rusticus Say, Amara cupreolata Putzeys, A. familiaris Duftschmidt, Calathus opaculus LeConte, and Stenolophus rotundata LeConte (all Coleoptera Carabidae); the rove beetles Meroneura venustula (Erichson), Neohypnus sp., Philonus sp., and Tachyporus jocosus Say (all Coleoptera: Staphylinidae) and ants, especially Phedole tysoni Forel. Birds also are common predators and where webworms are abundant the sod or soil often is heavily disturbed by birds probing for larvae. Flies are not uncommon parasitioids, including Aplomya caesar (Aldrich), A. confusionis (Sellars), and Stomatomyia floridensis (Townsend) (all Diptera: Tachinidae). Wasp parasitoids known from larger

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sod webworm include Macrocentrus crambi (Ashmead), M. crambivorus Viereck, Apanteles crambi Weed, Orgilus detectiformis Viereck (all Hymenoptera: Braconidae), and Diadegma obscurum (Cresson) (Hymenoptera: Ichneumonidae). Macrocentrus crambi and M. crambivorus have also been reared from corn root webworm. Species reared from striped sod webworm include Apanteles terminalis Gahan, A. ensiger (Say), and M. crambi (all Hymenoptera: Braconidae) and Campoletis argentifrons (Cresson) (Hymenoptera: Ichneumonidae). Life Cycle and Description. Following is a description of larger sod webworm, but the other sod webworm species are similar except for the larger size of P. trisectus, and the occurrence of a distinct silver stripe on the leading edge of the forewing in some species. Overwintering occurs in the larval stage. In Iowa and Tennessee, these larvae give rise to flights of moths in June, followed by additional generationproducing flights of moths in August. Light trap catches from Ontario indicate two flights of moths for numerous sod webworm species (Arnott Jr., 1934). Three generations are thought to occur for some sod webworms in the midwestern states, however. Egg. The eggs are dropped individually and randomly by females, while either at rest or while flying. The eggs are quite small, though large relative to other sod webworms. They measure 0.45–0.56 mm long and 0.31–0.36 mm wide. The eggs are white initially, and turn yellow with age. They are elongate-oval, with one end more broadly rounded than the other. Duration of the egg stage is 5–7 days. Dry air is lethal to eggs (Morrison et  al., 1972), but they tolerate a wide range of temperatures (Matheny and Heinrichs, 1971).

FIG. 10.29  Larger sod webworm larva. (Drawing by USDA.)

Larva. The larvae move downward upon hatching, usually hiding between the blades of grass. They produce silk readily, and if disturbed spin down on a silken thread. As the larvae increase in size they produce a silken tube or tubes beneath the surface of the soil, to which sand and soil particles adhere, as shelter. The larvae may leave the tube to feed, or if food is convenient it remains at least partially within the tube while feeding. The larvae may undergo 7–10 instars but normally display eight. Duration is about 3.0, 3.1, 3.0, 4.0, 3.6, 4.1, 5.9, and 9.9  days respectively, for instars 1–8. Head capsule widths average 0.21, 0.31, 0.45, 0.67, 0.99, 1.32, 1.55, and 2.20 mm and body lengths 1–2, 2.5–3.5, 3.7–5.5, 6.0–8.5, 10–12, 12–18, 18–24, and

21–28 mm, respectively, for instars 1–8. The young larvae are reddish brown with a blackish head capsule, but at instar four and thereafter the head becomes yellowish brown. The larvae bear reddish raised plates from which setae arise. The thoracic and caudal plates tend to be brownish or black until the final instar when they become lighter. Larvae that overwinter can do so in nearly any instar, but this is normally accomplished by instars 2–5. Pupa. The larvae abandon their feeding tubes at maturity and pupate in the soil nearby. The pupal cell is oval and lined with silk. It measures about 14 mm long and 6 mm wide. The pupa measures about 11 mm long and 3 mm wide. The cell is placed very near the soil surface so the moth has no trouble in escaping. Pupation may require 5–15 days, but 7–8 days is normal.

FIG. 10.30  Larger sod webworm adult. (Drawing by USDA.)

Adult. The moth’s front wings and body are yellowish gray or yellowish brown, the hind wings lighter and silvery at the base. A line of dark scales usually extends along the mid-line of the wing almost the entire length, but then turns up to the anterior tip. This moth measures about 21–35 mm in wingspan. Moths are nocturnal, but begin activity at dusk. They hide during the day, usually on the underside of broad-leaf weeds. They apparently require water or dew but have not been observed to feed on flowers. The longevity of adults is normally 7–10  days, though some individuals have survived for nearly a month. Females are believed to produce 200–250 eggs. The biology of corn root webworm was described by Runner (1914), striped sod webworm by Ainslie (1923a), silverstriped webworm by Ainslie (1923b), and larger sod webworm by Ainslie (1927). Forbes (1904) and Ainslie (1922) provided a brief description of several sod webworms. Artificial diets have been developed by Ward and Pass (1969) and Dupnik and Kamm (1970).

Damage Damage often occurs when larvae feed on the leaves of grasses, but feeding can also occur at the soil line or even on the roots. Under high-density conditions or if there is a shortage of leaf material, the plant stems and growing point

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of grasses may be eaten and the plants killed. Larvae commonly chew pits into the side of underground stems or leave the foliage ragged. Plant mortality is most common during periods of drought, and consumption of leaf material by webworms in the pasture environment often goes unnoticed during periods of adequate rainfall.

Management Sampling. Moths are highly attracted to light and can be captured in blacklight traps. Larval populations are assessed by careful examination of the soil surface. Insecticides. Insecticides are needed only when crops immediately follow sod or pasture infested with webworms. Webworms normally redistribute themselves within a year, dispersing from crop plants to grass-dominated areas. Liquid and granular insecticides can be applied at, or shortly after, planting to protect the seedlings. Cultural Practices. Rotation from sod or pasture to crops, especially corn, is risky if webworms have been abundant. Disking and tilling can destroy overwintering larvae, but intense soil disturbance is necessary. Both autumn and spring tillage are suggested for effective suppression.

Southern Cornstalk Borer

Diatraea crambidoides (Grote) (Lepidoptera: Crambidae)

Natural History Distribution. Southern cornstalk borer is, as its common name suggests, found predominantly in the southeastern states. It may be found as far north as Maryland and southern Ohio, and as far west as Kansas and New Mexico. However, these are the geographic extremes, and as a pest, its range is mostly limited to the southeastern states from Alabama and Florida to Virginia. It has been frequently confused with related species, particularly sugarcane borer, Diatraea saccharalis (Fabricius), so some records of occurrence are suspect. Southern cornstalk borer apparently is native to the southeastern United States, but also occurs in Mexico and South America. Host Plants. This insect is known principally from corn but occasionally damages sorghum and sugar cane. It may also be found in some of the wild grasses with thick stems such as Johnsongrass, Sorghum halepense; Paspalum spp.; panic grass, Panicum spp., and eastern gama grass, Tripsacum dactyloides. Natural Enemies. Mortality factors of southern cornstalk borer are not well documented. However, an undetermined fungus seems to be quite important in affecting the survival of overwintering larvae. Trichogramma sp. (Hymenoptera: Trichogrammatidae) egg parasitoids, as well as wasp parasitoids of the larval and pupal stages, have been noted on occasion, particularly Syntomosphyrum clisiocampae

(Ashmead) (Hymenoptera: Eulophidae), Macrocentris cingulum Reinhard (Hymenoptera: Braconidae), and Temelucha ferrunginea (Davis) (Hymenoptera: Ichneumonidae). The parasitic flies Lixophaga diatraeae (Townsend) and L. sphenophori (Villeneuve) (both Diptera: Tachinidae) have also been reared from this stalk borer. Probably several general predators consume larvae, and the goldenrod soldier beetle Chauliognathus pennsylvanicus De Geer (Coleoptera: Cantharidae) is among those insects known to attack larvae and pupae within the tunnels of the corn stalk. The fungus Beauveria bassiana has been isolated from larvae (Inglis et al., 2000). Life Cycle and Description. There are two generations per year, with the larval stage overwintering in the base of the corn stalk. In Virginia and North Carolina, pupation of overwintering larvae commences in late April and May. Adults and eggs of the first generation are found in May and early June, larvae from May to July, and pupae in July. Second-generation adults and eggs occur in July and early August, followed by the larval stage which persists until the following spring. Egg. The egg is flattened and oval. It measures about 1.6 mm long and 1.0 mm wide. They are deposited singly or in an overlapping, shingle-like fashion in small clusters. Initially, there may be as many as 20 eggs in a single cluster, but over time the female deposits smaller and smaller egg masses until she deposits only single eggs. Deposition usually occurs on the upper surface of leaf blades. The eggs are whitish when first deposited, but gradually assume a yellow or orange-yellow color. Duration of the egg stage averages about 9 days in the spring and 8 days during the summer (range about 7–13 days). Larva. Larvae increase in size from about 1.5 mm long to about 25 mm and display 5–6 instars during development. Newly hatched larvae are brownish, with a black head. Each body segment is darker toward the posterior, producing a transverse banded pattern. The appearance of the mature larvae differs slightly between seasons. Summer larvae bear a yellowish brown to a brown head capsule, with each body segment bearing several large dark spots dorsally and laterally on a whitish background. Overwintering larvae have a thicker, more robust appearance with the spots markedly lighter, barely darker than the whitish body color. Larvae of southern cornstalk borer are difficult to differentiate from other Diatraea spp. A key to the caterpillars boring in corn stalks is found in Appendix A. Mean development time of the first generation, or summer brood is about 30  days (range 24–38  days). Second brood larvae, of course, persist for about 9 months. Larvae tunnel freely within the stalk, with spring generation larvae usually feeding upward from the point of entrance. Overwintering larvae, however, eventually move down to

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the taproot to pass the winter in a more sheltered location. As they near the end of the larval stage the larvae eat to the outer edge of the stalk but leave a thin covering of the outermost layer of the stem in place or a few strands of silk across the exit hole. This presumably provides the mature larva and pupa protection from predators, but still allows the adult to escape. Second-generation larvae apparently do not feed in the spring but clear out the tunnel near the exit hole in preparation for pupation. Pupa. Pupation takes place within the tunnel produced by the larva. The pupa is mahogany brown and measures about 13–17 mm long and 3.5 mm wide. Duration of the pupal stage averages about 20 days in the spring and 12 days during the summer. Adult. The moths are yellowish brown except for the hind wings, which are white. Also, there is a small black spot near the center of the forewing, and the wing veins are darker brown. The tips of the front wings of the male moth are sharply angled rather than broadly rounded, approaching 90°, whereas the females are more rounded. The moths measure 25–35 mm in wingspan. As is characteristic of many pyralids, the palpi are enlarged and project forward, imparting the appearance of a pointed head. Oviposition usually commences 2 days after emergence and extends for 3–4 days. The moths are nocturnal and are not strong fliers. The females, in particular, fly only short distances. Females usually deposit about 200 eggs during their life span of 4–6 days, with nearly all eggs produced during the first two nights of oviposition. A good summary of southern cornstalk borer biology was given by Leiby (1920). The reports by Phillips et  al. (1921) and Cartwright (1934) are basically abbreviated versions of the same information. A key to the Diatraea larvae can be found in Peterson (1948), and Stehr (1987), and to the adults in Dyar and Heinrich (1927) and Solis and Mertz (2016). A key to stalk borers associated with corn in southern states is presented by Dekle (1976); this publication also includes pictures of the adults. A key to common stalk boring caterpillars also is found in Appendix A.

Damage The corn plant may be attacked at various stages of growth by southern cornstalk borer larvae, resulting in different types of injury. Early damage occurs when larvae feed on the unfolded leaves at the tip of the plant. If fewer than four larvae are present the result may be only ragged foliage bearing irregular holes. If several larvae are present, however, the bud or growing point of the plant may be killed, stunting the plant and preventing the production of a functional tassel. Larvae may also burrow through the veins of leaves and into the stalk. Tunneling larvae are found most frequently at the base of the stalk near the soil line and brace

roots. Stalks may support up to 15 larvae, but usually no more than 2–3 attain the pupal stage. Plants that have been tunneled by larvae are susceptible to wind damage.

Management Insecticides. Insecticides are not usually required for management of southern cornstalk borer. If necessary, insecticides can be applied in granular form to the whorl, where many larvae contact the insecticide. Insecticides can also be applied to the foliage, though because older larvae burrow within the plant, it is difficult to achieve high levels of suppression unless the insecticide is applied when the larvae are young. Systemic insecticides, particularly granular formulations applied to the soil, are also sometimes recommended. Cultural Practices. Several cultural practices can alleviate the damage by southern cornstalk borer. Destruction of stubble or lifting stubble from the soil can eliminate overwintering larvae. Burying stubble deeply can also prevent moths from escaping in the spring. Modified planting dates are sometimes recommended because late-planted corn can escape infestation. In Virginia and North Carolina, corn planted in June generally escapes damage by firstgeneration insects but is susceptible to infestation by the second generation. In general, the earlier the planting the more heavily it will be infested. Because moths, particularly females, do not fly far after emergence, crop rotation has considerable benefit. Transgenic corn varieties containing Cry toxins have come into use for suppression of Diatraea spp. worldwide.

Southwestern Corn Borer

Diatraea grandiosella Dyar (Lepidoptera: Crambidae)

Natural History Distribution. Southwestern corn borer is native to Mexico and appears to have been first found in the United States in New Mexico in 1891. By 1931, it had spread to the nearby states of Arizona, Texas, and Oklahoma. By 1970, the northern limits of its distribution were Kansas, Missouri, and Kentucky and the eastern limits were Tennessee and Alabama. Thus, it is found widely in the southern United States east of California. Host Plants. This species attacks only grasses, with corn as the principal host. Sorghum and sorghum hybrids, broomcorn, sugarcane, and pearl millet are other crops that sometimes serve as hosts. Among weeds, Johnsongrass, Sorghum halepense, may also serve as an alternate host, but development time is much longer and fecundity greatly reduced on this plant (Aslam and Whitworth, 1988). Natural Enemies. Several natural enemies are known, though their value is open to question. In Texas, egg or

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early instar mortality is critical to the survival of the spring population, whereas in the summer generation survival is most affected by the mortality of large larvae and diapausing larvae (Knutson and Gilstrap, 1989a, b, 1990). Among the egg parasitoids are Trichogramma spp. (Hymenoptera: Trichogrammatidae), and a larval parasitoid of Diatraea spp., Apanteles diatraeae Muesebeck (Hymenoptera: Braconidae) (Knutson and Gilstrap, 1989a, b). Parasitism by Trichogramma spp. usually occurs late in the oviposition period at low levels, but sometimes 20% or more of the eggs are parasitized (Moulton et  al., 1992). General predators such as insidious flower bug, Orius insidiosus (Say) (Hemiptera: Anthocoridae), and lady beetles such as Hippodamia spp. (Coleoptera: Coccinellidae) are common in cornfields but have little impact because they often seem to be out of synchrony with susceptible stages of southwestern corn borer. Pathogens such as the fungus Beauveria bassiana, and sometimes Bacillus sp. bacteria, also affect larvae, particularly the overwintering population. Knutson and Gilstrap (1989a) reported that up to 45% of large larvae were infected with Beauveria. Rolston (1955) indicated that unspecified nematodes were sometimes observed, and Knutson and Gilstrap (1990) reported minor incidence of Heterorhabditis sp. (Nematoda: Heterorhabditidae). Mortality also occurs during the winter months when the larval overwintering cell is penetrated by stalk rot fungi and termites, apparently because such penetration allows seepage of water into the cell. Thus, heavy rainfall and wet soils are detrimental to overwintering survival, and southwestern corn borer typically is more destructive in areas with sandy soil. Considerable overwintering mortality also results from bird predation, often 50%–80% of the total mortality. The yellowshafted flicker, Colaptes auratus (L.), is the most important avian predator (Black Jr. et al., 1970). Rodents also sometimes consume overwintering larvae, but this occurs erratically. A more recent study documented the presence of the entomopathogenic bacteria Enterococcus faecalis, Pseudomonas aeruginosa, Serratia marcescens, and Bacillus spp. Also, Nosema spp. fungi were observed (Inglis et al., 2000). Life Cycle and Description. In most locations, southwestern corn borer exhibits two to three generations per year. In Mississippi, moths are present in late April-early May, late June-early July, and in August. In northern Arkansas, corresponding moth flights appear in June, July, and August-September. In Texas, moths are found in late May-June, mid-July-early August, and sometimes in late August-early September. A generation normally requires 40–50  days. In most areas, larvae of the first or spring generation do not display diapause, but larvae from the subsequent generation(s) may enter diapause depending on photoperiod and temperature present when they are

d­ eveloping. A photoperiod of less than about 15 h often induces diapause. Larvae developing after early August usually enter diapause in Missouri. Egg. The eggs of southwestern corn borer are oval, measuring about 0.8 mm wide and 1.3 mm long. They are flattened, and deposited in an overlapping manner, resembling scales of a fish. The eggs are creamy white but bear three parallel, transverse, orange-red lines. They are deposited principally on the upper surface of foliage, but also on the lower surface, and occasionally on the stem (Poston et al., 1979). The number of eggs per mass varies from one to several, but averages 3–5 per cluster. Initially, egg masses tend to consist of several eggs, but the number diminishes as the females age, so by the fourth day of oviposition most females deposit but single egg. The incubation period of the eggs is 4–7 days.

FIG. 10.31  Southwestern corn borer larva. (Photo by J. Capinera.)

Larva. Most larvae pass through six instars, but 5–8 instars have been observed. Mean head capsule widths for instars 1–6 are 0.34, 0.51, 0.89, 1.34, 1.76, and 2.24 mm, respectively (Jacob and Chippendale, 1971). Mean duration of instars fed corn was determined to be 3.1, 2.9, 3.0, 4.2, 4.8, and 7.0 days for instars 1–6, respectively, when reared under variable insectary conditions. Larvae fed corn required about 22  days for development, whereas those fed sorghum, millet, and Johnsongrass required about 29, 49, and 45 days, respectively. Larval survival rates are much higher on young corn than old plant material due to the absence of succulent tissue on mature corn. Larvae in the first two instars are whitish but have a reddish prothorax and dorsal reddish stripe along the abdomen. The third and succeeding instars of nondiapausing larvae are distinctly marked with black spots on a yellowish-white background. Larvae that overwinter do not acquire the black spots, rather appearing mostly yellowish white, or marked only with faint brownish spots. Larvae are difficult to distinguish from other Diatraea stalk borers. Larvae generally feed externally on leaf or husk tissue during the first three instars, then bore

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into the stalk for the remainder of their larval development period. Once inside the stalk, larvae tunnel for about 7–30 cm. Following tunneling, nondiapausing larvae prepare an exit hole for the adult and pupate within the tunnel. Larvae about to enter diapause feed downward, or leave the feeding site and crawl downward externally, moving to the base of the stalk and entering the taproot where they construct a crude cell in preparation for overwintering. As part of the prediapause feeding behavior, larvae may girdle the stalk a few centimeters above the soil line. Only southwestern corn borer, among the several Diatraea spp. affecting corn, girdles the stalk. Girdling apparently is an acquired trait associated with dispersal into colder climates. Girdling is infrequent in Mexico, but common in the United States, where increased overwintering survival is associated with this behavior. Larvae tend to be cannibalistic, especially as they prepare for diapause, and though several larvae may develop in a single stalk, usually, only one successfully overwinters. In the spring, larvae clean the escape tunnel in preparation for the emergence of the adult stage. They close off the tunnel exit with strands of silk, probably to deter entry by predators that might attack the pupal stage. Pupa. Pupation usually occurs in the overwintering cell in the taproot, but occasionally above-ground within the tunneled area of the stalk. The pupa is yellowish brown with diffuse dark bands, especially dorsally. The tip of the abdomen is broadly rounded and bears thick spines. The pupa measures 13–25 mm long and 3.5 mm wide. Duration of the pupal stage averages about 14.8  days (range 11–20 days).

FIG. 10.32  Adult of southwestern corn borer. (Photo by J. Capinera.)

Adult. The moths are buff or tan in color, with seven faint narrow lines on each forewing that terminate in a minute dark spot. The hind wings are white with buff-colored veins. The male moth measures 15–30 mm in wingspan, whereas the female measures 30–38 mm. The palpi are prominent, projecting forward from the head in a manner

common among pyralid moths. The adult stage is short lived, persisting for about 4–5 days and does not feed. Adult females are ready to oviposit within 24 h of emergence. They are active at night, particularly 1–2 h before and after midnight. Females produce and release a sex pheromone during the first 3  days after emergence. Females produce 300–400 eggs. The biology of southwestern corn borer was reviewed by Davis et  al. (1933), Rolston (1955), Henderson and Davis (1969), and Chippendale (1979). Phenology was also presented by Walton and Bieberdorf (1948) and Knutson et al. (1982). Developmental biology was given by Whitworth and Poston (1979), Knutson et  al. (1989), and Ng et  al. (1993). An artificial diet for borer culture was presented by Whittle and Burton (1980). A bibliography was published by Morrison et al. (1977). A key to the Diatraea larvae can be found in Peterson (1948) and Stehr (1987), and to the moths in Dyar and Heinrich (1927) and Solis and Mertz (2016). Southwestern corn borer was included in the larval key by Capinera (1986) and the moth key by Capinera and Schaefer (1983). A key to stalk borers associated with corn in southern states was presented by Dekle (1976); this publication also includes pictures of the adults. A guide to the common stalk boring caterpillars is found in Appendix A.

Damage All stages of the plant may be injured by feeding of southwestern corn borer larvae. Early instars of the first generation feed on leaf tissue, especially new tissue within the whorl of young corn plants. This can result in the destruction of the terminal bud (a condition called “dead heart”), loss of apical dominance, and development of lateral buds. Such plants are stunted and bushy. Early instars of the second generation feed mostly on leaf sheaths, the husk, shank, kernels, and cobs of ears. In sweet corn it is difficult to detect the presence of the larvae in the ear until the husk is removed. Late instars of all generations bore within the stalk. Stalk damage may result in stunting if it occurs early in the growth of the plant; in more mature plants tunneling may disrupt translocation of nutrients to the ears, causing decrease in kernel size. Larvae about to enter diapause also girdle the stalk internally just above soil level. Girdling increases the likelihood that plants break. Stalk breakage is not often a problem in sweet corn due to early harvesting but is a severe threat to grain corn. Damage potential to grain corn was presented by Whitworth et al. (1984).

Management Sampling. Moths can be taken at blacklight traps, but they are not strongly attracted to light. Several types of traps baited with sex pheromone can be used to m ­ onitor

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populations (Knutson et al., 1987), but they differ in their ability to capture moths. The nonsticky Heliothis trap (Davis et  al., 1986) and bucket trap (Goodenough et  al., 1989) are most effective. The distribution of eggs and small larvae are highly aggregated, but older stages are progressively less clumped. Sequential sampling protocols have been developed (Poston et  al., 1983; Overholt et al., 1990). Insecticides. Several protocols are available for ­insecticide-based suppression of larvae (Daniels, 1978; Buschman et  al., 1985). Liquid formulations of insecticides are applied to the foliage to kill young larvae before they burrow into the stalk. Fairly precise timing or multiple applications may be necessary to produce good larval suppression, and application of insecticide in overhead irrigation systems is effective. Granules may be applied into the whorls because larvae tend to aggregate on this succulent tissue. Systemic insecticides applied to the soil at planting are effective for first-generation borers, and may also be applied to foliage for second-generation infestations. Cultural Practices. Several cultural practices are used to minimize the effects of southwestern corn borer on corn production. Early planting is often recommended, because damage to the growing point of the corn plant is minimized or prevented. Cultivation, especially if done early in the winter, prevents adults from emerging successfully (Archer et  al., 1983). Lifting stubble from the soil exposes larvae to more severe overwintering conditions, and also reduces adult emergence in the spring. Corn planted at high densities is more likely to suffer girdling, than corn at low densities, despite a tendency for the equal incidence of infestation; the basis for this disparity is unknown (Zepp and Keaster, 1977). Host-Plant Resistance. Corn cultivars possessing considerable resistance to corn borer feeding have been located and incorporated into commercial varieties, with mixed results (Davis et  al., 1991; Ng et  al., 1990; Thome et  al., 1992). Resistance is attributable to both limited damage by larvae and nonpreference by adults (Ng et  al., 1990). Transgenic corn varieties containing Cry toxins have come into use for suppression of Diatraea spp. worldwide. Biological Control. The wasp Pediobius furvus (Gahan) (Hymenoptera: Eulophidae) was imported from Africa to attack nocutid and pyralid borers, and evaluated against southwestern corn borer in Texas. Low levels of borer parasitism were obtained under field conditions, and the wasp failed to overwinter, thus limiting its usefulness to augmentative releases (Overholt and Smith Jr., 1989). Southwestern corn borer is also susceptible to infection by the bacterium Bacillus thuringiensis and the alfalfa looper baculovirus (Davis and Sikorowski, 1978; Nolting and Poston, 1982), but these materials have not come into general use for southwestern corn borer.

Sugarcane Borer

Diatraea saccharalis (Fabricius) (Lepidoptera: Crambidae)

Natural History Distribution. This species is native to the western hemisphere, but not to the United States. It apparently was introduced into Louisiana about 1855 and has since spread to the other Gulf Coast states. It inhabits only the warmer parts of these states, however. Sugarcane borer also occurs throughout the Caribbean, Central America, and the warmer parts of South America south to northern Argentina. Host Plants. Sugarcane borer attacks plants in the family Gramineae. Though principally a pest of sugarcane, this insect also feed on other crops such as corn, rice, sorghum, and sudangrass. Many wild or weed grasses are suitable hosts, including Johnsongrass, Sorghum halepense; Paspalum sp.; Panicum spp.; Holcus sp.; and Adropogon sp. Natural Enemies. The importance of natural enemies in corn-cropping systems is not known, because most studies involve only sugarcane. Ants are reported to be important predators of sugarcane borer in sugarcane fields, capable of reducing damage by over 90% (Bessin and Reagan, 1993). Although much of the attention has been focused on red imported fire ant, Solenopsis invicta Buren, other species such as Pheidole dentata Mayr and P. floridana Emery (all Hymenoptera: Formicidae) are also important (Adams et al., 1981). Effective parasitoids are not established in the United States. Egg parasitoids, Trichogramma sp. (Hymenoptera: Trichogrammatidae), are possibly the most important naturally occurring parasitic insects. Although they are not very abundant early in the season, by autumn they may inflict almost complete destruction of borer eggs. The most important imported parasitoid is Agathis stigmaterus (Cresson) (Hymenoptera: Braconidae), which was reported by King et al. (1981) to affect, on average, less than 12% of borers. Lixophaga diatraeae (Diptera: Tachinidae) has the potential to cause high levels of parasitism but does not persist well (see biological control, below). A wasp introduced from India, Cotesia flavipes Cameron (Hymenoptera: Braconidae), is an important late-season parasitoid late in the summer within Florida. Other parasitoids include Orgilus elasmopalpi Muesebeck, Apanteles diatraeae Musebeck, Apanteles impunctatus Musebeck Apanteles flavipes (Cameron) (all Hymenoptera: Braconidae), Euplectrus plathypenae Howard, and Syntomosphyrum clisiocampe (Ashmead) (both Hymenoptera: Eulophidae). The comparative assessment of natural enemies in sugarcane and sorghum conducted by Fuller and Reagan (1988) probably offers some insight into the role of natural enemies in corn, because cultural practices in sorghum and corn are similar. Predator densities were higher in sugarcane owing to the greater abundance of red imported fire ant. However, Orius spp.

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pirate bugs (Hemiptera: Anthocoridae), lacewings (Neuroptera: Chrysopidae), tiger beetles (Coleoptera: Cicindelidae), spiders, and foliage-dwelling ground beetle larvae (Coleoptera: Carabidae) were more abundant in sorghum fields. Suppression of predators with soil-applied insecticide affected predation in both agroecosystems, with borer populations 40%–60% higher where predator abundance was reduced. Weather. An inverse relationship between rainfall and borer abundance has been reported from both Louisiana and Puerto Rico. Heavy rainfall, and particularly winter rainfall resulting in flooding, depresses borer survival (Holloway et al., 1928). This is thought to result from prolonged emersion of stalks containing overwintering larvae in floodwater. Also, young larvae living in the whorl of corn or sugarcane are quite tolerant of short-term emersion, but heavy rainfall, while they are dispersing, could lead to death because they are washed from the plants. In addition to rainfall, cold winter temperature is reported to depress larval survival rates in Louisiana. Life Cycle and Description. Overwintering occurs in the larval stage, with pupation in the spring. In Louisiana and Texas, adults become active by April or May, and the borer population continues to cycle until autumn. Development time is highly variable, so the generations overlap considerably, obscuring population trends. There is potential for four to five generations to occur annually, but moths are abundant only in spring and autumn (Fuchs and Harding, 1979), so perhaps there are fewer generations. During the summer a complete generation may require only 25 days, whereas during the winter over 200 days are needed. Egg. The eggs are flattened and oval, measuring about 1.16 mm long and 0.75 mm wide. They are deposited in clusters and overlap like the scales of a fish. An egg cluster may contain from 2 to 50 eggs, with eggs deposited on both the upper and lower surface of leaves. The eggs are white initially, but turn orange with age and then acquire a blackish hue just before hatching. Duration of the egg stage is 4–6  days. Mean fecundity is about 700 eggs when borers are reared on corn and sugarcane, but only about 425 eggs when fed Johnsongrass (Bessin and Reagan, 1990).

FIG. 10.33  Sugarcane borer larva. (Photo by J. Capinera.)

Larva. The eggs within a cluster hatch about the same time, or at least within a few hours of one another. Larvae tend to congregate in the whorl of corn plants and begin feeding almost immediately. They may feed through the leaf tissue or tunnel through the midrib. After the first or second molt they burrow into the stalk. The larvae display both summer and winter forms. The larvae are whitish with a brown head, but the summer form also bears large brown spots on each body segment whereas the winter form lacks spots. A stout hair originates in each of the spots, or for the winter form, from the location where the spot might appear. Larvae during the winter are rarely found in corn; sugarcane and stalks of large grasses are more suitable and preferred. Instar number is quite ­variable. There are reports of about 3–10 instars, but 5–6 is normal. Holloway et al. (1928) reported instar duration of about 3–6, 4–8, 6–9, 4–6, and 4–9 days for instars 1–5, respectively, for larvae fed sugarcane. When reared on artificial diets, most larvae tend to display six instars. Roe et al. (1982) reported mean head capsule widths of about 0.29, 0.40, 0.62, 0.93, and 1.32 mm for instars 1–5 in larvae that had six instars; head capsule measurements were not reported for the final instar but probably were about 1.75 mm. Larval development time usually requires 25–30 days during warm weather and 30–35 days during cool weather except, of course, during the winter when development is arrested. Larvae attain a length of about 2–4, 6–9, 10–15, 15–20, and 20–30 mm during instars 1–5, respectively. Larvae of sugarcane borer are easily confused with southern cornstalk borer, and definitive separation involves microscopic examination of the mouthparts. Sugarcane borer, however, is much less likely than southern cornstalk borer, Diatraea crambidoides (Grote), to be found infesting corn. Pupa. Pupation occurs within the plant, in a tunnel created by the larva. The larva cleans and expands the tunnel before pupation, leaving only a thin layer of plant tissue for the moth to break through at emergence. The pupa is elongate and slender, and yellowish brown to mahogany brown. It measures 16–20 mm long and bears prominent pointed tubercles on the distal segments. Duration of the pupal stage is usually 8–9 days, but under cool conditions may extend for up to 22 days. Adult. The adult is a yellowish or yellowish-brown moth with a wingspan that measures 18–28 mm in males and 27– 39 mm in females. The forewing also bears numerous narrow brown lines extending to the length of the wing. The hind wing of females is white, but in males, it is darker. The adults are nocturnal and remain hidden during the daylight hours. Oviposition commences at dusk and continues throughout the evening. Females may deposit eggs for up to 4 days, but often less. Duration of the adult stage is 3–8 days. The biology of sugarcane borer was described by Holloway et al. (1928) and a bibliography was authored by Roe (1981). Several wheat germ-based diets are suitable for rearing (Roe et al., 1982). A key to the Diatraea larvae can be found in Peterson (1948), and Stehr (1987), and to

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the adults in Dyar and Heinrich (1927) and Solis and Mertz (2016). A key to stalk borers associated with corn in southern states was presented by Dekle (1976); this publication also includes pictures of the adults. A key to common stalk boring caterpillars is also included in Appendix A.

Damage Although generally regarded as a potentially serious pest of sugarcane, other crops are rarely at risk. Sugarcane borer is a minor pest of sweet corn even in Florida, where the weather favors its survival and sugarcane is abundant (Kelsheimer et  al., 1950). Damage by sugarcane borer to grain corn was described by Flynn and Reagan (1984) and Flynn et al. (1984). Larvae injure corn in two ways. Early in the season, they attack the whorl, feeding on the young developing tissue. If such damage is light, the result may be only series of holes across the leaf blade. If such damage is extensive, however, the growing point of the plant may be killed and plant growth stunted. This condition is called “dead heart.” Later in the season, the larvae descend to the stalk and burrow into it. Large larvae tunnel through the stalk, causing the plant to be prone to breakage. On occasion, especially during the second generation, larvae may burrow into corn ears (Rodriguez-del-Bosque et al., 1990).

Management Sampling. Sampling protocols have not been devised for sweet corn because this is a relatively minor pest. Monitoring of adult populations, egg density, and foliar feeding by young larvae are advisable if sweet corn is cultured in the vicinity of sugarcane. Larval distribution in sugarcane was described by Hall (1986). Sex pheromones have been identified for sugarcane borer, but there is variability among geographical “strains” (Cortés et al., 2010). Joyce et al. (2014) suggested that at least two separate introductions to the United States have occurred and that at least one cryptic species exists, thus perhaps explaining the variability in response to sex pheromones. Insecticides. Insecticides can be applied to the foliage of sugarcane, providing significant yield increases even in the presence of predation and resistant varieties (Bessin et al., 1990). Insecticides should be applied while the larvae are young before they burrow into the stalk. However, some control is possible even later, possibly because larvae leave their tunnel during the process of pushing out excrement. Cultural Practices. Sugarcane is the principal host of sugarcane borer, and proximity of corn to sugarcane is an important determinant of borer abundance in corn. Moths deposit more eggs on sugarcane than on corn when these hosts are in close proximity, and avoid pubescent cultivars (Sosa, 1990). It is advisable to destroy cane trash in the winter as it reduces overwintering by larvae, but the practice of burning does not always kill borers deep within the stalks. Borers also overwinter within corn stalks, but usually, only

late-planted corn is suitable. Some sugarcane cultivars display considerable resistance to sugarcane borer (Bessin et al., 1990; Bessin and Reagan, 1993), which presumably can reduce the overall abundance of borers and infestation potential in corn. Grain corn varieties with resistance to sugarcane borers also have been identified (Maredia and Mihm, 1991). Transgenic corn varieties containing Cry toxins have come into use for suppression of Diatraea spp. worldwide. Biological Control. The Caribbean region and tropical areas of South America have been surveyed extensively for natural enemies. Many species were introduced into the United States, but few were established (Clausen, 1978). Agathis stigmatera (Cresson) (Hymenoptera: Braconidae) was successfully imported from Argentina and Peru, and though it is well-established in both Florida and Louisiana, its effect on sugarcane borer is minimal. The fly Lixophaga diatraeae (Townsend) (Diptera: Tachinidae) was imported and released repeatedly but tends to disappear or dissipate after a few years. In some countries, augmentative releases are used to attain high levels of parasitism in sugarcane borer, and this has been attempted in Louisiana (King et al., 1981). Some authors have claimed success with augmentative releases of Trichogramma spp. (Hymenoptera: Trichogrammatidae), but this has proven difficult to implement in the United States (Long and Hensley, 1972). Entomopathogens have been shown to be effective for sugarcane borer suppression, though they are not used routinely as insecticides. For example, Alves et al. (1984) demonstrated that Metarhizium anisopliae could affect larvae as suppressive agents. However, the application of Bacillus thuringiensis is more common (de Medeiros Gitahy et al., 2007), particularly via transgenic host plants, as preventative agents) (Tan et al., 2011).

Sweetpotato Leaf Folder

Lygropia tripunctata (Fabricius) (Lepidoptera: Crambidae)

Natural History Distribution. This species is known from Hawaii, Texas, Louisiana, and Puerto Rico. It is also recorded from the Caribbean and Central American regions. Host Plants. This plant is found to feed only on the genus Ipomoea, both wild and cultivated varieties. Natural Enemies. Exorista pyste Walker (Diptera: Tachinidae) are known to attack the larvae, as are adults of Podisus maculventris Say (Hemiptera: Pentatomidae). Life Cycle and Description. This species apparently undergoes at least four and sometimes five generations per season. Each generation requires at least 25 days for ­completion, consisting of 4 days in the egg stage, 13 days in the larval stage, 2 days in the prepupal stage, and 6 days in the pupal stage. The eggs require about 3 days for development before they are laid. The biology is given by Jones (1917a, b).

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Egg. The egg is elliptical in shape. Initially it is colorless, though later in the development the embryo is evident. The eggs measure a mean length of 1.00 mm whereas the width is 0.79 mm. The eggs are deposited singly and in small groups of up to 5 per egg clutch. The eggs are typically found on the leaf tissue. Duration of the egg stage is about 4 days.

Adult. The upper color of the moth is light yellow, with the wings iridescent. A dark-grayish-brown band occurs along the periphery of the wings, especially along the margin of the forewings. Two brown spots are located along the costal margin of each forewing, and a small black spot is found on the hind wings. A wavy dark line is found at about the midpoint of the fore- and hindwings. The measurements of the male and female are about 27 mm (range of 25–29 mm) and about 25–27 mm, respectively.

Damage Damage results from feeding by the larvae of sweetpotato leaf folder on the leaf tissue.

Management The larvae can become quite numerous, and liquid insecticide is often applied to disrupt their development.

Sweetpotato Vine Borer FIG. 10.34  Larva of sweetpotato leaf folder. (Photo by L. Buss.)

Larva. Hatching larvae are about 1.5 mm long. At this stage, larvae are nearly colorless. After ingesting chlorophyll, they acquire a green color, and eventually may be brownish or bluish green. The head is pale yellow. Eventually, the larvae attain a length of about 27 mm. The larva typically constructs a shelter composed of the leaf material that shortens as the silk dries. Initially, the young larva creates small holes, but as the larva increases in size, the entire leaf may disappear. The larval stage persists for 13–14 days. Duration of the larval instars was about 3, 2, 2, 2, 2, and 6 days each for instars 1–6, respectively. Pupa. The pupa is dark brown in color. They measure about 15 mm in length, and 4 mm in width. Pupation requires about 6–9 days.

Omphisa anastomasalis (Guenée) (Lepidoptera: Crambidae)

Natural History Distribution. Sweetpotato vine borer is widespread in Asia where it is destructive in such countries as China, India, Indonesia, Japan, Philippines, and Vietnam. In the United States, its distribution is limited to Hawaii, where it was first observed in 1900. Host Plants. This species is associated with plants in the family Convolvulaceae. It is destructive only to sweet potato, but other Ipomoea spp. are common hosts. In Hawaii, it is also reported from Stictocardia campanulata. Natural Enemies. Larval parasitoids known from Hawaii include Chelonus blackburni Cameron (Hymenoptera: Braconidae), Enytus chilonis Cushman and Pristomeris hawaiiensis Perkins (both Hymenoptera: Ichneumonidae). Other parasitoids are known from Asia. Life Cycle and Description. Phenology of sweetpotato vine borer is not documented. Considering the importance of the insect and crop, relatively little work has been done on this species. Egg. The eggs of sweetpotato vine borer are elliptical with a flat base, measuring about 0.6 mm long, 0.5 mm wide, and 0.35 mm in height. They are greenish and laid singly or in small clusters of 2–3 in crevices on leaves, petioles, and stems. The incubation period is 5–7 days.

FIG. 10.35  Adult of sweetpotato leaf folder. (Photo by L. Buss.)

FIG. 10.36  Sweetpotato vine borer. (Drawing by USDA.)

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Larva. After hatching, larvae bore into the vine and move toward the base of the plant as they feed. Larvae are only about 1 mm long at hatching and are whitish with a black head and prothoracic shield. Mature larvae attain a length of about 30 mm and may be yellowish white or light purple. Large larvae are marked with brownish tubercles, which appear as spots over most of the body. The intersegmental membranes tend to be yellowish brown, especially in the anterior region of the body. The head and prothorax are brownish and the body bears scattered stiff hairs. There are six instars. The duration of the larval stage is usually 30–35 days, but values of 21–92 days have been reported.

Damage Larvae mine the vines of sweet potato, disrupting the flow of water and photosynthates. Infested vines show weak growth, poor foliage development, and poor tuber development. Larvae also may bore into the upper portions of tubers. Yield reductions are directly related to infestation levels, and yield loss of 30% or more are common in Asia. In Hawaii, heavy infestations have been reported to kill plants (Talekar and Cheng, 1987; Talekar and Pollard, 1991). When infestation levels are low the feeding damage is much less pronounced. Sometimes, the only outward ­evidence of larval feeding is the accumulation of fecal material near the opening of the larval tunnel, which usually is at the crown of the plant.

Management FIG. 10.37  Pupa of sweetpotato vine borer. (Drawing by USDA.)

Pupa. Pupation normally occurs within the base of the vine, but occasionally larvae pupate in the tubers if they are close to the soil surface. The larva spins a thin web-like cocoon and cuts an exit hole for the moth before pupation. The pupa is light to medium brown and measures about 16 mm long and 3 mm wide. Duration of the pupal stage is 14–18 days.

Insecticides. Insecticides can be applied for larval suppression, but the mining behavior of this insect requires that insecticides be in place at hatching, before larvae burrow into the tissue, or be systemic and translocated to the tissues where larvae feed. In areas of the world where sweetpotato vine borer is common, the use of insecticide often results in very sizable yield increases. Farmers are often discouraged from using insecticides, however, because of the low value of the crop. Host-Plant Resistance. Considerable effort has been directed to screening varieties for resistance to sweetpotato vine borer. Although some cultivars display the resistance (Talekar and Cheng, 1987) the level of resistance is moderate or the yield potential of these selections is low, so additional work is needed by breeders to improve resistance and yield.

FAMILY EREBIDAE—WOOLLYBEAR CATERPILLARS, TIGER MOTHS, AND OTHERS FIG. 10.38  Adult of sweetpotato vine borer. (Drawing by USDA.)

Adult. The moth is white but is heavily marked with yellowish brown. The base of the front wings bears a large, irregular dark spot. The abdomen is brown dorsally. The distal area of the front wings and the entire hind wings are marked with irregular dark lines. The wingspan is 30–40 mm. Moths are nocturnal and females produce sex pheromone, which has been identified (Yan et al., 2014). Adults survive for about 10  days, during which females deposit about 300 eggs. Fullaway (1911) and Talekar and Pollard (1991) provided the biology of sweetpotato vine borer. Yoshiyasu (1975) described the adult stage. A review of sweetpotato pests is provided by Johnson and Gurr (2016).

Banded Woollybear

Pyrrharctia isabella (J.E. Smith) (Lepidoptera: Erebidae)

Natural History Distribution. A native species, banded woollybear is found throughout the United States and southern Canada. Banded woollybear is the best known of the woollybears, because in American folklore its color pattern is said to foretell the severity of forthcoming winter weather. Banded woollybear, though common, is more of a curiosity than a pest. Host Plants. Banded woollybear consumes a wide breadth of flora, though damage to economic plants is infrequent, and therefore poorly documented. It is recorded from beet, corn and pea, and probably nibbles on nearly

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any garden vegetable, but its normal hosts are weeds and wild native plants. The normal foods are asters, Aster sp.; dandelion, Taraxacum officinale; dock, Rumex spp.; goldenrod, Solidago spp.; plantain, Plantago spp.; sweetclover, Melilotus; and some grasses. In the only broad study of banded woollybear feeding, a no-choice test, over 90 plant species from over 50 plant families were consumed to some degree (Shapiro, 1968). Natural Enemies. Several natural enemies are known, but they have not been the subject of many studies, so their relative importance is to be determined. Fly parasitoids associated with banded woollybear include Euexorista futilis (Osten Sacken), Exorista mella (Walker), Hubneria estigmenensis (Sellers), Mystacella spp., Parachaeta fusca Townsend, Thelaria spp., and Winthemia datanae (Townsend) (all Diptera: Tachinidae). Among wasps known to attack this woollybear are Apanteles flavovariatus Muesebeck (Hymenoptera: Braconidae); Coccygomius pedalis (Cresson), Dusona crassicornis (Provancher), and Enicospilus glabratus (Say) (all Hymenoptera: Ichneumonidae). A cytoplasmic polyhedrosis virus has been isolated from banded woollybear larvae; infected larvae displayed significantly slower rates of development (Boucias and Nordin, 1978). Life Cycle and Description. The phenology of banded woollybear is not well known, but there appears to be at least two generations in the southern United States, and only a single generation in the northern states and southern Canada (Goettel and Philogene, 1978a). Thus, there are reports of a single generation in Ontario, Quebec, the New England states, and New York, but two generations in Illinois. Egg. Banded woollybear deposits spherical eggs in clusters of 50–100 eggs. Duration of the egg stage is about 6 days at 27°C. Larva. Larval development is rather variable in this species. When reared on artificial diets, the larvae display 7–10 instars, with development times of about 3, 3, 3.2, 3.9, 4.9, 6.9, 14, 21 (and if present) 21 days, respectively for instars 1–9 (Goettel and Philogene, 1978b). The same authors (Goettel and Philogene, 1979) also provided head capsule measurements for larvae with 7–10 instars. Head capsule widths for larvae with eight instars, a common number, are 0.4, 0.5, 0.8, 1.2, 1.7, 2.3, 2.9, and 3.7 mm, respectively. Instar number is influenced by photoperiod but not by temperature; exposure of larvae to greater than 14 h photophase results in an increased number of instars (Goettel and Philogene, 1978a). The banded woollybear is thickly clothed with stout bristles. The head and body are black, with black or reddishbrown spines covering the body. Typically the caterpillar’s bristles are brown at the middle of the body and black at

both the anterior and posterior ends of the caterpillar. Young larvae tend to be about two-thirds black, with the amount of black dissipating as larvae mature. Sometimes only the head end remains black, and in California, caterpillars are sometimes uniformly brown. The larva overwinters and is often observed in the autumn as it disperses in search of a suitable overwintering shelter. The larvae typically overwinter beneath plant debris on the soil surface, where they are somewhat insulated from the extreme cold. However, they are quite tolerant of freezing, and their glycerol content increases with exposure to freezing, further increasing freeze tolerance (Layne et al., 1999). If cold weather commences early in the year, the larvae are relatively immature, and thus possesses a disproportionately large amount of black coloration as compared to years with long summers, when the larvae are more mature, and thus more brown. Thus, there is some meteorological basis to the American belief that banded woollybear foretells the weather, but the color registers the past weather, not the future. Banded woollybear larvae tend to have mostly one type of hair, of equal length, and have a rather short-cropped look as compared to some other long-haired species such as yellow woollybear, Spilosoma virginica (Fabricius). There may be a few long hairs at both the anterior and posterior ends of the caterpillar, but this does not detract from the overall impression of uniform hair length. The mature larva attains a length of about 30 mm and has a tendency to roll into a ball if disturbed. Pupa. Banded woollybear overwinters as a larva in leaf debris, under loose bark and similar shelter, and in the spring it feeds briefly before pupating. The pupal case is constructed principally from hairs from the caterpillars body, spun loosely together with silk. The cocoon is dark because it consists mainly of black and brownish body hairs. The pupa is light to dark brown. Pupation usually requires 14–21 days.

FIG. 10.39  Adult of banded woollybear. (Photo by L. Buss.)

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Adult. The moth is yellowish orange to light brown, with one to several dusky spots on the front wings and two on each hind wing, and has a wingspan of about 45 mm. In males, the hind wing tends to be paler. The abdomen is marked with three rows of black spots. Banded woollybear courtship is unusual in that it involves acoustical signals produced by females in response to male production of pheromone (Krasnoff et al., 1987; Krasnoff and Yager, 1988). The biology of this species is poorly documented. Banded woollybear biology was discussed by Saunders (1873) and a method of culture was given by Goettel and Philogene (1978b).

Damage Larvae are defoliators, but they feed principally on weeds and other noncrop plants. Young larvae are gregarious, feeding together on the underside of foliage and skeletonizing the plant tissue. Larger larvae disperse and feed sporadically, creating irregular holes in foliage.

Management It is highly unusual to experience banded woollybear in enough abundance to warrant concern. However, they are easily killed with foliar insecticides if it becomes necessary.

Green Cloverworm

Hypena scabra (Fabricius) (Lepidoptera: Erebidae)

Natural History Distribution. This native species is found widely in eastern North America. It is recorded from all states east of the Great Plains, and there are occasional records from the Rocky Mountain region. As a pest, however, it is best known from the soybean-growing areas of the midwest and southeast where it has an abundance of the suitable host material. In Canada, green cloverworm is known from southern Ontario, but it rarely causes serious damage. Host Plants. Larvae of green cloverworm develop successfully only on plants in the family Leguminosae. They have been observed to feed on weeds and crops from other plant families, but this occurs only after legumes have been consumed. Vegetable crops eaten include bean, cowpea, edamame, faba bean, lima bean, and pea. Field crops suitable for development include alfalfa, alsike clover, crimson clover, red clover, white clover, lespedeza, birdsfoot trefoil, velvet bean, and soybean. Pedigo et al. (1973) indicated that the most common food plants are soybean, alfalfa, clovers, field bean, lima bean, and pea, in that order. Adults feed on the nectar from blossoms. Because of the preference for soybean, most of this insect’s biology and management recommendations have

been derived from soybean-based research, but in large measure, the findings should be applicable to related crops. Natural Enemies. Many natural enemies are known, with their significance varying according to cloverworm population density. In Iowa, research has shown that low (endemic) densities, parasitoids and to a lesser extent predators are relatively important. During outbreak (epidemic) densities, resulting from invasion by many migrating moths early in the year, the entomopathogenic fungus Metarhizium (Nomuraea) rileyi becomes a key factor. The effectiveness of the fungus is principally dependent on the presence of high densities of larvae in the second generation. The fungal disease, but not the other mortality factors, is capable of controlling the cloverworm population (Pedigo et al., 1983; Thorvilson and Pedigo, 1984). Among the parasitoids commonly attacking green cloverworm are several wasps (Hymenoptera: mostly Braconidae and Ichneumonidae) and flies (Diptera: Tachinidae) (Lentz and Pedigo, 1975; Mueller and Kunnalaca, 1979; Bechinski and Pedigo, 1983; Bechinski et  al., 1983a; Daigle et  al., 1988). The most abundant larval parasitoids are Cotesia marginiventris (Cresson) and Rogas nolophanae Ashmead (both Hymenoptera: Braconidae). Egg parasitism is infrequent, but predation of eggs and young larvae by Nabis americoferus Carayon and N. roseipennis Reuter (both Hemiptera: Nabidae) is documented (Sloderbeck and Yeargan, 1983b). Predation assumes greater importance in the pupal stage, when such predators as ground beetles (Coleoptera: Carabidae), field crickets (Orthoptera: Gryllidae), and rodents inflict heavy mortality. In addition to the aforementioned entomopathogenic fungus, a granulosis virus sometimes occurs (Carner and Barnett, 1975; Daigle et al., 1988). Life Cycle and Description. Annually, there are normally three generations in Iowa, with four flights of moths present in May, June-July, August, and September. The fourth flight may not be evident in some years. Green cloverworm fails to overwinter successfully in cold climates such as Iowa and reinvades the northern states each spring. The green cloverworm is reported to overwinter in the pupal and adult stages in the south, and as far north as southern Ohio. The overwintering of this species has not been intensively studied in southern states, but it remains reproductively active throughout the year along the Gulf Coast. It is thought to overwinter in the south as far north as southern Virginia, Kentucky, southern Missouri, and most of Texas. In the spring, when sustained winds blow from the southcentral states northward, green cloverworm moths are carried into northern areas (Wolf et al., 1987). The length of the life cycle is 40 days during the summer months. When reared at 30°C, the duration of the egg, larval, pupal, and egg to adult periods are about 5, 15.5, 8, and 28 days, respectively (Scott and Pedigo, 1977).

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Egg. Females normally deposit 200–230 eggs, but up to 670 eggs have been recorded from a single female. They are deposited singly. The egg initially is greenish, becomes speckled with orange or red, and then purplish gray just before hatching. The egg is a slightly flattened sphere in shape; the base, in particular, is flattened. The egg measures about 0.51 mm in diameter and 0.35 mm in height, and bears 14–19 readily discernible ridges. Hatching occurs 2–5 days after oviposition. Larva. There are 6–7 instars. The larvae grow from about 1.5 mm to over 30 mm long as they mature. The larvae are green throughout their development. Larvae bear a pair of longitudinal white stripes along each side, with a less distinct along the back, but they are fairly indistinct until about the third instar. During the terminal instar the white stripes fade, the insect appears almost entirely green. One of the most distinctive features is the presence of only four pairs of prolegs. The larva walks with a looping motion. Mean head capsule widths (range) for the larvae are 0.23 (0.13–0.28), 0.35 (0.32–0.43), 0.57 (0.48–0.70), 0.89 (0.66–1.00), 1.24 (0.70–1.50), 1.69 (1.35–2.00), and 1.88 (1.73–2.30) mm for instars 1–7, respectively. Duration of the instars was estimated by Stone and Pedigo (1972) to be about 3.1, 1.4, 1.9, 2.1, 2.5, 3.7, and 5.5  days, respectively, for a total larval duration of about 19  days when reared on soybean. Hill (1925) reported an average larval development period of 22.8 days when fed alfalfa. Larvae are solitary in their feeding behavior. Pupa. As the larvae are near completion of their development they spin a loose web in preparation for pupation. Pupation may occur in the plant canopy, whereby larvae usually web together with a leaflet and pupate within a leaf fold. Most larvae, however, drop to the soil and pupate at the surface or just beneath the surface. The pupa is brown to brownish black, and measures 11–15 mm long. Duration of the pupal stage is 9–12 days.

Adult. The adults are mottled grayish brown with black and silver markings. The male and female differ somewhat in appearance, however. The male has less distinctive markings, bearing about three irregular, transverse black lines across the forewing. The female also has transverse black markings but with greater contrast, and silver and reddishbrown areas distally. The hind wings are blackish brown. Wingspan measures 27–34 mm. The mouthparts of both sexes protrude, forming a distinctive snout. Moths hide in vegetation during the daylight hours. They become active at dusk, and reportedly feign death if disturbed, suddenly dropping to the soil with their wings folded. Once they take flight they are strong fliers, and their flight may continue until the early morning hours. Oviposition commences about 4–5  days after adult emergence and continues for 10 days or longer. Oviposition may occur on both the upper and lower leaf surfaces. Apparently moths prefer to oviposit on leaf surfaces that contain leaf hairs and deposit eggs preferentially on the lower leaf surfaces of alfalfa and clover because of the greater pubescence. On soybean, which is hairy on both surfaces, the females do not discriminate between locations. Moths preferentially deposit eggs on pubescent varieties of soybean relative to glabrous varieties (Pedigo, 1971). A good summary of green cloverworm biology was given by Pedigo et al. (1973), but more detailed description, particularly of insect morphology, was found in Hill (1925). The larva was included in keys by Crumb (1956), Oliver and Chapin (1981), Capinera (1986), and in a key to loopers in Appendix A. The adult occurred in a key by Capinera and Schaefer (1983).

Damage Larvae feed principally on the leaf tissue between the main veins of leaves. Most authors indicate that pods, blossoms, or stem tissue are rarely consumed. Larvae usually feed on the lower leaf surface, and instars 1–2 or 1–3 do not eat completely through the leaf tissue but leave the upper epidermis intact. Each larva eventually consumes over 100 cm2 of bean leaf tissue, with about 90% occurring in the last two instars. As the beans are very tolerant of defoliation, withstanding about 30% leaf tissue loss before yields are depressed, at least 5–6 mature larvae likely are necessary to inflict damage. In Delaware, green cloverworm larvae were frequently observed to feed on small pods of lima bean (Burbutis and Kelsey, 1970). However, the beans were very tolerant of injury, and larval densities of up to 8 per plant did not suppress yield.

Management FIG. 10.40  Adult of green cloverworm. (Photo by L. Buss.)

Sampling. The adult populations can be monitored with blacklight traps, though more males than females

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are ­captured. Eggs are deposited principally on the upper surface of leaves. Egg dispersion is random. A sequential sampling plan for eggs was developed by Buntin and Pedigo (1981). Sweep nets are usually used to sample larvae. Larval dispersion is aggregated, and a sequential sampling protocol was presented by Bechinski et al. (1983b). Insecticides. Green cloverworm rarely attains pest status in vegetable crops but can be controlled easily with foliar insecticides. Also, insecticide-containing baits are effective (Morgan and Todd, 1975). Bacillus thuringiensis products are not usually recommended. Cultural Practices. Cloverworm is most abundant late in the season, in late-maturing cultivars, and in narrow-row plantings (Buschman et al., 1981). Planting date apparently has little influence on damage (McPherson et  al., 1988). Tillage practices similarly have few consistent effects on green cloverworm populations (Sloderbeck and Yeargan, 1983a; Thorvilson et al., 1985a). Green cloverworm oviposits readily in alfalfa, and the first generation often occurs in this crop or clover before soybean or bean are available. Alfalfa is harvested frequently, however, and harvesting results in mortality of most larvae. Thus, alfalfa acts as a “sink” for the cloverworm population, causing a decline in abundance. It is the presence of soybean that generally leads to the great abundance of green cloverworm late in the season (Buntin and Pedigo, 1983). Alfalfa also acts as an early-season source for parasitoids and disease (Thorvilson et al., 1985b).

Okra Caterpillar

Anomis erosa Hübner (Lepidoptera: Erebidae)

very closely resembles cotton leafworm, Alabama argillacea (Hübner), so it may be misidentified and its abundance underestimated (Creighton, 1936). Okra caterpillar is also reported to feed on Peperomia sp., family Piperaceae. Natural Enemies. The natural enemies of okra caterpillar are mostly generalists that attack other caterpillars. For example, paper wasps, Polistes spp. (Hymenoptera: Vespidae) commonly feed on larvae, as do ground beetles (Coleoptera: Carabidae), stink bugs (Hemiptera: Pentatomidae), and assassin bugs (Hemiptera: Reduviidae). Parasitoids of okra caterpillar include Trichogramma sp. (Hymenoptera: Trichogrammatidae), Apanteles bedelliae Viereck (Hymenoptera: Braconidae), Itoplectis conquisitor (Say) (Hymenoptera: Ichneumonidae), Copidosoma truncatellum (Dalman) (Hymenoptera: Encyrtidae), Syntomosphyrum esurus (Riley) (Hymenoptera: Eulophidae), and Eusisyropa blanda (Osten Sacken) Diptera: Tachinidae). Creighton (1936) indicated that S. esurus was the most important parasitoid in his studies conducted in Florida. Life Cycle and Description. The life cycle of this insect is poorly documented but has been observed in the field from March to October in northern Florida and throughout the year in southern Florida. As a life cycle can be completed in about 35 days, several generations are possible annually. It is thought to overwinter in the pupal stage, but this is not satisfactorily proven. Egg. The egg is basically spherical but flattened at the point of attachment. It measures about 0.8 mm in diameter and bears 31–38 ribs, but most ribs fade before reaching the apex of the egg. Initially whitish, the egg soon turns greenish. They are deposited singly on foliage. Duration of the egg stage is about 4 days (range 3–6 days).

Natural History Distribution. This species is found in South America, Africa, southern Asia, and Australia in addition to North America. It is generally considered to be a southeastern species, and indeed it is most common there. However, it has been reported from as far north as Massachusetts and Montreal, Canada, and as far west as Kansas and Texas. Its distribution also extends southward through Mexico and the Caribbean region. Host Plants. Okra caterpillar feeds primarily on plants in the family Malvaceae. Okra is the only vegetable crop affected, but its caterpillar also feeds on ornamental or weedy plants such as rose-of-sharon, Hibiscus syriacus; swamp rose, Hibiscus moscheutos; cotton rose, Hibiscus mutilabilis; chinese mallow, Hibiscus sinensis; roselle, Hibiscus sabdariffa; flour-of-an-hour, Hibiscus trionum; velvet leaf, Abutilon theophrasti; flowering maple, Abutilon striatum; hollyhock, Althaea rosea; and round-leaved mallow, Malva rotundifolia. It can also be found on cotton, and though it is not generally thought to be a common pest of this crop, it

FIG. 10.41  Okra caterpillar. (Photo by J. Capinera)

Larva. The larva is reported to display 5–7 instars. Mean duration of the larval stage is about 16 days (range 13–22  days). It grows from about 2 mm long at hatching to about 35 mm long at maturity. The larva feeds during

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daylight hours, moves with a typical looper-like gait, and frequents the underside of the leaves. The larva is yellowish green or green, with a dark stripe dorsally, though it acquires white dorsolateral stripes along the length of its body. As the larva matures, an irregular and broad yellow stripe develop laterally above the spiracles. There are four pairs of prolegs, three abdominal plus the anal prolegs, present on the large larvae. The first pair of abdominal prolegs are reduced in size, relative to the other pairs. About 2 days before termination of the larval stage the larva folds a section of leaf and anchors it with silk; this is the site of pupation. Pupa. Initially, the pupa is bright green, but soon turns brown and eventually almost black. It measures about 15 mm long. The tip of the abdomen possesses short bristles and hooks which anchor the pupal case to silk webbing. Duration of the pupal stage is about 12  days (range 8–15 days).

FIG. 10.42  Okra caterpillar moth. (Photo by J. Capinera.)

Adult. The body of the adult is yellowish or rust. The forewing is marked with irregular yellowish, rust, and gray areas, though the hind wings are brownish and darker distally. Adults commence oviposition about 5  days after emergence and may continue egg production for 25 days. Adults are nocturnal. The biology of okra caterpillar was described by Chittenden (1913b), Dozier (1917), and Creighton (1936). The larva was included in the key by Crumb (1956).

Damage The larvae eat a large irregular hole in the leaves of okra, sometimes defoliating entire plants. It is not a common pest, even in the deep south, where it is most abundant.

Management This caterpillar rarely is abundant enough to warrant suppression, but insecticides applied to the foliage are effective, especially if applied when the larvae are young.

Saltmarsh Caterpillar

Estigmene acrea (Drury) (Lepidoptera: Erebidae)

Natural History Distribution. This native insect is found widely in North and Central America. It is abundant enough to be damaging everywhere in the United States, and in Canada, it damaged crops in Ontario and Quebec. It is most serious as a pest in the southern United States, particularly the southwest. Host Plants. Saltmarsh caterpillar’s peculiar common name is derived from the initial description as a pest of saltgrass hay grown in the vicinity of Boston. This is an anomaly, and despite the wide host range of this insect, grasses are not particularly preferred. Broadleaf weeds are the normal host plants, but larvae commonly disperse from these late in the growing season to damage vegetable and field crops. Vegetables injured include asparagus, bean, beet, cabbage, carrot, celery, corn, lettuce, onion, pea, tomato, turnip, and probably others. Field crops damaged are alfalfa, clover, cotton, soybean, sugarbeet, and tobacco. The favored weed host seems to be pigweed, Amaranthus spp., but many others may be consumed, including anglepod, Gonolobus sp.; sicklepod, Cassia tora; dog fennel, Eupatorium capillifolium; ground cherry, Physalis spp.; and mallow, Anoda sp. Natural Enemies. Saltmarsh caterpillar larvae frequently are parasitized, particularly by tachinids (Diptera: Tachinidae). In Arizona, the most common parasitoids were Exorista mellea (Walker) and Leschenaultia adusta (Loew), but Gymnocarcelia ricinorum Townsend and Lespesia archippivora (Riley) were also observed (Taylor, 1954). Jackson et  al. (1970) documented the biology and importance of L. adusta. Arnaud Jr. (1978) reported additional species of tachinids associated with saltmarsh caterpillar. Hymenopteran parasitoids are known from both the larval and egg stages (Taylor, 1954; Taylor and Stern, 1971), and include Apanteles diacrisiae Gahan (Braconidae); Therion fuscipenne (Norton), T. morio (Fabricius), Casinaria genuina (Norton), Hyposoter rivalis (Cresson) (all Ichneumonidae); Psychophagus omnivorus (Walker), Tritneptis hemerocampae Vierick (both Pteromalidae); Anastatus reduvii (Howard) (Eupelmidae); and Trichogramma semifumatum (Perkins) (Trichogrammatidae). A cytoplasmic polyhedrosis virus is known (Langridge, 1983), but there are little data on importance. General predators such as lady beetles (Coleoptera: Coccinellidae), softwinged flower beetles (Coleoptera: Melydridae), and assassin bugs (Hemiptera: Reduviidae) prey on these caterpillars but are not thought to be very important in population regulation (Young and Sifuentes, 1959). Life Cycle and Description. Total generation time requires 35–40 days under ideal conditions, but most reports from the field suggest about 6 weeks between generations.

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The number of generations per year is estimated at one in the northern states to 3–4 in the south. Overwintering reportedly occurs in the mature larval stage, with pupation early in the spring. Saltmarsh caterpillars are usually infrequent early in the season, but may attain high numbers by autumn. Egg. The eggs are nearly spherical in shape and measure about 0.6 mm in diameter. Initially, they are yellow but soon become grayish. Females commonly produce 400–1000 eggs in one or more clusters. It is not unusual to find a single egg mass containing 1200 eggs. Eggs hatch in 4–5 days.

and 8 days, respectively, for a larval period of 20–22 days, depending on diet. However, some studies have reported longer larval periods, up to about 45 days. Larvae are active dispersers, a habit that is relatively uncommon among caterpillars. Most commonly, late instar larvae are found individually or in large numbers ambling over the soil, searching for suitable food. Damage to margins of crop fields often occurs as such larvae desert drying weeds for irrigated crops. Stracener (1931) reported that young larvae drop readily from plants when disturbed, spin a strand of silk, and are blown considerable distances by wind. Frequency of distribution by wind is unknown. Pupa. Pupation occurs on the soil among leaf debris, in a thin cocoon formed from silken hairs interwoven with caterpillar body hairs. The dark brown pupa measures about 30 mm long. Duration of the pupal stage is about 12–14 days.

FIG. 10.43  Saltmarsh caterpillar larva. (Photo by L. Buss.)

Larva. There are 5–7 instars. This description is based on Hinds (1904), who observed five instars in Texas. Upon hatching, the larvae are about 2 mm long, brown in color, and bear numerous long hairs over the entire length of the body. During this stage, and the subsequent instar, larvae feed gregariously on the lower leaf surface, usually failing to eat entirely through the leaf. Larvae attain a length of about 10 mm during the first instar. Second instars display longitudinal stripes, usually brown, yellowish, and white, and the body hairs become darker. Larvae attain a length of about 15 mm. During the third instar, larvae become darker, but a consistent color pattern is not apparent. Larvae attain a length of about 30 mm. In the fourth and fifth instars, larvae maintain the same general appearance as earlier stages, but grow to a length of about 45 and 55 mm, respectively. Larvae usually are dark, but sometimes are yellowish brown or straw colored. The larvae are marked by long body hairs, and these too range from cream or grayish to yellowish brown to dark brown. Although they are decidedly hairy, the hairs are not as dense or stiff as those found in woollybear larvae. Duration of larval development is 24–37  days. In contrast, Young and Sifuentes (1959) and Capinera (1978b) reported six instars in Mexico and Colorado, respectively. Development time of the six instars was about 3, 2, 2, 2, 3,

FIG. 10.44  Saltmarsh caterpillar adult. (Photo by L. Buss.)

Adult. Adults are fairly large moths, measuring 3.5–4.5 cm in wingspan, and are distinctive in appearance. They are predominantly white, though generally, the wings bear numerous, small, irregular black spots. The hind wings of the male are yellow; those of the female are white. The underside of the male’s front wings may also be tinted yellowish. Most of the abdominal segments are yellow and bear a series of large black spots dorsally. A sex pheromone is known (Hill and Roelofs, 1981). Females release pheromone in a bimodal manner, with peaks of release occurring about 4–6 h after onset of scotophase, and then again at about 10 after scotophase (Del Maxo-Cancino et al., 2004). Mating may occur in the evening following emergence, though most females call and mate days 2–4 after emergence. Egg deposition occurs the evening following mating. Females usually live only 4–5 days, but may produce more than one cluster of eggs. Accounts of saltmarsh caterpillar biology were provided by Hinds (1904), Stracener (1931), and Young and

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Sifuentes (1959). Keys that included saltmarsh caterpillar include Capinera (1986) and Sparks and Liu (2001). Rearing methods were given by Dunn et al. (1964) and Vail et al. (1967a).

Damage Larvae are defoliators. Young larvae feed gregariously and skeletonize foliage. Older larvae are solitary and eat large holes in leaf tissue. On celery, most feeding is restricted to leaf tissue, unlike many other lepidopterous species, which may also feed readily on petioles (stalks) (Jones and Granett, 1982). Older larvae may disperse long distances in search of food, sometimes moving in large numbers. Commonly this is associated with maturation of cotton or weeds in the autumn. Thus, these caterpillars tend to be damaging to fall-planted vegetable crops. Foliage consumption at least doubles with each succeeding instar and mature larvae can consume over 13 cm2 of thick-leaved foliage, such as sugarbeet, daily (Capinera, 1978b). Capinera et al. (1987) measured bean foliage consumption by each instar and recorded over 400 cm2 of foliage consumed during the life of a caterpillar. Further, they estimated that 1.0–1.5 mature caterpillars per plant could inflict 20% defoliation, a level adequate to cause yield loss.

Management Insecticides are commonly used to suppress saltmarsh caterpillars if they become abundant in vegetable crops. Baits are not effective. Most damage occurs at field margins as larvae disperse into crops from nearby senescent vegetation. Both chemical insecticides and Bacillus thuringiensis are recommended. Physical barriers, including ditches or trenches with steep sides, can be used to interrupt invasion of crops by caterpillars.

Yellow Woollybear

Spilosoma virginica (Fabricius) (Lepidoptera: Erebidae)

Natural History Distribution. Yellow woollybear is a native insect and is found throughout the United States and southern Canada. Yellow woollybear’s range as a pest is generally restricted to the Great Plains region to the west coast. Even within this area, however, it is infrequently numerous enough to be damaging. Host Plants. Yellow woollybear is a very general feeder and reported from over 100 different plants. Yellow woollybear has been observed to damage such vegetable crops as asparagus, bean, beet, cabbage, cantaloupe, carrot, cauliflower, celery, corn, eggplant, lima bean, parsnip, pea, potato, pumpkin, radish, rhubarb, squash, sweet potato, Swiss chard, turnip, and watermelon. Other economic plants

damaged include field crops such as alfalfa, peanut, and ­sugarbeet; fruits such as blackberry, cherry, currant, gooseberry, grape, and raspberry; and flowers such as canna, dahlia, geranium, hollyhock, hyacinth, and verbena. Among weeds fed upon are dandelion, Taraxacum officinale; dock, Rumex sp.; pigweed, Amaranthus spp.; lambsquarters, Chenopodium album; plantain, Plantago major; Russian thistle, Salsola kali; Spanish needle, Bidens bipinnata; and sunflower, Helianthus spp. Natural Enemies. The number of parasitoids found in association with these insects is quite large, though the importance of these natural enemies has not been well studied. Arnaud Jr. (1978) listed several tachinids reared from yellow woollybear, including Aplomya caesar (Aldrich), Blondelia hyphantriae (Tothill), Bombyliopsis abrupta (Wiedemann), Carcelia diacrisiae Sellers, C. reclinata (Aldrich and Webber), Compsilura concinnata (Meigen), Exorista mella (Walker), Gymnocarcelia ricinorum Townsend, Hubneria estigmenensis (Sellers), Lespesia aletiae (Riley), L. frenchii (Williston), Mericia ampelus (Walker), Thelaira americana Brooks, and Winthemia datanae (Townsend) (all Diptera: Tachinidae). Wasps reared from yellow woollybear include Apanteles diacrisiae Gahan, A. scitulus Riley (both Hymenoptera: Braconidae); Coccygomius sanguinipes (Vierick), Cratichneumon unifasciatorius (Say), Vulgichneumon subcyaneus (Cresson), Therion morio (Fabricius), Hyposoter rivalis (Cresson), Enicospilus glabratus (Say) (all Hymenoptera: Ichneumonidae); Psychophagus omnivorous (Walker), Tritneptis hemerocampae Vierick (both Hymenoptera: Pteromalidae); Elachertus marylandicus Girault, E. spilosomatis Howard (both Hymenoptera: Eulophidae); Telenomus nigriscapus Ashmead, and T. spilosomatis Ashmead (both Hymenoptera: Scelionidae). The fungus Beauveria bassiana has been reported to cause low levels of mortality, and a granulosis virus has been observed (Boucias and Nordin, 1977). Life Cycle and Description. There are likely three generations of yellow woollybear annually, despite the numerous reports of only two generations. The discrepancy is due to the overlapping flights of the moths from overwintering pupae with those of the spring generation. In the most complete study of yellow woollybear population dynamics, conducted in Iowa, the apparent spring flight of moths was shown to consist of two reproductive populations, each represented by separate peaks in abundance within the overall spring flight period. There was also a late summer flight which produced overwintering pupae (Peterson et al., 1993). Elsewhere, moths also are abundant in spring (April-June) and autumn (July-October), but there is considerable geographic variation in the timing of flights. For example, the early-season flight activity occurs in mid-April in Arkansas and North Carolina, but not until late May in Maine. Lateseason flight occurs in August in Maine, September in

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North Carolina, and October in Arkansas. The pupal stage reportedly overwinters throughout this insect’s range. Egg. The spherical eggs of yellow woollybear are yellow, 0.6 mm in diameter, and deposited in clusters of 50–200 both on plant foliage and on inedible substrata. Duration of the egg stage is about 7 days.

FIG. 10.46  Adult of yellow woollybear. (Drawing by J. Capinera.)

FIG.  10.45  Larva of yellow woollybear caterpillar. (Drawing by J. Capinera.)

Larva. Upon hatching, the larvae are hairy and bluish white. During the first two instars, the larvae feed gregariously and then disperse. As they mature, they retain their hairy characteristic and develop both long, fine, soft hairs and shorter, stout bristles. The hairs are not so thick as to hide the larval body, which is quite variable in coloration. The most common color for larvae is yellow, which is the basis for the common name, but they also may be cream, light brown or dark brown. Also, there is a dark line along each side of the caterpillar, and the membrane between each body segment tends to be marked by dark pigment. The head is principally yellow, and this character is useful to distinguish it from saltmarsh caterpillar, Estigmene acrea (Drury), which tends to have a black head. Larvae attain a length of up to 5 cm at maturity. Larvae reared on bean foliage at 25°C required about 39 days to progress through nine instars (Capinera et  al., 1987). Instar duration was 3.1, 2.4, 3.9, 4.9, 3.6, 3.6, 3.6, 4.6, and 10.0 days for instars 1–9, respectively. Although Peterson et  al. (1993) suggested that this long development time or large instar numbers might be indicative of a suboptimal host, Dethier (1988) similarly reported larval development times of 35  days on suitable host plants. Larvae often move to another plant after completing a meal, even though the plant is relatively suitable for growth and development. They spend less than 1% of their time eating, 2.5% in wandering, and the remaining of their existence in resting (Dethier, 1988). Pupa. Pupation occurs in plant debris, under the bark of trees, and in other sheltered locations. The pupal case is constructed from the larval hairs, which is held together loosely with silk. Duration of the pupal stage is 7–14  days, and the reddish-brown pupa measures about 15–16 mm long.

Adult. Adults are medium-size moths measuring about 38–50 mm in wingspan. The wings are white, but the front wings bear a small black spot near the center, and the hind wings usually are with three black spots. The head and thorax are covered with white scales. The abdomen is yelloworange with three rows of black spots, one row dorsally and one on each side. The biology of yellow woollybear is not well documented. A brief treatment of yellow woollybear was provided by Riley (1871), Marsh (1912b), and Maxson (1948). Keys that included woollybear caterpillar include Capinera (1986) and Sparks and Liu (2001).

Damage Larvae are defoliators. Young larvae are gregarious and tend to feed together on the underside of foliage and skeletonize the plant tissue. Larger larvae disperse and feed sporadically, creating irregular holes in foliage. When larvae are particularly numerous, and succulent vegetation scarce, many of the large larvae remain on crops and inflict injury. This most often occurs in irrigated cropland when adjacent weedy vegetation senesces, or dries up due to drought. Typically it is only the late summer generation that attains densities adequate to inflict injury. Capinera et  al. (1987) measured bean foliage consumption by each instar and recorded over 300 cm2 of foliage consumed during the life of a caterpillar. Further, they estimated that 1.2–2.2 mature caterpillars per plant could inflict 20% defoliation, a level adequate to cause yield loss.

Management Yellow woollybear is common among weeds growing along roadsides, fence rows, and irrigation ditches. Larvae disperse into crops only when native or weedy vegetation is depleted or otherwise unsuitable. Such sources of infestation should be monitored. Larvae are easily killed with foliar insecticides, though this is rarely warranted. Treatment of the source of infestation or the borders of crops is generally adequate to prevent damage. Burning of crop residues in the autumn is sometimes recommended to destroy overwintering larvae and pupae

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because they are located above ground and very susceptible to fire. However, it is normally better to leave such organic matter on the soil surface, or tilled into the soil; the exception might be ditch banks or other small areas that are not tilled.

FAMILY GELECHIIDAE—LEAFMINER MOTHS Eggplant Leafminer

Keiferia inconspicuella (Murtfeldt) (Lepidoptera: Gelechiidae)

Natural History Distribution. Eggplant leafminer is present in the southeastern and midwestern United States north to about New Jersey and Iowa, and west to Nebraska and Texas. This species has a rather confused history, but records from most western states appear to be due to related species. Eggplant leafminer is a native insect. Host Plants. Eggplant leafminer apparently limits its attacks to eggplant and horsenettle, Solanum carolinense. Horsenettle is the natural host. Natural Enemies. Several parasitoids are known and apparently effective in keeping this insect from becoming very numerous. Among the common species are Apanteles epinotiae Viereck, Bracon gelechiae Ashmead, Cardiochiles sp., Macrocentrus delicatus Cresson, Orgilus mellipes Say, Agathis gibbosa (Say) (all Hymenoptera: Braconidae); and Chrysonotomyia sp. and Miotropis sp. (both Hymenoptera: Eulophidae). Cirrospiloides bicoloriceps (Girault) and Campoplex phthorimaeae (Cushman) also parasitize this leafminer. A eumenid, Parancistrocercus fulvipes (Saussure) (Hymenoptera: Eumenidae), has been observed to prey upon larvae by digging them from their tunnels. Gross and Price (1988) observed mean parasitism rates of about 33% in Illinois, and equivalent levels of parasitism on larvae developing in both eggplant and horsenettle. Life Cycle and Description. Eggplant leafminer can complete its development, from the egg to the adult stage, in about 25 days when cultured at 27°C. In Illinois, eggplant leafminer was active from June to September, and underwent three generations. Egg. The eggs are deposited singly on the leaf surface, with deposition occurring on both the upper and lower surfaces, but the lower surface is heavily favored. They are somewhat cylindrical, but with rounded ends. Mean egg length (range) is 0.34 mm (0.30–0.38 mm); mean width (range) is 0.19 mm (0.16–0.21 mm). Egg color is yellow. Duration of the egg stage is about 7 days.

FIG. 10.47  Eggplant leafminer larva. (Drawing by USDA.)

Larva. Larvae burrow within the leaf along the edge of the leaf blade. If the larvae do not hatch at the leaf edge, they may construct a small linear mine near the egg, but soon relocate to the leaf margin. Other than to move to the leaf edge, larvae do not leave the mine and do not web together foliage. Larvae form a blotch-shaped mine and deposit feces and silk within the mine. There are five instars, with mean head capsule widths of 0.17, 0.27, 0.38, 0.57, and 0.76 mm for instars 1–5, respectively. The larva initially is white or pale yellow except for the head and thoracic shield, which are brown. By the third instar the larva acquires a brownish or greenish color, and in the fourth or fifth instar becomes dark green, turquoise, or dark blue. The thoracic legs are light in color. Mature larvae attain a length of 7–8 mm and are slightly flattened in form. Pupa. When ready to pupate, mature larvae spin down to the soil on a strand of silk. Pupation normally occurs within a silken cocoon in the soil, and usually quite close to the soil surface. The pupa is dark blue when first formed, but becomes dark brown with maturity. Eggplant leafminer pupae have no distinctive features, resembling most moth pupae. They measure 3.4–5.2 mm long and 1.0–1.7 mm wide.

FIG. 10.48  Eggplant leafminer adult. (Drawing by USDA.)

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Adult. The adult is a small grayish-brown moth that is marked with yellowish brown. The forewings, but especially the hind wings, bear a long fringe of hairs. The wingspan measures 10–14 mm. Adults are nocturnal. Eggplant leafminer is quite similar in appearance to potato tuberworm, Phthorimaea operculella (Zeller), and they share common hosts. However, eggplant leafminer tends to mine leaf margins, whereas potato tuberworm attacks the central areas, particularly the area of the main veins. Also, eggplant leafminer does not leave the mine to web leaves. Poos and Peters (1927) gave the morphological characters useful in distinguishing the species, but accurate determination is best accomplished by an authority. The biology of eggplant leafminer was given by Jones (1923) and Gross (1986). A key that included eggplant leafminer larvae (as Keferia glochinella Zeller) is provided by Capps (1946).

Damage Though commonly found mining eggplant in southern states, eggplant leafminer is not considered to be a serious pest. The leaf edge is preferentially mined, acquiring a dry, sometimes swollen, blotch.

Management This insect is not known to be a serious pest and normally, its presence should not be a cause for concern. However, insecticides applied to the foliage should be effective if suppression is warranted.

Potato Tuberworm

Phthorimaea operculella (Zeller) (Lepidoptera: Gelechiidae)

Natural History Distribution. Potato tuberworm occurs widely in the United States but normally is absent from the northernmost states and Canada. It is most common in southern areas, particularly California and the southeastern states north to Maryland. Potato tuberworm appears to be native to North America but is now spread throughout the world. Areas with warm, dry climates such as southern Europe, northern and southern Africa, India, Australia, and Central and South America all experience problems with potato tuberworm. The transport of tubers infested with insects causes extensive dissemination of this pest and also results in occurrence records where this insect does not exist permanently. Host Plants. This insect feeds almost entirely on members of the plant family Solanaceae. Vegetable crops supporting potato tuberworm include eggplant, pepper, potato, and tomato, though potato is the only frequent host. Tobacco is occasionally affected, and potato tuberworm is sometimes called “tobacco splitworm” when it is ­associated

with this host. Solanaceous weeds such as bittersweet, Solanum dulcamara; black nightshade, S. nigrum; groundcherry, Physalis spp.; henbane, Hyoscyamus sp.; horsenettle, S. carolinense; jimson weed, Datura stramonium; and matrimony vine, Lycium europaeum; also serve as hosts. A worldwide host list was provided by Das and Raman (1994). Natural Enemies. Natural enemies affect the egg, larval, and pupal stages of potato tuberworm, though they are much more effective when the tuberworms are feeding on the aerial portions of the plant rather than within tubers. Among the parasitoids known to affect potato tuberworm are numerous species of Braconidae, Encyrtidae, Eulophidae, Ichneumonidae, Mymaridae, Pteromalidae, Scelionidae, and Trichogrammatidae (all Hymenoptera). Reported to be the most abundant in Virginia were Bracon gelichiae Ashmead (Braconidae) and Campoplex sp. (Ichneumonidae) (Hofmaster, 1949). In California, Apanteles dignus Muesebeck (Braconidae) was the dominant parasitoid (Oatman and Platner, 1989), though in an earlier report by the same authors (Oatman and Platner, 1974), Agathis gibbosa (Say), A. scutellaris Muesebeck, and Campoplex phthorimaeae (Cushman) (all Hymenoptera: Braconidae) were most common. Many of the parasitoids of tuberworm were discussed and pictured by Graf (1917). There have been several attempted introductions of parasitoids to North America, but with few successes (Clausen, 1978). Other natural enemies are less important. Several general predators have been noted to feed on tuberworm, including the ants Pheidole and Lasius spp. (Hymenoptera: Formicidae), pirate bugs (Hemiptera: Anthocoridae), shield bugs (Hemiptera: Pentatomidae), and rove beetles (Coleoptera: Staphylinidae). Diseases have been noted (Briese and Mende, 1981; Trivedi and Rajagopal, 1992), but seem to be of little natural significance. Weather. Weather is thought to affect the abundance of potato tuberworm. Summers that are unusually warm and dry favor increase in tuberworm populations (Langford and Cory, 1932; Hofmaster, 1949). Life Cycle and Description. A life cycle may be completed in 15–90  days, resulting in about five generations annually in both California and Virginia; in other locations around the world the number of generations is reported to range from 2 to 13 annually. In warm climates, the generations overlap and cannot be distinguished easily. Potato tuberworm normally cannot withstand freezing, so in cold climates overwintering survival by larvae is poor except within potatoes in storage or in cull piles. Populations develop over a range of 10–35°C with an optimum of 28– 30°C. Degree-day requirements are about 65, 165, and 107 degree-days for the egg, larval, and pupal stages, respectively (Sporleder et al., 2004). Egg. The eggs are deposited singly or in poorly defined clusters, usually on the underside of leaves. If deposited

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on potatoes in storage, however, the egg clusters tend to be larger, up to 30 eggs. Tubers tend to be heavily infested if they are exposed, that is, not covered with soil. There are also reports of oviposition on soil adjacent to plants (Traynier, 1975). The egg is elliptical and measures about 0.48 mm long and 0.36 mm wide. Initially white in color, they turn yellow and acquire a distinct iridescence with age. Duration of the egg stage is only about 5 days during the summer but may reach 30 days during cool weather.

a silk cocoon that is usually covered with leaf trash, fecal material, soil particles, and other debris. Initially white or yellow, the pupa eventually becomes dark mahogany. The form of the pupa is typical of Lepidoptera, wider at the anterior end, tapering to a point at the posterior end, and with the partially developed wings twisted ventrally. The tip of the abdomen bears a hook and a circle of spines. It measures about 6 mm long. Mean duration (range) of the pupal period is 11.6 days (range 8–14 days).

FIG. 10.50  Adult potato tuberworm. (Drawing by USDA.)

FIG. 10.49  Potato tuberworm larvae (Drawing by USDA.)

Larva. At hatching, larvae normally begin to burrow almost immediately. Larvae normally mine the leaves, but occasionally the petioles and stems, and sometimes burrow into tubers. The older or lower leaves are preferred. Larvae often plug the entrance to their burrow with excrement but extrude the cast skins and head capsules. Sometimes considerable amounts of silk are produced by larvae, usually when they are forced to traverse the leaf surface, but also to plug larval burrows and to web together leaves. There are four larval instars. Mean head capsule widths are about 0.20, 0.36, 0.60, and 1.13 mm for instars 1–4, respectively. Body lengths are about 1.1, 2.0, 4.5, and 7.0 mm, respectively. Mean duration (range) of the instars is about 3.5 (2–6), 2.5 (2–3), 3.1 (2–4), and 7.3 (5–12) days, respectively. Initially white in color with a black head and thoracic plate, the larva acquires additional color as it grows. In the mature larva, the head, thoracic plate, and thoracic legs are black. The body is principally white, with pink or greenish pink dorsally. There are five pairs of prolegs. The anal plate is yellow. Duration of the larval period may require only 14  days during the summer, but up to 70  days during the winter months. Pupa. Pupation occurs in the soil, or just beneath the epidermis of the leaf or tuber. Before pupation, the larva spins

FIG. 10.51  Adult potato tuberworm. (Drawing by USDA.)

Adult. The adult stage is a small grayish-brown moth with a wingspan of 12–16 mm. The wings, especially the hind wings, are fringed. The front wings are marked with dark spots, which usually coalesce to form a dark longitudinal streak or a row of dark spots. The wings, abdomen, and legs also are tinged with yellow scales. The moths are nocturnal, hiding during the day beneath debris and clods of soil. Mating occurs within 2  days of moth emergence. Oviposition is usually completed in 6–17  days, with females each producing about 150–250 eggs. Longevity rarely extends beyond 21 days. A sex pheromone has been identified and can be used for trapping under field conditions (Persoons et al., 1976). Potato tuberworm is easily confused with eggplant leafminer, Keiferia inconspicuella (Murtfeldt). These species are similar in appearance and have overlapping host range. Potato tuberworm moths are usually larger, with yellow scaling more distinct and forming longitudinal streaks, but accurate differentiation is best accomplished by an authority. Poos and Peters (1927) presented differences in the genitalia of adults and discussed procedures to distinguish the other stages. Eggplant leafminer does not attack potato or tobacco though it shares eggplant and horsenettle with

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potato tuberworm. On eggplant and horsenettle, mines of potato tuberworm begin at the midrib on one of the principal veins, whereas mines of eggplant leafminer begin near the leaf margin. Excellent treatment of potato tuberworm biology was given by Graf (1917) and Poos and Peters (1927). The reports of Clarke (1901) and Hofmaster (1949) were also useful. A brief worldwide review of this insect was published by Trivedi and Rajagopal (1992). A key to vegetable insects that included potato tuberworm was published by Sparks and Liu (2001). Culture of potato tuberworm was described by Platner and Oatman (1968).

Damage Leaf mining is the most common habit of potato tuberworm, but mining of the tuber is the most damaging. Mining normally is restricted to the foliage so long as it is green and succulent. Larvae may also mine the stems, usually working downward. If the tuber is attacked, the mining may occur near the epidermis, or “skin” of the tuber, or larvae may burrow deeply. The tunnels in potato tubers normally fill with fungus (Graf, 1917). Tunneling not only destroys the food quality of the tubers but also the sprouting potential of tubers that are used for propagation. In tomato, foliage is initially attacked, but larvae can mine through the fruit stem into the fruit (Gilboa and Podoler, 1995).

Management Sampling. Pheromone traps are effective for monitoring potato tuberworm populations, and usually, there is a good correlation between trap catches and damage levels (Shelton and Wyman, 1979b; Yathom et  al., 1979). Pheromonebaited water pan traps are more effective than pheromonebaited sticky traps (Bacon et al., 1976), though funnel traps seem to be as effective as water traps (Raman, 1988). Sticky traps are prone to be covered with dust, thereby reducing catch (Kennedy, 1975). Larval sampling was discussed by Horne (1993), and a binomial sequential sampling plan for tuberworm in tomato was developed by Gilboa and Podoler (1995). Insecticides. Tuberworm often is controlled by the application of insecticide to foliage (Bacon, 1960), though in some parts of the world resistance to insecticides is a problem (Collantes et al., 1986). Also, insecticides interfere with predators and parasitoids of tuberworm, which can be quite effective, so it is prudent to determine that tuberworm is present in potentially damaging numbers before implementing an insecticide-based management effort (Shelton et al., 1981). Integration of chemical insecticides with cultural practices is effective (Fuglie et al., 1993). Biological insecticides, particularly the bacterium Bacillus thuringiensis, are recommended for protection of potato tubers in storage, but not usually in the field.

Suppression in the field is possible, but several applications may be required (Broza and Sneh, 1994). A granulosis virus has been used experimentally as a suppressive bioinsecticide under field conditions (Kroschel et  al., 1996). The repellency of vegetation to potato tuber moth was evaluated with vegetation from five fragrant plant species: Ambrosia, Anemone, Eupatorium, Eucalytus, and Lantana. Lantana was most effective at protecting potato tubers in storage from infestation, reducing damage from 70% in the check to only 5% in the presence of Lantana (Lakshman, 1987). Cultural Practices. Cultural practices can greatly affect the susceptibility of potato-to-potato tuberworm. Overwintering population tend to be low, with tuberworm populations increasing through the year. Thus, areas, where more than one potato crop is cultivated, tend to experience greater loss by tuberworm, and greatest damage occurs late in the season. In some regions, potato production is limited to the spring months to eliminate the nearly year-long availability of potatoes for tuberworm breeding. Sanitation is extremely important in potato tuberworm management. Potatoes held in storage or in cull piles are potential sources of infestation. Similarly, potatoes left in the field, volunteer plants, and solanaceous weeds can support tuberworms. Harvested potatoes should not be left in the field overnight as this is when oviposition occurs. If vines are killed before senescence and tubers harvested soon thereafter, the level of tuber infestation is low. Delayed harvest increases the exposure of tubers to ovipositing moths. Infestation of tubers is especially likely if there are cracks in the soil, allowing access by tuberworm. Soil depths of 5 cm or more protect tubers from infestation. Sandy soil can also be a problem if rainfall washes away soil, exposing tubers. Irrigation practices greatly affect soil condition, with furrow irrigation producing more cracks than overhead irrigation. Frequent irrigation helps to prevent soil cracking. Hilling of the soil, wherein soil is scraped from between the rows and deposited at the base of the plants, helps to deny access by tuberworm to tubers (Langford, 1933; Shelton and Wyman, 1979a, b; von Arx et al., 1990). Deep planting of potato seed, and culture of varieties that do not produce shallow tubers, also reduce the incidence of tuberworm damage. Ali (1993) assessed various cultural practices in Sudan and reported that early planting, deep planting, frequent hilling-up, irrigation, mulch, and neem mulch all reduced infestation by potato tuber moth. Light but frequent irrigation, and neem mulch were the most beneficial practices. Some differences in tuber susceptibility or suitability exist among cultivars (Fenemore, 1980). Oviposition preference, percent pupation, and moth fecundity are affected, but the significance of preference diminishes when moths are confronted with a no-choice situation. Jansens et al. (1995) and Davidson et al. (2004) demonstrated that

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Bacillus thuringinesis genes could be inserted into potato, resulting in decreased damage to leaves and tubers. The sex pheromone can be used to manipulate populations. Mass trapping can be used to reduce damage in the field. Trapping, and disruption of mating by saturation of the atmosphere and confusion of the moth works best for potatoes in storage (Raman, 1988).

Tomato Leafminer

Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae)

Natural History Distribution. This species is native to South America, where it occurs widely in tropical and subtropical regions, and north to (including) Panama. It recently has become established in southern Europe, most of Africa, and portions of the Middle East. In Europe, it also occurs in northern areas, but mostly in greenhouses. Currently, it does not occur in North America. In southern regions of the United States, a closely related species, tomato pinworm (Keiferia lycopersicella [Walshingham] [Gelechiidae]), occupies the ecological niche of tomato leafminer on tomatoes. Apparently, T. absoluta and K. lycopersicela can co-occur, as both are reported from northern South America. Tomato leafminer is a major threat to North America and given its redistribution in recent years to previously unoccupied parts of the world, it is a likely invader. The history and distribution of T. absoluta are described by Desneux et al. (2010), Zappalá et al. (2013), and Campos et al. (2017). Host Plants. Tuta absoluta feeds mostly on plants in the family Solanaceae and is a pest principally of tomato. However, it also feeds on eggplant, pepper, potato, and (rarely) bean (Fabaceae). Among weeds that can host this insect are jimson weed, Datura stramonium L.; black nightshade, Solanum nigrum L., Jerusalem cherry, Solanum pseudocapsicum L.; and sticky nightshade, Solanum sisymbriifolium Lamb. (all Solanaceae). Natural Enemies. A number of predators and parasitoids are known. Their geographic distribution varies and they differ in effectiveness. Desneux et al. (2010) provide a worldwide list of beneficial insects, though emphasizing the Mediterranean region. Zappalá et al. (2013) provide a more review of natural enemies that is not materially different. These reviews document that wasps in the families Eulophidae and Braconidae are important in parasitizing leafminer larvae, and the Trichgrammatidae parasitize eggs. Among the Hemiptera, several zoophytophagous mirids are quite important, both in fields and in greenhouses. Nabids can also be effective under field condition. These Hemiptera feed on both eggs and young larvae. The mirid Nesidiocoris tenuis Reuter has been shown to be especially effective as an augmentative release organism in greenhouses (Calvo et al., 2012). The other important taxon is mites (Acari) in

the family Pytoseiidae, which feed on leafminer eggs. The bioinsecticides Bacillus thuringiensis and Beauveria bassiana are compatible with most natural enemies. Miranda et al. (1998) provide a life table analysis that documents the importance of natural enemies under Brazilian conditions; in this study, predators were particularly important. Life Cycle and Description. In suitable habitats, this species undergoes 10–12 generations per year. The life cycle can be completed in about 30–35 days. Apparently, this insect can overwinter in the egg, pupal, or adult stage, and whether diapause occurs is not known, but does not seem likely. Egg. The eggs are elongate-oval, and about 0.36 mm long and 0.22 mm wide. Normally they are deposited singly on the underside (abaxial surface) of leaves. The eggs initially are light-colored, either white or yellow, but darken as the embryos develop, and are nearly black as the eggs near the hatch. Mating and oviposition normally commence 1–3 days after emergence of the moth. Most of the eggs are deposited within 5  days of commencing oviposition, and 90% within 10 days. Eggs hatch in about 4–7 days. Larva. Newly hatched larvae are white or nearly transparent, with a dark head and two dark spots on the pronotal shield, but become mostly green or pink as they feed and grow. Like many leafminers, the larvae are slightly flattened dorsoventrally, allowing them to live within relatively thin leaves. Usually, four instars are found in this species. Vargas (1970) reported that the mean (range) values for body length were 1.61 (1.40–1.90), 2.8 (2.45–3.10), 4.69 (3.85–5.65), and 7.72 (5.50–9.20) mm for instars 1–4, respectively. Similarly, maximum head capsule widths were given as 0.15 (0.15–0.18), 0.25 (0.24–0.28), 0.39 (0.35– 0.43), and 0.83 (0.70–0.98) mm for instars 1–4. Duration of the larval stage is often about 10–30 days when cultured at 25°C on various diets, but on tomato, the most favored host, it is about 10 days (Mihsfeldt and Parra, 1999). The mature larva (prepupa) often leaves the mine or fruit where it was feeding and constructs a silk cocoon on the foliage or soil. Alternatively, the larva may remain in the leaf mine or fruit for pupation, in which case a cocoon in not constructed. Pupa. Initially, the pupa is green but turns dark brown or reddish brown before the emergence of the adult. Pupation may occur on the plant or in the soil. In the soil, it is typically found at a depth of 1–2 cm. Duration of the pupal stage is 8–10 days when reared at 25°C on various diets, but normally 10–13  days under field conditions. The pupal period when reared on tomato at 25°C is about 8  days. Male pupae are about 4.3 mm long and 1.2 mm wide, whereas female pupae are about 4.7 mm long and 1.4 mm wide.

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Adult. Moths are about 5–7 mm long and bear silvergray scales interspersed with darker scales. The wingspan is 8–10 mm. The wings are fringed at the hind margins. The antennae are long and thin, with alternating light and dark segments. The legs similarly are banded. The abdominal scales are gray in males and cream colored in females. Adults mate repeatedly, with mating occurring at dawn. Adults can persist for about 30 days. Over the course of her life, the female can produce up to 260 eggs, though production of 60–120 eggs is more typical. Adults deposit eggs mostly on foliage, and stems. Occasionally eggs are deposited on green fruit, but not red fruit. These moths are not easily distinguished from tomato pinworm. On average, the forewings of tomato leafminer have more defined dark patches, whereas those of tomato pinworm have less distinct brownish streaks. Dissection of the male genitalia generally is required for accurate identification.

Damage The larvae most often attack the leaves and stems, but will also burrow into the fruit. The blotch-shaped mines in the leaves are diagnostic. The larvae extrude excrement (frass) from the mines through the entrance hole. Extensive mining can cause the leaves to drop from the plant. Damage to the fruit often results in entry of plant decay pathogens, and contamination of the fruit with larvae.

Management Sampling. Visual inspection can be used to detect these insects, especially examination for mines, though larvae and eggs are also detectable. Care must be taken, however, not to confuse the mines of tomato leafminer with tomato pinworm or one of the leafmining flies. Yellow sticky, McPhail, light, and pheromone-baited traps can be used to assess insect abundance. Also, the adults are active at dusk and dawn and can be flushed from the foliage and captured with a sweep net. Insecticides. Insecticide application is traditionally are the most commonly used insect management technique for this insect. Unfortunately, insecticide resistance has become a problem. Thus, minimal application frequency and insecticide rotation are recommended. Systemic insecticides are popular. At low pest population densities, organic insecticides such as neem, Bacillus thuringiensis, and Beauveria bassiana are reasonable options. Dustable sulfur, though not insecticidal, can be applied to foliage as an oviposition deterrent (Zappalá et al., 2012). Fruits that are moved from infested to uninfested areas are often fumigated as a precaution against moving the insects. Cultural Practices. Introduction into new fields or new locations often results from attachment of eggs or larvae to seedlings or to fruit, so sanitation should be a p­ riority.

Seedling transplants are a potential source of insects. Similarly, crop residue from a previous crop can provide the inoculum for a new infestation. Tomato cultivars vary in their susceptibility to T. absoluta (Sohrabi et al., 2017), though host plant resistance is not fully exploited with respect to this pest. In the greenhouse environment, insect exclusion nets should be fitted over vents, and doors should close tightly. Preferably, a double-door system should be used. Humans can be a good vector for the movement of insects, so care must be exercised so that workers do not transport moths. Mass trapping is feasible in closed environments such as greenhouses and can use color- or pheromone-based attractants as a basis for trapping. Crop rotation is desirable, particularly with nonhost crops such as cucurbits. Solanaceous weeds are another source of insects and so should be controlled. Pheromones. The pheromone of T. absoluta is highly attractive to males and can be used in conjunction with sticky, McPhail, or water pan traps of several designs to reduce the number of moths, especially in greenhouses. These traps can also be used in the field to assess the occurrence and abundance of the moths. Mating disruption by the continuous release of pheromone has been shown to be somewhat effective in greenhouses (Cocco et al., 2013).

Tomato Pinworm

Keiferia lycopersicella (Walsingham) (Lepidoptera: Gelichiidae)

Natural History Distribution. Tomato pinworm was first found in the United States in southern California in 1923. It now occurs regularly as a field pest in warm areas such as California, Arizona, Texas, and Florida. It also is known from Hawaii, the Caribbean, and Central and South America. Tomato pinworm commonly overwinters in greenhouses and may be shipped northward in the spring on seedlings cultured in warm areas, so the potential area of infestation is quite large. The origin of tomato pinworm is thought to be Central America. Host Plants. Tomato pinworm develops only on plants in the family Solanaceae. Tomato is undoubtedly the preferred host, but eggplant and potato also serve as suitable hosts (Schuster, 1989). The role of weeds in tomato pinworm biology is less certain, but among the weeds reported to support pinworm are bitter nightshade, Solanum dulcamara; black nightshade, S. nigrum; horsenettle, S. carolinense; and silverleaf nightshade, S. elaeagnifolium (Elmore and Howland, 1943; Batiste and Olson, 1973). Natural Enemies. Over 20 species of wasp parasitoids of tomato pinworm are known, including species in the families Braconidae, Ichneumonidae, Eulophidae, Pteromalidae,

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Benthylidae, and Trichogrammatidae (Oatman and Platner, 1989). In southern California, six species are common, but the dominant species there, and elsewhere, usually is Apanteles dignus Muesebeck (Hymenoptera: Braconidae). Other important parasitoids include Apanteles scutellaris Muesebeck, Chelonus phthorimaeae Gahan, Parahormius pallidipes (Ashmead) (all Hymenoptera: Braconidae), Sympiesis stigmatipennis Girault (Hymenoptera: Eulophidae); and Campoplex phthorimaeae (Cushman) (Hymenoptera: Ichneumonidae) (Oatman et  al., 1979). Oatman (1970), working in California, observed that parasitism of pinworm increased from practically undetectable in April to about 70% in June; six species contributed to pinworm suppression. Many of the parasitoids affecting tomato pinworm also are associated with potato tuberworm, Prthorimaea operculella (Zeller). In Florida, Pena and Waddill (1983) similarly observed an increase in parasitism rates as the season progressed, and also observed the egg parasitoid, Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae) to be important, particularly in fields with low pinworm densities. The importance of parasitism was also observed by Wang and Shipp (2004) who studied T. petiosum releases on greenhouse tomatoes. Life Cycle and Description. A generation can be completed in just 30 days under summer conditions. The number of generations is estimated at 7–8 annually in California, with 4–5 occurring during the summer months. The pinworm does not enter diapause, but development slows greatly during cool weather, with the pupal stage most important for survival during cool weather or the absence of hosts.

Larva. Upon hatching, young larvae spin a small web of silk on the leaf surface, then dig beneath the web and enter the leaf. There are four larval instars. The lower developmental threshold for larvae is about at 11°C. Mean head capsule width (range) is 0.15 (0.14–0.16), 0.25 (0.23–0.28), 0.37 (0.36–0.39), and 0.56 (0.52–0.61) mm, respectively, for instars 1–4. Young larvae are yellowish gray with a brown head capsule and measure about 0.85 mm long. As the larva matures it develops dorsally on the abdominal segments a darker pigmentation which is initially orangish or brownish, but eventually purplish. The dark region is irregular in shape but contains two small circular light spots, and two elongate light spots that connect to the light-colored background. The mature larva measures 5.8–7.9 mm long. The first two instars are leafminers, but they become too large for mining and third instars construct a leaf fold in which the last two instars dwell. Leaf mining pinworms deposit nearly all their fecal material in a single mass at the entrance to the mine. This characteristic is useful to distinguish pinworm mining from feeding by dipterous leafminers, Liriomyza spp., because the latter species deposit their feces throughout the mines. Pupa. Mature larvae drop to the soil on a strand of silk, spin a loosely woven pupal cell intermingled with soil particles, and pupate. Pupae are found at or near the soil surface, about 90% within the upper 1 cm of soil. Cool-weather is often passed in the pupal stage. The pupa is green when first formed but soon turns brown. The pupa measures 4–5 mm long. Duration of the pupal stage is normally 8–20 days.

Egg. The eggs of tomato pinworm are deposited on both the upper and lower surface of leaves in small clusters of 3–7. The small eggs are elliptical, though somewhat flattened where they attach to the substrate. They measure 0.30–0.45 mm long and 0.20–0.25 mm wide, and are very difficult to detect under field conditions. When first deposited they are light yellow, but gradually turn light orange. The lower developmental threshold for eggs is estimated at 11.4°C. Duration of the egg stage is 4–7 days.

FIG. 10.53  Tomato pinworm adult. (Photo by J. Capinera.)

FIG. 10.52  Tomato pinworm larva. (Photo by J. Capinera.)

Adult. The moth is grayish in general appearance, with a wingspan of 9–12 mm. The oval forewings, though mostly gray, are marked with diffuse orangish or brownish longitudinal spots or streaks. The hind wing is a more uniform yellowish brown, narrow in form, and pointed apically. The forewings, but especially the hind wings, are heavily fringed. Adults live only about 7–9  days and are nocturnal. Mating commences within 24 h of emergence, and

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most mating occurs shortly after sunset (McLaughlin et al., 1979). Most of the eggs are deposited within 3–4 days of adult emergence. The biology of tomato pinworm was described by Thomas (1936) and Elmore and Howland (1943). Thermal relations were given by Weinberg and Lange (1980) and Lin and Trumble (1985). Tomato pinworm is included in a key to vegetable insects by Sparks and Liu (2001). Rearing was described by Schuster and Burton (1982) and Burton and Schuster (1986).

Damage Young larvae mine the leaves of host plants, forming narrow serpentine or straight mines. Later, as larvae mature they leave the mines, fold the leaves, and dwell within the folds, causing large blotch-shaped mines adjacent to each leaf fold. Most important, however, is the tendency of larvae to enter fruit from the area of the stem. In the fruit they may feed shallowly beneath the skin of the fruit, causing blotches, or feed within the fruit, usually in the core. In general, there is a positive relationship between numbers of pinworm larvae and fruit damage, though it is much stronger in the fruit of the lower canopy of the plant than the upper region (Pena et al., 1986). In warm weather tomato-producing areas such as southern California and Florida, fruit can be severely injured, with infestation levels of 70%–100% recorded in the absence of cultural and chemical management of pinworm. In northern areas, such injury can also occur in greenhouse tomatoes.

Management Sampling. Wellik et al. (1979) suggested that the lower canopy stratum, particularly the large fruit, was sampled for pinworm. Pena et  al. (1986) also suggested sampling the lower canopy because of the strong relationship between foliar injury and yield loss in this stratum. They also indicated that population densities of less than one larva per plant could cause economic loss in Florida due to the high market value of winter tomatoes. However, Wolfenbarger et al. (1975) suggested that feeding damage on the top of the plant was related to tomato yield, and up to 0.99 feeding areas on the top three leaves was not a threat to the crop, whereas 3.3 or more feeding areas on these leaves was a serious problem. Pheromone traps can also be used to monitor populations (Wyman, 1979) and to time insecticide applications. van Steenwyk et  al. (1983), for example, recommended an insecticide application threshold of 10 moths per trap per night. Toscano et al. (1987), however, reported a relatively poor relationship between trap catches and larval populations. Sex pheromone can also be applied to crops to confuse the males and disrupt mating (van Steenwyk and Oatman, 1983).

Insecticides. Applications of insecticide are often made to foliage to prevent an increase in pinworm number and damage to fruit (Batiste et  al., 1970b; Schuster, 1982). Weisenborn et al. (1990) reported that weekly applications of insecticide made after a threshold of about 0.5 larvae per plant was attained provided protection of tomato in California. Application of granular systemic insecticides is often ineffective (Schuster, 1978), and use of the bacterium Bacillus thuringiensis is not recommended. Insecticide resistance has developed in many areas (Brewer et al., 1993). Cultural Practices. Sanitation is a very helpful practice. Tomato plants and cull tomatoes should be destroyed immediately after harvest or pinworm will continue to breed. If crops are grown sequentially, it is highly desirable to grow new crops at some distance from old sites. Infested tomato transplants moved from southern to northern areas also serve to transport pinworm into new areas or fields (Batiste et al., 1970a).

FAMILY HESPERIIDAE—SKIPPERS Bean Leafroller

Urbanus proteus (Linnaeus) (Lepidoptera: Hesperiidae)

Natural History Distribution. Bean leafroller is a tropical species but apparently is native to the southeastern United States. It is found throughout Florida and in coastal areas from South Carolina west to eastern Texas. It also invades most of the southeastern states, even attaining New England during warm years, and regularly invades the southernmost areas of the Southwest. It cannot tolerate prolonged freezing temperatures, however, and in the United States, it persists only in the southern coastal plain, perhaps only in southern Florida. The range of bean leafroller also includes Mexico, Central America and the Caribbean, and south to Argentina. A closely related species, Urbanus dorantes Stoll, has expanded its range from the tropical Americas to include the permanent range of U. proteus in the United States. Found throughout Florida, and southern Texas and Arizona, it is readily confused with bean leafroller, particularly because it also feeds on legumes, including bean (Heppner, 1975). Thus far, however, it is not particularly abundant. Host Plants. Larvae of bean leafroller feed exclusively on legumes and normally are found inhabiting open, disturbed habitats. Vegetable hosts include cowpea, lima bean, pea, and snap bean. Other hosts include soybean; wisteria, Wisteria sp.; tick trefoil, Desmodium spp.; butterfly pea, Clitoria spp.; and hog peanut, Amphicarpa bracteata. The adult form, a longtail skipper, feeds on the nectar from numerous flowers.

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Natural Enemies. Natural enemies are poorly documented. Wasp and fly parasitoids were observed in Colombia (van Dam and Wilde, 1977). Two tachinids with a very wide host range, Lespesia aletiae (Riley) and Nemorilla pyste (Walker), have been reported from bean leafroller (Arnaud Jr., 1978). In Florida, Chrysotachina alcedo (Loew) (Diptera: Tachinidae) was reared from larvae, and predation was observed by a Polistes sp. wasp (Hymenoptera: Vespidae) and Euthyrhynchus floridanus (Linnaeus) stink bugs (Hemiptera: Pentatomidae). Also, a nuclear polyhedrosis virus (NPV) was found to infect and to kill up to 40%–50% of larvae late in the season when larvae were numerous (Temerak et al., 1984). Life Cycle and Description. The bean leafroller can complete its life cycle in about 30  days during warm weather. They breed in southern Florida throughout the year, but are relatively infrequent in northern Florida until June, and only become abundant late in the season, usually September-October. The number of generations is not known but has been estimated at 3–4 in Florida. In the northern areas invaded during the summer months, usually, there is only a single generation. In Florida, large numbers of adults are frequently observed migrating southward in the autumn (Balciunas and Knopf, 1977); they make a similar northward migration in the spring and summer, but it is less apparent.

FIG. 10.54  Eggs of bean leafroller. (Photo by L. Buss.)

Egg. The eggs of bean leafroller may be deposited singly on the lower epidermis of foliage, but often are found in small clusters of 2–6 eggs. Also, clusters of up to 20 eggs have been observed, and on occasion, they may be stacked on one another to form a column. Initially the eggs are white, but soon turn yellow. The egg is a slightly flattened sphere and marked with about 12 vertical ridges. It measures about 1 mm in diameter and about 0.8 mm long. Eggs hatch in about 4.5 days when held at 24°C and 2.8 days at 29°C.

FIG. 10.55  Caterpillar of bean leafroller. (Photo by J. Capinera.)

Larva. The larva of bean leafroller increases in size from about 2 mm at hatching to 30 mm at maturity. Initially, the larva cuts a small triangular patch at the edge of the leaf, folds over the flap, and takes up residence within this shelter. The larva leaves the shelter to feed, and lines the shelter with silk. These flaps are used until the third or fourth instar when the larva constructs a larger shelter formed by folding over a large section of the leaf or webbing together two separate leaves. Larvae feed nocturnally. Eventually, the larva pupates within the leafy shelter. There are five instars. Mean head capsule widths (range) for the five instars are 0.6 (0.5–0.7), 1.1 (1.0–1.2), 1.8 (1.5–2.1), 3.2 (2.9–3.5), and 4.7 (4.2–5.2) mm, respectively. When maintained at 24°C the mean duration is about 2.8, 3.1, 3.5, 5.6, and 15 days, respectively, for instars 1–5, whereas instar duration at 29°C is about 2.0, 1.7, 2.2, 2.8, and 5.9 days. Initially, the larva is yellowish with a brownish-black head and prothoracic shield, and this general color pattern is maintained though the markings become more distinct as the larva matures, and the larva may also acquire considerable green color. With the molt to the second instar, the dorsal surface of the insect is marked with numerous small black spots. Beginning with the third instar, lateral yellow lines become quite distinct. The last two instars are similar to the preceding: brownish-black head, black prothoracic shield, yellowish body sprinkled with black spots but lighter below, and yellow lateral lines. Also evident are orange spots on the head near the base of the mandibles, and red on the ventral portion of the thoracic segment. The body tapers sharply toward both the anterior and posterior ends. Perhaps the most striking attribute of this insect, but a character shared with other members of the family Hesperiidae is the greatly enlarged head, which is connected to the body by a narrow “neck.” Pupa. The larva pupates on the plant within the shelter formed by the larva from leaf material. The pupa measures about 20 mm long and about 6 mm wide. The pupa

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is ­yellow to brown and is covered by a bluish-white pubescence. Duration of the pupal stage is about 17.1 day at 24°C and only 8.7 days at 29°C.

densities of 140 eggs or 70 first instar larvae per plant must occur to cause damage.

Management Sampling. The distribution of eggs and larvae is clumped (Shepard, 1972). Populations are normally sampled by visual observation because the larvae are sheltered within leaf folds and are difficult to dislodge by sweeping and because the leaf damage caused by shelter-building activity is readily apparent. Insecticides. Insecticides applied to the foliage at the first sign of damage are very effective for leafroller suppression. This should only be necessary for late-season bean crops. The microbial insecticide Bacillus thuringiensis is currently not recommended.

FIG. 10.56  Bean leafroller moth. (Photo by J. Capinera.)

Adult. The bean leafroller moth is fairly large for a hesperiid, measuring about 50 mm in wingspan. The most pronounced feature is the prolonged extensions or “tails” of the hind wings. Not surprisingly, the butterfly is commonly known as “long-tailed skipper.” The front and hind wings are chocolate brown dorsally and pale brown ventrally. The front wings are also marked with 5–7 squares or rectangular white spots. Green iridescent scales are found dorsally on the wings and body. The presence of green scales serves to differentiate bean leafroller from the other bean-feeding skipper, Urbanus dorantes. Skippers are active principally at dawn and dusk, often seen darting from flower to flowerseeking nectar. This insect has not been well studied. The most complete description was provided by Quaintance (1898a), but van Dam and Wilde (1977), Young (1985), Nava and Para (2002), Greeney and Sheldon (2008), and Cock (2015) provided valuable observations. Phenology and developmental biology were given by Greene (1970a, 1971a). Greene (1970b) made reference to culture on standard bean-based artificial diet, but the relative success of this approach is not clear.

Damage Larvae are defoliators, feeding only on leaf tissue of legumes. Greene (1971b) determined that larvae consumed about 0.5, 1.3, 5.1, 26.1, and 162.4 cm2 of foliage during instars 1–5, respectively. Beans can tolerate up to about 30% leaf loss without a reduction in yield, so Greene estimated that about 4.4 larvae must complete their development on a “typical” bean plant with 2175 cm2 of foliage to inflict a damaging level of defoliation. As about one-half of the individuals perish in each life stage, Greene estimated that

FAMILY LYCAENIDAE—HAIRSTREAK BUTTERFLIES Gray Hairstreak

Strymon melinus Hübner (Lepidoptera: Lycaenidae)

Natural History Distribution. This insect is found throughout the United States and southern Canada. Its range also extends south through Central America to South America. Apparently, it is absent from the Caribbean. Host Plants. The caterpillar stage of gray hairstreak feeds on the widest array of plants of any butterfly, but few are crop plants. Serious damage is limited to bean and cotton. Because of its injury to the latter plant, this species is also known as “cotton square borer.” Vegetable crops eaten include cowpea, lima bean, okra, pea, and snap bean. This insect is sometimes associated with field crops such as alfalfa, hops, lespedeza, and sweet clover. Numerous other plants are consumed by this insect, including Leguminocae such as Astragalus, Casia, Desmodium, Lupinus, and Trifolium spp.; Malvaceae such as Hibiscus, Malva, and Sphaeralcea spp.; Polygonaceae such as Eriogonum, Polygonum, and Rumex spp; and many others. Although the preferred habitat is in open areas inhabitated by early successional plants and shrubs, most agricultural weeds are exempt from attack by this insect. Scott (1986) provided a list of larval hosts. Adults collect nectar from goldenrod, Solidago spp.; dogbane, Apocynum spp.; milkweed, Asclepias spp.; sweet clover, Melilotus spp.; and other flowering plants (Opler and Krizek, 1984). Natural Enemies. Natural enemies are generally effective at suppression of gray hairstreak numbers, so little crop damage occurs. In Texas, for example, the first generation is abundant, but in subsequent generations caterpillars tend to be less and less abundant as parasitoids take a toll. The most

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important parasitoid is a gregarious species, Apanteles thecloe Riley (Hymenoptera: Braconidae). This species attacks the caterpillar, with larvae emerging from the host to pupate in white silken cocoons on the back of the caterpillar. Other, less important parasitoids are Octosmicra sp. (Hymenoptera: Chalcididae), Aplomya theclarum (Scudder), and Lespesia sp. (both Diptera: Tachinidae). Predators are poorly known, but adults are taken by robber flies (Diptera: Asilidae). Life Cycle and Description. There are two generations annually in the northern portion of gray hairstreak’s range, and three to four generations in the central and southern states. About 40 days are required for a complete generation. In Texas, butterflies are observed in all months except December and January. The winter is passed in the pupal stage. Females begin oviposition in February or March in the south, but not until May in the North.

Larvae normally feed on the leaf surface during the first three instars and thereafter display a tendency to bore into pods or other reproductive tissues. Pupa. Near the completion of larval development, the larva attaches itself with silk to the plant, usually adjacent to the feeding site. After a quiescent period of 2–3 days the larva molts into the pupa. The pupa is yellow to brown, and bear blackish spots that vary in size, frequency, and intensity. It measures 7.5–10.5 mm long. The sides of the pupa bear spines arranged regularly in rows. Duration of the pupal stage is about 9.6 days (range 8–11 days).

Egg. The eggs are spherical in shape but slightly flattened. They are pale-green to yellowish white. They measure 0.6–0.7 mm in diameter, and 0.2–0.3 mm long. They are deposited singly, usually on the underside of leaves. Caged adults have been observed to produce 40–70 eggs each, but this is believed to be an underestimate of true fecundity because this species fares poorly in captivity. Oviposition occurs most frequently in the morning and early evening. Mean duration of the egg stage is 5.5 days (range 4–6 days). FIG. 10.58  Adult of gray hairstreak butterfly. (Photo by J. Capinera.)

FIG. 10.57  Larva of gray hairstreak butterfly. (Photo by J. Castner.)

Larva. There are five instars. The mean (range) duration of each instar is 3.6 (3–5), 3.4 (3–5), 3.5 (3–6), 4.8 (3–6), and 7.7 (5–11) days, respectively. Thus, the total larval development requires about 23 days. At hatch, larvae measure only about 1 mm long, but eventually grow to measure 12– 16 mm long and 4.5–6.5 mm wide. Initially, the larvae are slender, but with each succeeding molt the larva becomes relatively broader, so at maturity it is rather stout. The caterpillar in all instars is covered with stout hairs or bristles. The body color is generally green, but sometimes brownish.

Adult. The adults usually emerge in the morning, and 2–3 days are required before oviposition commences. The butterflies are small, measuring only 25–32 mm in wingspan. In general appearance they are blue gray. The dorsal surface of the wing is steel gray, with the lower surface pale gray. The hind wing is marked dorsally with a reddishorange spot. The wings are fringed with long scales, and each hind wing bears two thin extensions or tails—one long and the other short. Adults live at least 10 days in confinement but likely live considerably longer in nature. In southern latitudes, butterflies apparently are dormant during the brief cold periods of winter and become active during warm weather. Adult males perch on bushes and trees and patrol their territory to watch for intruders. They are aggressive about defending their territory not only from males of the same species, but other butterflies, wasps, and even hummingbirds (Alcock and O’Neill, 1986). A comprehensive treatment of gray hairstreak biology was given by Reinhard (1929).

Damage Foliage consumption by early instar larvae is insignificant, but the damage may occur as larvae burrow into bean or

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okra pods. On beans, larvae often burrow only partially into the pod, then move to another location, which may be on the same or another pod. This damage is likely to be significant only in home gardens.

Management These insects cause only incidental damage and infrequently require suppression. Foliar insecticides are generally very effective.

FAMILY NOCTUIDAE—ARMYWORMS, CUTWORMS, LOOPERS, STALK BORERS, AND NOCTUID MOTHS Alfalfa Looper

Autographa californica (Speyer) (Lepidoptera: Noctuidae)

present were Microplitis spp. and Apanteles yakutatensis Ashmead (Hymenoptera: Braconidae), and Campoletes sonorensis (Cameron) and Patroclides montanus (Cresson) (Hymenoptera: Ichneumonidae). Other parasitoids, of uncertain importance, were noted by Hyslop (1912a). A NPV is very common in alfalfa looper populations, usually killing the larvae as they reach maturity. Life Cycle and Description. The number of generations per year is reported to be two in Alberta and two to three in Washington. A complete life cycle is estimated to require 30–40 days. Overwintering occurs in the pupal stage. Miller et al. (1984) reported that the developmental threshold was 11.5°C and about 360 degree-days are required for complete development (oviposition to adult emergence). Egg. The eggs are deposited singly, usually on the underside of the foliage. They are pale yellow, hemispherical in shape, and marked with narrow vertical ridges. The eggs normally hatch in 3–5 days.

Natural History Distribution. Alfalfa looper is native to North America, and is western in distribution. In the United States it occurs occasionally in Nebraska and Kansas, and frequently in all states further west. It is most damaging along the West Coast. In Canada its distribution is similar, and is considered a pest only in British Columbia and Alberta. Alfalfa looper is also known from Mexico. Host Plants. This insect feeds on numerous plants. Vegetable crops reportedly injured include beet, bean, cabbage, carrot, cantaloupe, celery, cucumber, lettuce, onion, pea, potato, radish, rhubarb, spinach, squash, tomato, turnip, watermelon, and probably others. The most frequently damaged vegetables are lettuce, bean, and the crucifer crops. Field crops damaged include alfalfa, cotton, red clover, sweet clover, flax, white clover, sugarbeet, and sunflower; as its common name suggests, alfalfa is the most common host. Grasses and grains are not eaten, with the exception of rare feeding on corn. Fruit crops such as apple, currant, gooseberry, raspberry may be damaged. Many of the hosts recorded in the literature are based on outbreak conditions, when hordes of larvae, after totally consuming their preferred hosts, disperse in search of food. Under such conditions, relatively unpreferred plants such as the aforementioned fruit crops may be eaten, but this is not the normal situation. Numerous common weeds support larval development, including dock, Rumex crispus; lambsquarters, Chenopodium album; wild lettuce, Lactuca canadensis; and many others. Natural Enemies. Mortality due to natural enemies was estimated at 70%–80% in California (Puttarudriah, 1953), of which parasitoids accounted for 30%–35% and the remaining from undetermined diseases. The principal parasitoid was Voria ruralis Fallen (Diptera: Tachinidae). Also

FIG. 10.59  Larva of alfalfa looper caterpillar. (Photo by J. Capinera.)

Larva. Larvae have only three pairs of abdominal prolegs, and greatly resemble cabbage looper, Trichoplusia ni (Hübner). The body is broader at the posterior end and narrower at the anterior end. As is the case with cabbage looper, the mature larva is predominantly green but is usually marked with a distinct white stripe on each side. Dorsally, the larva bears several narrow, faint white stripes clustered into two broad white bands. When larvae hatch they measure less than 2 mm long, but at the completion of the five instars they attain a length of about 25–35 mm. At cooler temperatures, six instars may occur. Unlike the cabbage looper, the alfalfa looper tends to have dark thoracic legs and a dark bar on the side of the head, or even an entirely black head capsule. Also, alfalfa looper lacks the small, n­ ipple-like structures located ventrally on the third and fourth abdominal segments of cabbage looper. Alfalfa looper can be distinguished from bilobed looper, Megalographa biloba (Stephens), and celery looper, Anagrapha falcifera (Kirby) by the absence of numerous dark microspines on the body

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of alfalfa looper. A key to common vegetable-feeding loopers can be found in Appendix A and in Capinera (1986). Duration of the larval stage is about 14 days. Pupa. The mature larva spins a loose, whitish silk cocoon, often incorporating leaves into the structure, and pupates within. The pupa is 18–20 mm long, and its color is blackish brown. Duration of the pupal stage is about 10–14 days.

Management Alfalfa looper infrequently is a vegetable pest, though in the southwest it sometimes damages lettuce, especially during spring in California (Vail et al., 1989). Populations of moths, including both males and females, can be monitored using traps baited with floral chemical lures (Landolt et al., 2001). Foliar application of Bacillus thuringiensis or chemical insecticides is effective. Careful monitoring is recommended, especially if late-season lettuce is grown in proximity to alfalfa or cotton. Lettuce is sensitive to injury, and an average of 0.25–0.5 larvae per plant likely warrants suppression. Components of the sex pheromone are known and can be used to bait traps for population monitoring (Steck et  al., 1979a, b). Moths also can be captured in blacklight traps.

Army Cutworm

Euxoa auxiliaris (Grote) (Lepidoptera: Noctuidae)

Natural History FIG. 10.60  Adult of alfalfa looper caterpillar. (Photo by J. Capinera.)

Adult. The moth measures 35–45 mm in wingspan. The front wings are gray with irregular light-brown and darkbrown patches. Near the center of each forewing, there is a prominent silvery white or yellow-colored mark, which in general shape is said to resemble the hind leg of a dog. The hind wing is light brown or gray basally, and dark brown distally. The edge of both front and hind wings is marked with a series of dark spots. Unfortunately, the color pattern of the moths is not completely diagnostic, as there are other species with similar markings. LaFontaine and Poole (1991) should be consulted for keys to moths. The biology of alfalfa looper is not well cataloged. Hyslop (1912a) and Parker (1915) provided accounts of its ecology. Alfalfa looper is easily reared on bean-based artificial diet (Shorey and Hale, 1965). The larvae are included in the keys of Crumb (1956), Okumura (1962), Capinera (1986), and Stehr (1987). Adults are included in the keys of Eichlin and Cunningham (1978), Oliver and Chapin (1981), and Capinera and Schaefer (1983).

Damage Larvae of alfalfa looper are defoliators. Initially, young larvae may be gregarious and skeletonize leaves. They soon disperse, however, and chew irregular holes in foliage. They feed on the underside of leaves. A sign of their presence is large quantities of wet fecal matter adhering to foliage. When larvae are abundant they may disperse in large numbers from favored to less-favored plants. Alfalfa is often the source of such infestations.

Distribution. This native insect is abundant in the Great Plains and Rocky Mountain regions in the United States and Canada. It has been recorded from all states west of the Mississippi River, and as far east in Canada as Ontario, but it attains high densities only in semiarid areas. Host Plants. Army cutworm has been reported to feed on numerous plants. It is known principally as a pest of small grains, perhaps because these crops dominate the landscape where army cutworm occurs. Among vegetable crops, it has been reported to damage beet, cabbage, celery, corn, onion, pea, potato, radish, rhubarb, tomato, and turnip. Other crops injured include fruit crops such as apple, apricot, blackberry, cherry, currant, gooseberry, peach, plum, prune, raspberry, and strawberry, and field crops such as alfalfa, barley, clover, flax, rye, sanfoin, sunflower, sweet clover, timothy, vetch, and wheat. Army cutworm also feeds on noncultivated plants such as bluegrass, Poa spp.; bromegrass, Bromus spp.; buffalograss, Buchloe dactyloides; gramagrasses, Bouteloua spp.; field pennycress, Thlaspi arvense; dandelion, Taraxacum officinale; lambsquarters, Chenopodium album; and lupine, Lupinus spp. Natural Enemies. Many natural enemies have been found associated with army cutworm, and both hymenopterous parasitoids and disease have been documented to cause considerable mortality. Walkden (1950), working in the central Great Plains, reported mortality trends over a 20-year period and observed parasitism levels of up to 33% and disease incidence of up to 57%. Not surprisingly, the incidence of disease was greatest at high armyworm population densities. Snow (1925) reported 30% parasitism in Utah. In a 3-year study in Oklahoma, researchers found that less than 12% of larvae were parasitized, with most parasitism due to two species, Meteorus leviventris (Wesmael)

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and Apanteles griffini Viereck (both Hymenoptera: Braconidae) (Soteres et al., 1984). A polyembryonic wasp, Copidosoma bakeri (Howard) (Hymenoptera: Encyrtidae), causes larvae to consume more food, to become larger, and live longer; this can result in the appearance of artificially high rates of parasitism, which sometimes exceeds 50% (Byers et al., 1993). Among the other parasitoids known from army cutworm are wasps such as Cotesia marginiventris (Cresson), A. militaris Walsh, Chelonus insularis Cresson, Macrocentrus incompletus Muesebeck, Microplitis feltiae Muesebeck, M. melianae Viereck, Rogas sp., Zele melea (Cresson) (all Hymenoptera: Braconidae); Campoletis flavicincta (Ashmead), C. sonorensis (Cameron), Diphyus nuncius (Cresson), Exetastes lasius Cushman, and Spilichneumon superbus (Provancher) (all Hymenoptera: Ichneummonidae). Flies known to parasitize these species include Bonnetia comta (Fallen), Euphorocera claripennis (Macquart), Mericia spp., Peleteria sp., Periscepsia cinerosa (Coquillett), P. helymus (Walker), and P. laevigata (Wulp) (all Diptera: Tachinidae). Several viruses are known to infect army cutworm, including entomopox, granulosis, and nonoccluded viruses (Jackson and Sutter, 1985; McCarthy et  al., 1975; Sutter, 1972, 1973). The relative importance of each is uncertain, but the granulosis virus is unusually pathogenic. Life Cycle and Description. There is a single generation per year throughout the range of this insect. The eggs are deposited on soil in August-October. They hatch in autumn or early winter, and larvae overwinter, feeding actively in the spring. Pupation occurs about a month before adults appear. Adults first become active in April-May in southern locations such as Kansas and Texas, whereas in more northern locations such as Alberta and Montana they may not appear until June-July. The moths migrate from the plains, where the larvae develop, to higher elevations in the Rocky Mountains, where the adults feed on nectar from flowers. The adults return to the plains in September-October. Egg. The eggs are deposited singly or in small clusters just beneath the soil surface on a solid substrate (Pruess, 1961). Soil particles adhere to the eggs so they are difficult to detect in the field. In shape, the eggs are a slightly flattened sphere, measuring about 0.6 mm in diameter and 0.5 mm in height. The egg is white to yellow initially, becomes gray to brown as the embryo matures. The egg is marked with about 18 very narrow ridges that radiate from the apex. Survival of eggs apparently is affected by moisture, and Seamans (1928) suggested that above-average rainfall in late summer and autumn assured good insect survival and damaging populations in the subsequent year; this concept appears not to have been independently confirmed, however. Field-collected females were reported by Pruess (1963) to produce 200–300 eggs, with the potential to produce about

500 eggs. However, Jacobson and Blakeley (1959) suggested that 1000–2500 eggs could be produced by a female based on laboratory studies in which larvae were fed dandelion, a highly suitable host.

FIG. 10.61  Army cutworm caterpillar. (Photo by J. Capinera.)

Larva. The eggs hatch in the autumn or early winter but the larvae are usually not noticed until spring when they increase in size and begin to consume considerable foliage. There are 6–7 instars, with head capsule widths of 0.26–0.30, 0.40–0.45, 0.65–0.72, 1.04–1.21, 1.70–2.10, and 2.90–3.40, mm, respectively, for instars 1–6 among larvae with only six instars (Jacobson and Blakeley, 1959). In comparison, head capsule widths of 0.25–0.30, 0.36–0.43, 0.55–0.70, 0.88–1.28, 1.40–1.90, 1.95–2.50, and 2.95–3.55 mm were reported for instars 1–7, respectively, in larvae with seven instars (Sutter and Miller, 1972). Additional instars apparently occur when larvae feed on less suitable host plants. Duration of the instars is estimated at 16–48, 13–73, 4–70, 3–42, 6–11, 4–18, and 9–25 days for instars 1–7, respectively (Burton et al., 1980). The body color of the larvae is grayish brown, but bears numerous white and dark brown spots. There is usually evidence of three weak light-colored dorsal stripes. Laterally, it tends to be a broad dark band and the area beneath the spiracles is whitish. The head is light brown with dark spots. Larvae attain a length of about 40 mm. They are usually found beneath the surface of the soil, emerging in the late afternoon or early evening to feed. On cloudy days, however, they may be active during the daylight hours. Larvae assume a migratory habit when faced with food shortage, and numerous larvae proceed in the same direction, consuming virtually all vegetation in their path. It is this dispersive behavior that is the basis for their common name, and larvae are observed to disperse over 4 km. Pupa. Pupation occurs in the soil, in a cell prepared by the larva. The walls of the cell are formed with salivary secretion, which hardens and provide a degree of rigidity. The depth of pupation varies according to soil and moisture

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conditions, but it may be any depth up to 7.5 cm. The larva spends about 10 days in the cell before pupation. Duration of pupation is 25–60  days. The pupa is dark brown, and measures about 17–22 mm long and 6 mm wide.

Damage These insects principally are pests of small grain crops grown in arid regions, though many irrigated crops are also at risk. Larvae readily climb plants to consume foliage, eating holes in vegetation initially, and eventually destroying the entire plant. Although they burrow into the soil during the daylight hours, they do not normally feed below-ground. However, when succulent food is in short supply they follow the plant stem down into the soil. When food supplies are exhausted numerous larvae may disperse in search of additional food.

Management

FIG. 10.62  Army cutworm moth. (Photo by J. Capinera.)

Adult. The adults measure 35–50 mm in wingspan. They are quite variable in appearance, with five named subspecies (Pruess, 1967), but moths generally assume two basic forms. One common form has the leading edge of the forewing marked with a broad yellowish stripe, and the remainder of the wing blackish but marked with white-rimmed bean-shaped and round spots, and a light transverse line. In another common color form, the forewing is mottled brown, bearing bean-shaped and round spots but lacking bands and stripes. In all cases, the hind wings are brownish with dark veins, and darker distally. The brown body of the moth is quite hairy. As earlier noted, the adults are migratory, dispersing from the plains to the mountains annually (Pruess, 1967; Pruess and Pruess, 1971). In transit and in the mountains they feed on nectar from flowering plants (Kendall et  al., 1981). They are nocturnal and seek shelter during the daylight hours. They have the habit of aggregating in houses, automobiles, and other sheltered locations where they become a nuisance, soil walls, and induce allergic reactions among some individuals (Storms et  al., 1981). They may also aggregate in natural shelters in mountainous regions, where they become prey for bears (Chapman et al., 1955; Mattson et al., 1991). In the Rocky Mountain region, they are commonly called “miller moths.” An excellent summary of army cutworm biology was given by Burton et al. (1980). Rearing procedures using vegetation were provided by Blakeley et al. (1958) and using an artificial diet by Sutter and Miller (1972). Sex pheromones have been identified (Struble and Swailes, 1977b; Struble, 1981b). Larvae are included in keys by Whelan (1935), Walkden (1950), Okumura (1962), Capinera (1986), and Stehr (1987), and are included in a key to armyworms and cutworms in Appendix A. Moths are included in pictorial keys of Rings (1977a), and Capinera and Schaefer (1983).

Sampling. Adults can be captured in light and pheromone traps. However, males are attracted to the sex pheromone only during the autumn flight. Pheromone traps positioned at a height of 1 m or lower are more effective than those placed higher (Swailes and Struble, 1979). Larvae can be recovered from soil by raking through the top 5–7 cm. Insecticides. Persistent insecticides can be applied to vegetation to kill army cutworm larvae when they emerge from the soil to feed; Bacillus thuringiensis is not effective (McDonald, 1979; Bauernfeind and Wilde, 1993) though pyrethroids are used successfully. Larvae also accept bran bait containing insecticide. Cultural Practices. Cultural manipulations are not generally effective to prevent oviposition because moths deposit eggs on barren soil. Delayed planting of crops can be effective, however, as larvae complete their development on weeds or starve before crops are planted. If larvae are dispersing, creation of deep ditches with steep sides, or filled with running irrigation water, may prevent the invasion of fields. To protect plants grown in the home garden, barriers are occasionally used to decrease access by cutworms to seedlings. Metal- or waxed-paper containers with both the top and bottom removed can be placed around the plant stem to deter consumption. Aluminum foil can be wrapped around the stem to achieve a similar effect. Because larvae burrow and feed below the soil line, the barrier should be extended below the soil surface.

Armyworm

Mythimna unipuncta (Haworth) (Lepidoptera: Noctuidae)

Natural History Distribution. Armyworm is a native species occurring throughout North America. It is most common, however, in the eastern United States and Canada west to the Rocky Mountains. It does not overwinter at northern latitudes such as Canada and the northern states. Rather, armyworm disperses northward each spring, principally along the

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Mississippi River Valley, and then disperses southward during the autumn, principally along the east coast (McNeil, 1987). It is often called “true armyworm” to distinguish it from other armyworms such as fall armyworm, Spodoptera frugiperda (J.E. Smith), and yellow-striped armyworm, Spodoptera ornithogalli (Guenée). Armyworm also occurs in Central and South America, southern Europe, central Africa, and western Asia. Host Plants. Armyworm generally prefers to oviposit and feed upon plants in the family Gramineae, including weedy grasses. Thus, such grain and grass crops as barley, corn, millet, oats, rice, rye, sorghum, sugarcane, timothy, and wheat may be consumed, as well as wild or weed grasses. During periods of abundance, larvae feed more generally, damaging vegetables such as artichoke, bean, cabbage, carrot, corn, celery, cucumber, lettuce, onion, parsley, parsnip, pea, pepper, radish, sweet potato, watermelon, and others. Field crops damaged during such population outbreaks include alfalfa, dry bean, and sugarbeet in addition to the aforementioned grain crops. Adults feed on the nectar of various flowers and sometimes feed on other sweet foods such as ripe and decaying fruit. Natural Enemies. The importance of natural enemies, especially parasitoids, has been studied, though nearly all data are derived from the periods of high armyworm density, which is not typical for this insect. Krombein et al. (1979) listed 35 species of Hymenoptera reared from armyworm, of which 19 species are braconids and 12 species are ichneumonids. Similarly, Arnaud Jr. (1978) listed 35 species of tachinids from armyworm. Breeland (1958) studied armyworm in Tennessee and gave a long list of natural enemies known to affect armyworm around the world. He reported rates of parasitism to be 30%–40% in Tennessee studies. The most important parasitoids found by Breeland during 1956 were Glyptapanteles militaris (Walsh) and Rogas terminalis (Cresson) (both Hymenoptera: Braconidae), which accounted for 27% and 5% of total parasitism, respectively; Winthemia rufopicta (Bigot) (Diptera: Tachinidae), which accounted for 12% parasitism; and Eniscospilus merdarius Gravenhorst (Hymenoptera: Ichneumonidae), which accounted for 9% parasitism. In contrast, during 1957, W. rufopicta accounted for only 1.5% parasitism; parasitism by A. militaris and R. terminalis increased to 36% and 22%, respectively; and parasitism by E. merdarius dropped to 4.5%. Interestingly, Hyposoter sp. (Hymenoptera: Ichneumonidae) was not observed in 1956 but accounted for nearly 20% of total parasitism in 1957. Guppy and Miller (1970) provided keys to the immature stages of armyworm parasitoids. Predators readily consume armyworm larvae. Ground beetles (Coleoptera: Carabidae) are especially effective because larvae spend most of their time in association with soil, but various predatory bugs (Hemiptera: various families), ants (Hymenoptera: Formicidae), and spiders (Araneae:

Lycosidae and Phalangiidae) also feed on armyworm (Clark et  al., 1994). Avian predators are often credited with the destruction of armyworms. The bobolink, Dolichonyx oryzivorus (Linnaeus), prospers during outbreak years and has sometimes been called the “armyworm bird.” Other birds of note include the crow, Corvus brachyrhynchos Brehm, and starling, Sturnus vulgaris Linnaeus. Diseases commonly infect armyworms, especially during periods of high density. Bacteria and fungi, particularly the fungus Metarhizium anisopliae, are reported in the literature. In Arkansas, Steinkraus et al. (1993a) reported an epizootic caused by the fungus Furia virescens, and a significant incidence of mermithid nematodes, with mortality by the nematodes estimated at 13.5%. Kramer (1965) documented the pathogenicity of two additional fungi, Thelohania diazoma (J.P. Kramer) and Vairimorpha (Nosema) necatrix (J.P. Kramer), to this insect. However, undoubtedly the most important diseases are viruses; several granulosis, cytoplasmic polyhedrosis, and NPVs often kill virtually all armyworms during periods of an outbreak, especially when larvae are also stressed by lack of food or inclement weather (Tanada, 1959, 1961). Weather. Armyworm attains high densities irregularly, often at 5- to 20-year intervals. The exact cause is unknown, but outbreaks often occur during unusually wet years and are preceded by unusually dry years. Armyworm is not well-adapted for hot temperature; survival decreases markedly when temperatures exceed about 30°C. Consequently, at southern latitudes, populations are higher early and late in the year, but at northern latitudes, it is a mid-season pest. Life Cycle and Description. Larvae apparently overwinter at least as far north as Tennessee, though they are unsuccessful in the northernmost states and in Canada, which are invaded annually by moths dispersing northward. In the south, all stages may be found during the winter months (Moran and Lyle, 1940). The number of generations varies among locations, but two generations occur in Ontario, two to three occur in Minnesota and New York, four to five are reported in Tennessee, and five to six in southern states (Knutson, 1944; Frost, 1955; Breeland, 1958; Guppy, 1961; Chapman and Lienk, 1981). In Tennessee, moth flights are observed in March-May, June, July-mid-August, September, and November. In New York, moths are common from March to September, but sometimes as late as November. A complete generation requires 30–50 days. Egg. Females deposit eggs in clusters consisting of two to five rows, in sheltered places on foliage, often between the leaf sheath and blade, especially on dry grass. Often females seem to deposit large numbers in the same vicinity, resulting in very high densities of larvae in relatively small areas of a field. Nevertheless, the eggs are very difficult to locate in the field. The eggs are white or yellowish but turn gray immediately before hatching. They are spherical and

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measure about 0.54 mm (range 0.4–0.7 mm) in diameter. The egg surface appears to be shiny and smooth, but under high magnification fine ridges can be observed. Their clutches are covered with an adhesive secretion that is opaque when wet but transparent when dry. As the adhesive material dries it tends to draw together the foliage, almost completely hiding the eggs. Mean duration of the egg stage is about 3.5  days at 23°C, and 6.5  days at 18°C, but the range is 3–24  days over the course of a season. Hatching rates are affected by temperature, with cool weather more favorable for embryonic survival. In Tennessee, about 98% egg hatch occurs in early spring and autumn, with hatching rates dropping to less than 30% during the summer; this probably accounts for the evolution of the dispersal behavior in this species.

readily extrude silk and spin down to the soil. Larvae in instars 3–6 are active at night, seeking shelter during the day on the soil beneath debris or clods of soil. Pupa. Larvae pupate in the soil, often under debris, at depths of 2–5 cm. Pupation occurs in an oval cell that contains a thin silken case. The pupa is moderate in size and robust, measuring 13–17 mm long and 5–6 mm wide. The pupa is yellowish brown initially but soon assumes a mahogany brown color. The tip of the abdomen bears a pair of hooks. Duration of the pupal stage is 7–14 days during summer but longer early and late in the season, sometimes lasting 40 days.

FIG. 10.63  Armyworm caterpillar. (Photo by J. Capinera.) FIG. 10.64  Armyworm moth. (Photo by J. Capinera.)

Larva. Larvae normally display six instars, though up to nine instars have been observed. Mean head capsule widths (range) are 0.34 (0.30–0.37), 0.55 (0.49–0.63), 0.94 (0.83– 1.12), 1.5 (1.29–1.70), 2.3 (2.08–2.56), and 3.3 (3.04–3.68) mm, respectively, for instars 1–6. Head capsule widths increase slightly with increased temperature up to about 30°C (Guppy, 1969). Larvae attain a body length 4, 6, 10, 15, 20, and 35 mm, respectively, during instars 1–6. Except for the first instar, which is pale with a dark head, the larvae of armyworm are marked with longitudinal stripes throughout their development. The head capsule is yellowish or yellow brown with dark net-like markings. The body color is normally grayish green but a broad dark stripe occurs dorsally and along each side. A light subspiracular stripe is often found laterally beneath the dark stripe. Development time varies with temperature. During summer, larvae complete their development in about 20 days, but this is extended to about 30  days during the spring and autumn, and greatly prolonged during winter. Instar-specific development times recorded during early summer in Tennessee are 2–3, 2–3, 2–4, 2–3, 4–5, and 7–10 days for instars 1–6, respectively. The larvae tend to disperse upward following hatching, where they feed on tender leaf tissue. If disturbed, they

Adult. The adult is a light reddish-brown moth, with a wingspan measuring about 4 cm. The forewing is fairly pointed, appearing more so because a transverse line of small black spots terminates in a black line at the anterior wing tip. The forewing is also marked with a diffuse dark area centrally containing 1–2 small white spots. The hind wings are grayish and lighter basally. Adults are nocturnal. Mating commences 1–3 days after moths emerge from the soil, and usually 4–7 h after sunset. Eggs are normally deposited within a 4–5 day period (range 1–10 days). Females produce an average of 4.9 egg masses (range 1–16 masses). Breeland (1958) reported that about 450 eggs are produced by each female (range 15–1350 eggs) in Tennessee, but Guppy (1961) found a mean fecundity of 1450 (range 250– 1900). Feeding is necessary for normal oviposition. Mean longevity at warm temperature is about 9 days in males and 10 days in females (range 3–25 days) whereas at cool temperature mean longevity of males is 19  days and females 17 days. An excellent treatment of biology was given by Breeland (1958). Other informative publications include Davis and Satterthwait (1916b), Walton and Packard (1947),

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Pond (1960), and Guppy (1961). Developmental biology was presented by Guppy (1969). A sex pheromone is described by Steck et  al. (1982) and female response by Turgeon et al. (1983). Armyworm is included in the larval keys of Walkden (1950), Crumb (1956), Rings and Musick (1976), Oliver and Chapin (1981), Capinera (1986), Stehr (1987), and Sparks and Liu (2001). Larvae are also included in a key to armyworms and cutworms in Appendix A. Moths are included in the keys by Rings (1977a) and Capinera and Schaefer (1983), and pictured in Rockburne and Lafontaine (1976) and Chapman and Lienk (1981).

Damage Larvae initially skeletonize foliage, but by the third instar, they eat holes in leaves and soon afterwards consume entire leaves. Larvae of armyworm are notorious for appearing out of nowhere to inflict a high level of defoliation. This occurs for several reasons: a highly clumped distribution of young larvae, with most of the crop uninfested until larvae are nearly mature and highly mobile; a tendency by larvae to feed on grass weeds preferentially, only moving to crops after the grass is exhausted; occurrence of a preponderance of feeding, about 80%, in the last instar; the nocturnal behavior of larvae, which makes them difficult to observe during the day; and the gregarious and mobile behavior of mature larvae, which form large aggregations or bands (hence the common name “army” worm). As earlier noted, grasses and grains are preferred, but as these plants are consumed larvae disperse, often in large groups, to other plants. During outbreaks, few plants escape damage.

Management Sampling. Adults can be captured with blacklight traps. A sex pheromone has been identified and can be used for population monitoring (Kamm et al., 1982; Lopez Jr. et al., 1990). Light-colored pheromone traps capture more moths than dark-colored traps (Hendrix III and Showers, 1990). It is advisable to examine crop fields for larvae, especially if moths have been captured in light or pheromone traps. Fields should be examined at dawn or dusk because larvae are active at this time. If it is necessary to check fields during the day, it is important to sift through the upper surface of the soil and under debris for resting larvae. Insecticides. Larvae consume wheat bran or apple pomace baits treated with insecticide, but foliar and soil-applied insecticides are also effective, and used frequently (Musick and Suttle, 1973; Harris et al., 1975a; Harrison et al., 1980a). Cultural Techniques. Cultural practices have limited effect on armyworm abundance due to their highly dispersive behavior. However, grass weeds are a focal point of infestation and should be eliminated, if possible. Not surprisingly, no-till and minimum tillage fields experience

greater problems with armyworm than conventional tillage fields (Harrison et  al., 1980a; Willson and Eisley, 1992). Proximity to small grain crops is considered to be a hazard owing to the preference of moths for such crops, and the suitability of grains for larval development. In Virginia, destruction of winter cover crops by herbicide application is more favorable to armyworm survival than mowing of cover crops, apparently because predators are more disrupted by herbicide treatment (Laub and Luna, 1992). Before the availability of effective insecticides, deep furrows with steep sides were sometimes plowed around fields to prevent invasion by dispersing armyworm larvae. Although this approach remains somewhat useful, it is rarely practiced. Biological Control. Some suppression of armyworm can be achieved with Bacillus thuringiensis, though it is not as effective as with some other caterpillars. Larvae are also susceptible to infection by the entomopathogenic nematode Steinernema carpocapsae, though neonate larvae are fairly resistant (Kaya, 1985).

Bean Leafskeletonizer

Autoplusia egena (Guenée) (Lepidoptera: Noctuidae)

Natural History Distribution. This insect is a native to tropical areas of the western hemisphere. The northern limits of its range are the Gulf Coast states and California. Its distribution extends southward through Central and South America to Brazil, and the Caribbean Islands. Host Plants. As suggested by its common name, the principal host is bean. However, other vegetables sometimes consumed include cabbage, carrot, and celery. Soybean, tobacco, mint, spearmint, comfrey, chrysanthemum, marigold, hollyhock, verbena, and larkspur are other economic plants eaten. Natural Enemies. Several parasitoids were found to affect bean leafskeletonizer in California; Copidosoma truncatellum (Dalman) (Hymenoptera: Encyrtidae) was the only parasitoid affecting a significant proportion of the leafskeletonizers (Lange, 1945), although Cotesia marginiventris (Cresson) and Apanteles yakutatensis (Ashmead) (both Hymenoptera: Braconidae) and Voria ruralis (Fallén) (Diptera: Tachinidae) are also known to parasitize bean leafskeletonizer (Clancy, 1969). In Florida, Meteorus autographae Muesbeck and Apanteles autographae Muesbeck (both Braconidae) as well as C. truncatellum were important parasites (Genung, 1960). An undetermined fungal disease was also noted to affect the larvae. There has not been adequate study to determine the relative importance of the natural enemies, but based on observation they are important mortality factors.

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Life Cycle and Description. The complete life cycle requires about 45 days. The number of generations appears to be undetermined, but Lange (1945) reported “several” in California. Egg and larva. The duration of the egg is about 5  days. Eggs are laid in small clusters of 2–3 eggs. In Puerto Rico, Casanova (1977) determined that larvae have five instars, the duration of which were 4.4, 3.4, 4.0, 4.1, and 7.8 days, respectively. Mean (range) of head capsule widths are 0.25 (0.25–0.25), 0.45 (0.42–0.46), 0.93 (0.66–1.10), 1.40 (1.21–1.57), and 2.16 (2.03–2.32) mm for instars 1–5, respectively. In Brazil, Specht et al. (2007) reported the egg, larval, prepupal, and pupal stages to average 3.0, 15.7, 1.9, and 8.8  days, respectively. Specht also gave mean head capsule measurements of 0.30, 0.50, 0.85, 1.42, and 2.74 mm for instars 1–5, respectively. The body color is green, and pale white stripes are usually evident. The head is green but marked with black spots at the base of setae. Numerous black tubercles are found dorsally on the body. The body bears microspines, but they are quite small and difficult to detect. The larva attains a length of about 22–33 mm at maturity. The lack of nipple-like structures on abdominal segments three and four separates bean leafskeletonizer from cabbage looper, Trichoplusia ni (Hübner), and soybean looper, Chrysodeixis includens (Walker). The lack of dark lateral bands on the head helps to distinguish bean leafskeletonizer from most of the other common loopers. A key to common vegetable-feeding loopers can be found in Appendix A. Pupa. Larvae roll the foliage, form a silken cocoon within the rolled leaves, and pupate within. The pupa measures about 17–18 mm long, 4.5 mm wide, and 0.2 g in weight. Duration of the pupal stage averages about 13.7 (range 9–12) days. Adult. The front wings of the adult are distinctively rusty brown, blending into gray, cream, or yellowish brown. The front wings lack the distinctive silver or white spot found centrally in most loopers. The hind wings are light grayish-brown basally and darker brown distally. Under laboratory conditions, the adults survived only about 5.3  days. A sex pheromone is known; the chemical blend is similar to that of other Plusiinae (Kaae et al., 1973). The only detailed reports on biology were provided by Casanova (1977) and Specht et al. (2007). Other useful information and diagnostic characters were given by Crumb (1956), Genung (1960), Eichlin and Cunningham (1978), and LaFontaine and Poole (1991). Larvae are included in the key by Stehr (1987).

Damage Both the leaf tissue and pods of beans are eaten. Although generally regarded as only a minor pest, this insect reportedly was very damaging at times to several varieties of beans in California. In Puerto Rico, it is most damaging to tobacco.

Management Foliar insecticides are effective if needed.

Beet Armyworm

Spodoptera exigua (Hübner) (Lepidoptera: Noctuidae)

Natural History Distribution. Beet armyworm is a tropical insect and is native to Southeast Asia. It is now found around the world, except South America. It was first discovered in North America about 1876 when it was found in Oregon. It rapidly spread to Hawaii (1880), California (1882), Colorado (1899), Texas (1904), Arizona (1916), Mississippi (1920), and Florida (1924). It has since spread to Central America and the Caribbean (Mitchell, 1979). As it is a tropical insect and lacks a diapause mechanism, it can overwinter successfully only in warm areas or in greenhouses. Daytime temperatures below 10°C are deleterious, and it rarely overwinters in areas where frost kills its host plants. Thus, overwintering is generally limited to Arizona, Florida, Hawaii, and Texas. Despite its inability to overwinter in most of the United States, beet armyworm nevertheless invades the southern half of the United States (Maryland to Colorado to northern California, and south) annually. Sometimes it is found as far north as New York and Ontario in the east, and British Columbia in the west. It is rarely viewed as a serious pest anywhere but the southern states, however, except sometimes in greenhouses. Beet armyworm is routinely a long-distance migrant, but unlike many migrants, migration does not precede the commencement of oviposition by the migrating generation. Instead, flight and oviposition occur at the same time, which is probably necessary because the adult has a relatively short life span (Jiang et  al., 2010). After migrating north in the summer, beet armyworm migrates south during autumn (Feng et al., 2003). Migration is aided by seasonal winds. Host Plants. This insect has a wide host range, occurring as a serious pest of vegetable, field, and ornamental (flower) crops; even trees are sometimes attacked. Among susceptible vegetable crops are asparagus, bean, beet, broccoli, cabbage, cauliflower, celery, chickpea, corn, cowpea, eggplant, lettuce, onion, pea, pepper, potato, radish, spinach, sweet potato, tomato, and turnip. Field crops damaged include alfalfa, corn, cotton, peanut, safflower, sorghum, soybean, sugarbeet, and tobacco.

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Weeds are also suitable for larval development, including common plants such as lambsquarters, Chenopodium album; mullein, Verbascum sp.; pigweed, Amaranthus spp.; purslane, Portulaca spp.; Russian thistle, Salsola kali; parthenium, Parthenium sp.; and tidestromia, Tidestromia sp. Although its host range is wide, there are significant differences in suitability even among hosts considered to be suitable. For example, in a comparison among diets consisting of sugarbeet, pigweed, or lambsquarters, the ­sugarbeet-fed larvae had the shortest development time and the highest fecundity, lambsquarter had the longest development time and the lowest fecundity, and pigweed was intermediate (Al-Zubaidi and Capinera, 1986). Greenberg et al. (2001) similarly compared growth and survival on different hosts and reported that larval development time was shortest on pigweed, intermediate on cotton, and longer on cabbage, pepper, and sunflower. Similarly, overall survival was highest on pigweed, intermediate on cotton and pepper, and longer on cabbage and sunflower. Natural Enemies. Throughout most of the world, numerous native natural enemies have adapted to this pest, resulting in a very long list of parasitoids and predators. Among the most common parasitoids in North America are Chelonus insularis Cresson, Cotesia marginiventris (Cresson), and Meteorus autographae (Muesbeck) (all Hymenoptera: Braconidae), and the tachinid Lespsia archippivora (Riley) (Diptera: Tachinidae) (Oatman and Platner, 1972; Tingle et al., 1978; Ruberson et al., 1994). Predators frequently attack the eggs and small larvae; among the most important are minute pirate bugs, Orius spp. (Hemiptera: Anthocoridae); big-eyed bugs, Geocoris spp. (Hemiptera: Lygaeidae); damsel bugs, Nabis spp. (Hemiptera: Nabidae); and a predatory shield bug, Podisus maculiventris (Say). Pupae are subject to attack, especially by the red imported fire ant, Solenopsis invicta Buren. Fungal diseases, Erynia sp. and Nomurea rileyi, and a NPV also inflict some mortality (Wilson, 1933, 1934; Harding, 1976b; Ruberson et al., 1994). The important mortality factors vary among crops and geographic regions. None except the NPV are highly specific to beet armyworm, which may explain why they are not especially effective. Virus is considered to be the most important mortality factor in Mexico (Alvarado-Rodriguez, 1987). Life Cycle and Description. Seasonal activity varies considerably according to climate. In warm locations such as Florida, all stages can be found throughout the year, though development rate and overall abundance are decreased during the winter months (Tingle and Mitchell, 1977). The life cycle can be completed in just 24  days, and six generations have been reared during 5  months of summer weather in Florida (Wilson, 1934). However, generation times of 50–126  days have been observed, with a total of five generations annually, in southern California (Campbell and Duran, 1929).

Egg. Eggs are laid in clusters of about 50–150 per mass. Females may deposit over 1200 eggs during their lifetime, but normal egg production is about 300–600. They are usually deposited on the lower surface of the leaf, and often near blossoms and the tip of the branch. The individual egg is circular when viewed from above, but when examined from the side it is slightly peaked, tapering to a point. The eggs are greenish to white and are covered with a layer of whitish scales that gives the egg mass a fuzzy or cottony appearance. They hatch in 2–3 days during warm, but the incubation period is extended to about 4  days in cool weather. The developmental threshold for eggs is estimated at 12.4°C.

FIG. 10.65  Beet armyworm larva. (Photo by L. Buss.)

Larva. Normally there are five instars, though additional instars are sometimes found. Duration of the instars under warm (summer) conditions is reported to be 2.3, 2.2, 1.8, 1.0, and 3.1 days, respectively (Wilson, 1932), and at constant 30°C instar development time was reported by Fye and McAda (1972) to be 2.5, 1.5, 1.2, 1.5, and 3.0 days, respectively. Total larval development time is also influenced by diet quality (Al-Zubaidi and Capinera, 1984). The developmental threshold for larvae is estimated at 13.6°C. Only 1 mm long at hatching, the larvae attain a mean length of 2.5, 5.8, 8.9, 13.8, and 22.3 mm during instars 1–5, respectively (Wilson, 1932). Head capsule widths average 0.25, 0.45, 0.70, 1.12, and 1.80 mm, respectively. The larvae are pale green or yellow during instars 1–2, but acquire pale stripes during instar three. During instar four, larvae are darker dorsally and possess a dark lateral stripe. Larvae during instar five are variable in appearance and tend to be green dorsally, with pink or yellow color ventrally, and a white stripe laterally. A series of dark spots or dashes is often present dorsally and dorsolaterally. Sometimes larvae are very dark. The spiracles are white with a narrow black border. The body is practically devoid of hairs and spines. In the western states, the larva

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of beet armyworm is easily confused with clover cutworm, Discestra trifolii (Hufnagel), but beet armyworm lacks the black pigment adjacent to the spiracles that is so evident in clover cutworm. In the southern states, the larva of beet armyworm is easily confused with southern armyworm, Spodoptera eridania (Cramer), but southern armyworm can be distinguished by the presence of a large dark spot laterally on the first abdominal segment that disrupts the lateral stripe. Beet armyworm occasionally bears a spot laterally, but if present it occurs on the mesothorax, not on the first abdominal segment. Initially, the larvae of beet armyworm are gregarious, feeding as a group and skeletonizing plant foliage. As they mature, larvae become solitary and quite mobile, often traveling from plant to plant. Cannibalism may occur when larvae are at high densities or feeding on food low in nitrogen (Al-Zubaidi and Capinera, 1983).

An overview of biology was given by Wilson (1932), and developmental biology by Wafa et al. (1969), Fye and McAda (1972), and Ali and Gaylor (1992). Brown and Dewhurst (1975) provided a detailed description of all stages and a comprehensive list of host plants. Rearing technology was discussed by many authors, including Cobb and Bass (1975) and Hartley (1990). A sex pheromone has been identified (Persoons et  al., 1981; Mitchell et  al., 1983). Effects of irradiation and potential for release of sterile insects have been investigated (Debolt and Wright, 1976). Larvae are included in keys by Okumura (1962), Oliver and Chapin (1981), Capinera (1986), Stehr (1987), Sparks and Liu (2001), and are included in a key to armyworms and cutworms in Appendix A. Adults are included in keys by Rings (1977a, b), and Capinera and Schaefer (1983). Heppner (1998) provided very useful keys to the adults and larvae of North American Spodoptera.

Pupa. Pupation occurs in the soil. The larva generally constructs a pupal chamber near the soil surface, digging only about 1 cm beneath the surface. The chamber is constructed from sand and soil particles held together with an oral secretion that hardens when it dries. The pupa is light brown and measures about 15–20 mm long. Duration of the pupal stage is 6–7  days during warm weather. Fye and McAda (1972) reported a pupal duration of 5.1  days at 30°C.

Damage

FIG. 10.66  Beet armyworm moth. (Photo by L. Buss.)

Adult. The moths are moderately sized, the wingspan measuring 25–30 mm. The front wings are mottled gray and brown, and normally with an irregular banding pattern and a light-colored bean-shaped spot. The hind wings are a more uniform gray or white color, and trimmed with a dark line at the margin. Mating occurs soon after the emergence of the moths, and oviposition begins within 2–3 days. Oviposition extends over a 3–7 day period, and the moths usually perish within 9–10 days of emergence.

Larvae feed on both foliage and fruit. Young larvae feed gregariously and skeletonize foliage. As they mature, larvae become solitary and eat large irregular holes in foliage. They also burrow into the crown or center of the head of lettuce, or on the buds of crucifers. As a leaf feeder, beet armyworm consumes much more cabbage tissue than diamondback moth, Plutella xylostella (Linnaeus), but it is less damaging than cabbage looper, Trichoplusia ni (Hübner) (East et al., 1989). This insect is also regarded as a serious pest of celery in California, and the damage is directly correlated with the abundance of late-instar larvae late in the season. However, damage to foliage and petioles (stalks) during the first half of the growing season is of little consequence because these plant parts are removed at harvest (van Steenwyk and Toscano, 1981). Tomato fruit is quite susceptible to injury, especially near fruit maturity, but beet armyworm is not considered to be as threatening to tomato as is corn earworm, Helicoverpa zea (Boddie) (Zalom et al., 1986a). In tomato crops, infestation early in the growing period is more damaging than later infestation, and as little as one caterpillar per 20 tomato plants can cause economic loss (Taylor and Riley, 2008). Larvae not only damage t­omato fruit but may appear as contaminants in processed ­tomato (Zalom and Jones, 1994).

Management Sampling. Pheromone traps can be used to detect the presence of adult beet armyworm (Trumble and Baker, 1984; Mitchell and Tumlinson, 1994). Visual sampling for damage and larvae, combined with an action threshold of 0.3 larvae per plant, was used successfully on cabbage in South Texas to determine the need for crop treatment with insecticides (Cartwright et  al., 1987). A binomial sequential sampling program for armyworm-damaged tomato fruit

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was ­developed by Wilson et al. (1983b). Egg distribution on tomato was studied by Zalom et al. (1983). Regular monitoring of crops, probably about twice per week, is recommended because adults frequently invade from surrounding crops or weeds (Edelson et al., 1988). Insecticides. In the southeast and southwest, the relatively high abundance of beet armyworm has stimulated the frequent application of insecticides. Chemical insecticides and Bacillus thuringiensis are commonly applied to foliage to protect against defoliation. Insecticide resistance is a major problem in management of this insect, possibly because it attacks crops such as flowers, cotton, and vegetable crops that are treated frequently with insecticides (Brewer et al., 1990; Brewer and Trumble, 1994). Beet armyworm abundance is favored by frequent insecticide use, and it is considered to be a secondary or induced pest in some crops (Eveleens et al., 1973). Also, intensive use of insecticides for beet armyworm control in vegetables such as celery has stimulated outbreaks of other pests, principally American serpentine leafminer, Liriomyza trifolii (Burgess). Beet armyworm larvae are susceptible to management with other products such as neem formulations (Prabhaker et  al., 1986). Eggs can be killed with petroleum oil (Wolfenbarger et al., 1970), and both eggs and young larvae can be controlled with foliar applications of 5% cottonseed oil, though at some concentrations oil is damaging to some plants (Butler Jr. and Henneberry, 1990a, b). Kaolin film particles applied to leaves inhibit oviposition by moths, and complete mortality to first instars (Showler, 2003). Pheromones can also be used to disrupt mating and to inhibit or eliminate reproduction. Saturation of the atmosphere around beet armyworm-susceptible crops has been estimated to decrease mating by 97% (Wakamura and Takai, 1992, 1995) and production of eggs and larvae by 57% and 95%, respectively (Mitchell et al., 1997). Cultural Practices. Host-plant resistance in several crops has been studied for its contribution to beet armyworm pest management. In tomato, for example, resistance is correlated with total glycoalkaloid concentration in the fruit tissue. However, leaf tissue does not have any effective antibiotic chemistry, so larvae are able to develop on plants even if they have unsuitable (glycoalkaloid-rich) tomato fruit (Eigenbrode and Trumble, 1994). The future for beet armyworm-resistant celery is most promising (Meade and Hare, 1991; Diawara et al., 1996). Fertilization of plants with nitrogen has numerous effects on beet armyworm. High rates of nitrogen fertilization increase the rate of development when larvae feed on these tissues, larvae feed preferentially on plants tissue containing high nitrogen, and female moths oviposit preferentially on plants with high nitrogen levels (Chen et al., 2008). Biological Control. Several insect pathogens may prove to be useful for suppression of beet armyworm. A NPV isolated from beet armyworm is fairly effective as a

bioinsecticide under greenhouse conditions, where inactivation by ultraviolet light in sunlight is not a severe problem (Smits et  al., 1987). It is as effective as commonly used insecticides (Gelernter et  al., 1986), but it is not commercially available. The fungus Beauveria bassiana has the same attributes and limitations (Barbercheck and Kaya, 1991). Entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) successfully infect both larvae and adults of beet armyworm, and infected adults can fly short distances, helping to spread the pathogens (Timper et  al., 1988). Use of nematodes is similarly constrained by environmental conditions, but these biological control agents are available commercially.

Bertha Armyworm

Mamestra configurata Walker (Lepidoptera: Noctuidae)

Natural History Distribution. Bertha armyworm is found widely in western North America but is known principally as a pest in Canada’s Prairie Provinces and British Columbia. This native species occurs from Ontario west to British Columbia, and south through the Rocky Mountain states to Mexico. It is usually associated with dry grassland areas. Host Plants. Although considered to be a general feeder, this species prefers plants in the families Cruciferae and Chenopodiaceae. The importance of bertha armyworm as a crop pest has grown as the popularity of canola, an important oilseed crop, has increased. However, it has long been known as a pest of potatoes in southern British Columbia. It is reported from several vegetables including bean, beet, cabbage, cauliflower, corn, lentil, lettuce, pea, potato, rhubarb, Swiss chard, tomato, and turnip. Other crops injured include alfalfa, canola, flax, red clover, sugarbeet, sweet clover, sunflower, tobacco, and wheat. Several flower crops are reported injured, including geranium, gladiolus, hollyhock, larkspur, poppy, petunia, sunflower, and zinnia. Among common weeds consumed are lambsquarters, Chenopodium album; mustard, Brassica spp.; and Russian thistle, Salsola kali. The natural hosts of bertha armyworm are not known, though plants in the family Chenopodiaceae are likely candidates. In many cases, crops are damaged after the larvae develop on weeds, despite not being able to complete their growth on the crops. The dispersal of hungry caterpillars to a host unsuitable for the growth of young larvae, but which they damage, is a common phenomenon among cutworms and armyworms. Natural Enemies. Parasitoids and diseases are known to affect bertha armyworm larvae. Of the several parasitoids known from this insect, Banchus flavescens Cresson (Hymenoptera: Ichneumonidae) and Athrycia cinerea

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(Coquillett) (Diptera: Tachinidae) are most common, but it has proved to be difficult to accurately assign causes to much of the mortality observed among larvae (Wylie and Bucher, 1977; Turnock, 1988). Among other parasitoids found in association with bertha armyworm are Apanteles xylinus (Say) (Hymenoptera: Braconidae), Ichneumon canadensis Cresson (Hymenoptera: Ichneumonidae), Eulophus sp. and Euplectris bicolor (Swederus) (both Hymenoptera: Eulophidae); and Panzeria ampelus (Walker), Exorista mella (Walker), and Phryxe pecosensis (Townsend) (all Diptera: Tachinidae). Evendon et al. (2017) have provided a list of known parasitoids. Diseases agents are important mortality factors. Among those known from bertha armyworm include two species of baculovirus (NPV), a fungus, and a microsporidian (unspecified and usually considered now to be a fungus). The NPV occurs infrequently at low armyworm densities, but its incidence increases at higher host densities. For example, Erlandson (1990) reported the occurrence of over 95% infection of late-instar larvae in Saskatchewan. Fairly high doses of virus are needed to infect late-instar larvae, so higher levels of infection occur if the virus is distributed early, among young larvae. The virus probably survives in the soil and is distributed naturally by blowing soil (Bucher and Turnock, 1983). Life Cycle and Description. There is a single generation annually in Canada, though museum records suggest two generations in southern areas of its range (Evendon et al., 2017). In Canada, this species overwinters in the pupal stage, with moths emerging principally in June and July to mate and oviposit. Larvae mature in late summer and autumn, and pupate in the soil. Biology has not been studied in more southern areas. Egg. The eggs are deposited in clusters of 50–500 on the lower surface of leaves. They are white initially but turn brown with age, and black just before hatching. In shape, the eggs resemble a slightly flattened sphere and bear about 38 ridges radiating from the apex. Duration of the egg stage is about 10, 5.5, 4, and 3  days at 15°C, 20°C, 25°C, and 30°C (Jones and Heming, 1979), respectively. Fecundity is reported to be up to 2100 eggs, but more typically 700–900 eggs per female when fed favorable host such as canola, and only about 280 eggs when fed less suitable hosts such as potato. Moths prefer to oviposit on plants that are flowering. Larva. The larval stages are variable in appearance. There are six instars. During the first four instars, the larva is green with narrow, white or yellow dorsal and subdorsal stripes, and has the appearance and habits of loopers. Young larvae are gregarious but older larvae are solitary. Throughout their larval development, they tend to feed nocturnally. If disturbed, young larvae quickly spin down from the plant on a strand of silk. The later instars may be

g­ reenish with conspicuous stripes, or brownish to blackish with less apparent stripes. The darker forms tend to occur where larvae feed in exposed habitats, with the green form often found feeding in sheltered locations. Diet also influences color. Mature larvae usually have narrow dorsal and subdorsal stripes that are yellow. An irregular orangish or brownish band may be present below the spiracles. The mature larva measures about 32 mm long, and the head capsule width is 3.1–3.2 mm. Bailey (1976a) studied larval development on several natural diets at 20°C and reported development times of about 3.1, 2.2, 2.3, 2.4, 2.5, and 8.5 days for instars 1–6, respectively, and total larval development time of about 20.5 days. Under field conditions, however, 6  weeks may be required for larval development. Larvae can develop at temperatures ranging from 8°C to 32°C. The head of the mature larva is yellowish to orange with darker spots. Diapause is induced by short daylengths or cool temperatures. The critical daylength for diapause induction varies with latitude. Even in diapause, pupae are susceptible to freeze-induced mortality. Snow cover is important in protecting pupae from cold weather, but survival can be estimated based on soil temperatures. Pupa. Pupation occurs in the soil, usually at a depth of 5–15 cm. The pupa is reddish brown, and measures about 18–20 mm long and 5–6 mm wide. The posterior end bears a pair of unusually long spines with curved tips. Although nearly all pupae remain in diapause, invariably a few continue their development and emerge late in the season, only to perish due to cold weather. Pupae usually perish during the winter in Manitoba in the absence of snow cover on the soil, but about half of the pupae survive if 5–10 cm of snow is present (Lamb et  al., 1985). The pupal stage is intolerant of warm conditions; exposure to temperatures of 25°C or higher results in the sterilization of adults (Bucher and Bracken, 1977). Duration of the pupal stage is commonly 8–9 months. Adult. The moths measure about 35–40 mm in wingspan. The forewing is predominantly gray, bearing patches of brown, black, olive, and white scales. The bean-shaped spot near the center of the forewing is mostly white, as is an irregular transverse line near the wing tip. The hind wing is grayish-white basally, and darker distally. Moths are nocturnal and feed avidly at flowers. The calling behavior of females occurs during the latter two-thirds of the scotophase, copulation begins at dawn, and oviposition occurs shortly afterwards. This is different from most noctuids, which often mate early in the dark period (Howlader and Gerber, 1986). The mated female typically lays 75% of her eggs within the first week of adult life. Summaries of bertha armyworm biology are given by King (1928b), Mason et al. (1998), and Evendon et al. (2017).

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A sex pheromone has been described (Underhill et al., 1977; Struble et al., 1984). Beirne (1971) provided good historical treatment in Canada. Thermal biology was described by Bailey (1976a, b) and Bodnaryk (1978). Bucher and Bracken (1976) described culture techniques. The larva is included in keys to noctuid larvae by Crumb (1956), Godfrey (1972), and Stehr (1987). Capinera and Schaefer (1983) included the adult in a pictorial key of moths.

Damage This is one of the species considered to be a “climbing cutworm” because it spends little time in association with the soil. Larvae initially feed on foliage, creating irregularly shaped holes in the leaves. This type of damage is usually negligible on pod-producing plants. As they mature, however, they will feed on other plant parts, including the flowers of clover; pods of canola, lentil, pea, and bean; bolls of flax; silk and ears of corn; fruit of tomato; and the head of cabbage.

Management Sampling. Moths can be captured in blacklight and pheromone traps, but light traps are not considered to be efficient (Bucher and Bracken, 1979). Pheromone traps are useful, though usually not completely specific for bertha armyworm (Steck et al., 1979a, b; Struble et al., 1984; Landolt, 2000a, b). Although pheromone traps give good indications of broad trends in armyworm abundance, larval populations in adjacent fields may differ considerably, depending on the relative attractiveness of the crops when the eggs are being deposited (Turnock, 1987). Thus, it is advisable to sample plants for larvae, using caution to prevent the larvae from dropping to the soil. Turnock and Bilodeau (1985) described sampling techniques for larvae. Fermented sugar solutions can also be used to attract moths. Food bait lures have the advantage of attracting both sexes, whereas pheromones only attract males. Based on the attractiveness of fermented sugars, the active chemicals were identified, and a combination of acetic acid plus 3-methyl1-butanol was shown to be highly attractive to bertha armyworm (Landolt, 2000a, b; Landolt and Alfaro, 2001). Insecticides. Application of chemical insecticides to foliage provides good suppression of larvae. Larvae are only weakly susceptible to the common formulations of Bacillus thuringiensis (Morris, 1986; Trottier et  al., 1988). Neem products act as growth regulators and feeding deterrents (Isman, 1993), but benefit under field conditions has yet to be demonstrated. Cultural Methods. Weed control is an important component of bertha armyworm management because weeds such as lambsquarters and wild mustard attract ovipositing females and serve as a source of larvae, which damage crops only after the preferred weeds are consumed. Therefore, it is advisable to keep crops free of weeds during the flight

period of adults, so they do not oviposit on favored weeds within otherwise nonpreferred crops. Tillage in the autumn can be destructive to overwintering pupae. Mortality can result from direct damage to pupae from tillage equipment, with increased exposure of pupae to cold temperatures, and to differential mortality to parasitoids. Parasitoids seem to be less affected than their armyworm hosts by the tillage of the soil (Turnock and Bilodeau, 1984). Biological Control. Entomopathogenic nematodes (Nematoda: Heterorhabditidae and Steinernematidae) have been evaluated for suppression of bertha armyworm larvae under laboratory conditions (Morris and Converse, 1991). Although bertha armyworm is more susceptible than other soil-dwelling insects under laboratory conditions, field evaluation is needed before nematodes can be recommended for bertha armyworm suppression. Potential naturals enemies have been imported from Europe, based on the similarities of bertha armyworm and Mamestra brassica L (Lepidoptera: Noctuidae). However, despite laboratory studies showing the potential for parasitism, the introduced parasitoids apparently did not survive on the Canadian Prairie (Evendon et al., 2017).

Bilobed Looper

Megalographa biloba (Stephens) (Lepidoptera: Noctuidae)

Natural History Distribution. This native insect is found throughout North and South America, including Hawaii. It is relatively infrequent, however, in western Canada and the northwestern United States. Apparently, it cannot overwinter in northern latitudes and reinvades the northern United States and southern Canada each summer. Host Plants. Bilobed looper is reported to feed on several families of plants, but only a few crops are affected. Among vegetable crops eaten are bean, cabbage, and lettuce. Other crops accepted include alfalfa, clover, and tobacco, as well as some ornamental plants such as geranium, gladiolus, ivy, and salvia. Weeds consumed include hedge nettle, Stachys sp.; sunflower, Helianthus sp.; vervain, Verbena sp.; and yellow thistle, Cirsium horridulum. Natural Enemies. The natural enemies of bilobed looper are unknown. Life Cycle and Description. Adults, eggs, and larvae have been found during the period of January-June in Florida, with most larvae observed in late spring (Martin et al., 1981a). Presumably, they disperse northward during the spring and summer months. Egg. The eggs of bilobed looper are hemispherical in shape, white in color, and bear ridges that radiate vertically. They hatch in 3–5 days, usually during the morning hours.

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FIG. 10.67  Bilobed looper caterpillar. (Photo by J. Capinera.)

Larva. There are five instars, with development times of about 3, 2, 3, 3, and 4 days, respectively, when reared at 25°C. Total larval development time is about 13, 16, and 25 days at 30°C, 25°C, and 20°C, respectively. Head capsule widths for the five instars are about 0.2, 0.4, 0.7, 1.2, and 1.9 mm, respectively. The larvae attain a length of about 30 mm at maturity. In form, larvae resemble most other related loopers; the body is distinctly broader at the posterior end and tapers toward the head. The general color of the larva is green, but there is a dark green dorsal stripe, 3–4 weak white lines running parallel to the dorsal green line, and a thin white line on each side just above the lateral spiracles. The head is green and has a strong black band on each side of the head. The thoracic legs are normally black and three pairs of thoracic legs are present. The larva of bilobed looper is easily confused with other loopers, but because it bears black thoracic legs and black bars on the side of the head it is most easily confused with alfalfa looper, Autographa californica (Speyer). In contrast to alfalfa looper, however, bilobed looper is weakly marked with stripes, and bears microspines on the abdomen. A key to c­ommon ­vegetable-feeding loopers can be found in Appendix A. Pupa. Pupation occurs in a thin, nearly transparent silk cocoon that is attached to the host plant or nearby vegetation. Pupae of bilobed looper are variable in color, usually mottled black with irregular tan or light green areas. Pupation requires about 6, 9, and 15 days at 30°C, 25°C, and 20°C, respectively.

FIG. 10.68  Moth of bilobed looper. (Photo by J. Capinera.)

Adult. The wingspan of the bilobed looper moth measures about 4 cm. The forewing is irregularly marked with pale brown to medium brown, and a silver bilobed spot is located near the center of the wing. The hind wing is gray to tan basally and darker brown distally. The prereproductive period of the adult is estimated at 2–4  days, and the reproductive period at 5–6  days. Adults often mate more than once. Total egg production is about 500 eggs per female. Adult longevity is estimated at 8.5 days for males and 10.0 days for females. Developmental biology and rearing information were provided by Beach and Todd (1988). Adult and larval descriptions, and keys to differentiate bilobed looper from related species were provided by Eichlin and Cunningham (1978), Crumb (1956), Capinera and Schaefer (1983), and Capinera (1986). Bilobed looper is also included in the key to vegetable-atttacking loopers in Appendix A.

Damage Bilobed looper is a defoliator, and mature larvae can consume considerable quantities of foliage during the final days of larval life. However, it rarely is sufficiently abundant to be cause for concern.

Management Moth populations can be monitored with blacklight traps. A sex pheromone component that attracts numerous noctuids in the subfamily Plusiinae, cis­-7-dodecenyl acetate, also is attractive to bilobed looper (Roelofs and Comeau, 1970). Larvae are readily controlled with foliar applications of insecticides, including Bacillus thuringiensis.

Black Cutworm

Agrotis ipsilon (Hufnagel) (Lepidoptera: Noctuidae)

Natural History Distribution. Black cutworm is found widely around the globe, though it is absent from some tropical regions and cold areas. Its origin is uncertain. It is more widespread and damaging, in the northern than the southern hemispheres. It is found annually throughout the United States and southern Canada but apparently does not overwinter in northern states and Canada. There is strong evidence that black cutworm disperses northward from the Gulf Coast region each spring. Long-distance dispersal of adults has long been suspected in Europe, China, and North America. The basic pattern is to move north in the spring, and south in the autumn. Studies in the United States demonstrated northward displacement of moths during the spring in the range of 1000 km in 2–4 days when assisted by northward ­flowing

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air (Kaster and Showers, 1982; Showers et  al., 1989; Smelser et al., 1991). Similar displacement to the south and ­southwest has been documented in the autumn (Showers et al., 1993). Host Plants. Black cutworm has a wide host range. Among vegetables injured are artichoke, asparagus, bean, beet, broccoli, cabbage, cantaloupe, carrot, cauliflower, celery, Chinese cabbage, corn, cowpea, cucumber, eggplant, garbanzo, garlic, kale, kohlrabi, lettuce, mustard, okra, onion, pea, pepper, potato, sweet potato, radish, spinach, squash, tomato, turnip, and watermelon. This species also feeds on alfalfa, clover, cotton, rice, sorghum, strawberry, sugarbeet, tobacco, and sometimes grains and grasses. Suitability of several grasses and weeds for larval development was studied by Busching and Turpin (1977); among the relatively suitable plants were bluegrass, Poa pratensis; curled dock, Rumex crispus; lambsquarters, Chenopodium album; yellow rocket, Barbarea vulgaris; and redroot pigweed, Amaranthus retroflexus. The preference by black cutworm for weeds is sometimes quite pronounced. Genung (1959), for example, demonstrated how black cutworm, the dominant cutworm species in southern Florida during the winter, would avoid feeding on beans if spiny amaranth, Amaranthus spinosus, was present. Adults feed on nectar from flowers. Deciduous trees and shrubs such as linden, wild plum, crabapple, and lilac are especially attractive to moths (Wynne et al., 1991). Natural Enemies. Numerous species of natural enemies have been associated with black cutworm, but data on their relative importance are scarce. However, Puttler and Thewke (1970) collected black cutworm larvae in Missouri and reported 69% parasitism, so natural enemies probably exact a significant toll on cutworm populations. Among the wasps known to attack this cutworm are Cotesia marginiventris (Cresson), Microplitis feltiae Muesebeck, Microplitis kewleyi Muesebeck, Meteorus autographae Muesebeck, Meterorus leviventris (Wesmael) (all Hymenoptera: Braconidae); Campoletis argentifrons (Cresson), Campoletis flavicincta (Ashmead), Hyposoter annulipes (Cresson), and Ophion flavidus Brulle (all Hymenoptera: Ichneumonidae). Larvae parasitized by Meteorus leviventris (Wesmael) consume about 24% less foliage and cut about 36% fewer seedlings (Schoenbohm and Turpin, 1977), so considerable benefit is derived from parasitism in addition to the eventual death of the host larva. Other parasitoids known from black cutworm include flies often associated with other ground-dwelling noctuids, including Archytas cirphis Curran, Bonnetia comta (Fallén), Carcelia formosa (Aldrich and Webber), Chaetogaedia monticola (Bigot), Eucelatoria armigera (Coquillett), Euphorocera claripennis (Macquart), Gonia longipulvilli Tothill, G. sequax Williston, Lespesia archippivora (Riley), Madremyia saundersii (Williston), Sisyropa eudryae (Townsend), and Tachinomyia panaetius (Walker) (all Diptera: Tachinidae). Predatory ground-dwelling

i­nsects such as ground beetles (Coleoptera: Carabidae) apparently consume numerous larvae (Best and Beegle, 1977; Lund and Turpin, 1977). Although Genung (1959) indicated that 75%–80% of cutworms could be killed by a granulosis virus, there is surprisingly little information on epidemiology and of natural pathogens. Rather, such pathogens as viruses, fungi, bacteria, and protozoa from other insects have been evaluated for black cutworm susceptibility; in most cases, only relatively weak pathogens have been identified (Ignoffo and Garcia, 1979; Grundler et al., 1987; Johnson and Lewis, 1982a, b). However, the Agrotis ipsilon nucleopolyhedrovirus was characterized in 1999 and is quite lethal to young larvae. Virus-induced mortality is enhanced by addition of an optical brightener (Boughton et al., 2001). An entomopathogenic nematode, Hexamermis arvalis (Nematoda: Mermithidae), is known to parasitize up to 60% of larvae in the midwestern states (Puttler and Thewke, 1971; Puttler et al., 1973). An ectoparasitic nematode, Noctuidonema guyanense, parasitizes the adult and is thought to be debilitating (Simmons and Rogers, 1996). Life Cycle and Description. The number of generations occurring annually varies from one to two in Canada to two to four in the United States. In Tennessee, moths are present in March-May, June-July, July-August, and September-December. Based on light trap collections, moths are reported to be abundant in Arkansas during May-June and September-October (Selman and Barton, 1972), and in New York, they occur mostly in June-July (Chapman and Lienk, 1981). However, light traps are not very effective during the spring flight and underestimate early-season densities (Willson et al., 1981; Levine et al., 1982). Thus, the phenology of black cutworm remains uncertain, or perhaps is inherently variable owing to the vagaries associated with long-range dispersal. Overwintering has been reported to occur in the pupal stage in most areas where overwintering occurs, but larvae persist throughout the winter in Florida. Pupae have been known to overwinter as far north as Tennessee, but apparently are incapable of surviving farther north (Story and Keaster, 1982a). Thus, moths collected in the midwestern states in March and April are principally dispersing individuals that are past their peak egg production period (Clement et  al., 1985). Nonetheless, they inoculate the area and allow production of additional generations, including moths which disperse north into Canada. Duration of the life cycle is normally 35–60 days. Egg. The egg is white initially but turns brown with age. It measures 0.43–0.50 mm long and 0.51–0.58 mm wide and is nearly spherical in shape, with a slightly flattened base. The egg bears 35–40 ribs that radiate from the apex; the ribs are alternately long and short. The eggs normally are deposited in clusters on foliage. Females may deposit 1200–1900 eggs. Duration of the egg stage is 3–6 days.

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FIG. 10.70  Adult of black cutworm. (Photo by J. Capinera.) FIG. 10.69  Larva of black cutworm. (Photo by J. Capinera.)

Larva. There are 5–9 instars, with a total of 6–7 instars most common. Head capsule widths are about 0.26–0.35, 0.45–0.53, 0.61–0.72, 0.90–1.60, 2.1–2.8, 3.2–3.5, 3.6– 4.3, and 3.7–4.1 mm for instars 1–8, respectively. Head capsule widths are very similar for instars 1–4, but thereafter those individuals that display 8–9 instars show only small increments in width at each molt and eventually attain head capsule sizes no larger than those displaying only 6–7 instars. Larval body length is reported to be 3.5, 5.3–6.2, 7, 10, 20–30, 30–45, 50, and 50 mm for instars 1–8, respectively. Duration of the larval stage is normally 20–40  days. Mean duration of instars 1–6 was reported to be 6.0, 5.0, 4.6, 4.3, 5.6, and 4.0 days, respectively, at 22°C. Larval development is strongly influenced by temperature, with the optimal temperature of about 27°C. Humidity is less important, but instars 1–5 thrive best at higher humidities. In appearance, the larva is rather uniformly colored on the dorsal and lateral surfaces, ranging from light gray or gray brown to nearly black. On some individuals, the dorsal region is slightly lighter or brownish, but the larva lacks a distinct dorsal band. Ventrally, the larva tends to be lighter in color. Close examination of the larval epidermis reveals that this species bears numerous dark, coarse granules over most of its body. The head is brownish with numerous dark spots. Larvae usually remain on the plant until the fourth instar, when they become photonegative and hide in the soil during the daylight hours. In these latter instars, they also tend to sever plants at the soil surface, pulling the plant tissue below ground. Larvae tend to be cannibalistic. Pupa. Pupation occurs below-ground at a depth of 3–12 cm. The pupa is 17–22 mm long and 5–6 mm wide, and is dark brown. Duration of the pupal stage is normally 12–20 days.

Adult. The adult is fairly large in size, with a wingspan of 40–55 mm. The forewing, especially the proximal twothirds, is uniformly dark brown. The distal area is marked with a lighter irregular band, and a small but distinct black dash extends distally from the bean-shaped spot. The hind wings are whitish to gray, and the veins marked with darker scales. The adult preoviposition period is about 7–10 days. Moths select low-growing broadleaf plants preferentially for oviposition, but lacking these, they deposit eggs on dead plant material. Soil is an unsuitable oviposition site (Busching and Turpin, 1976). The life cycle of black cutworm was described by Harris et  al. (1962b) and Abdel-Gawaad and El-Shazli (1971). Developmental data were provided by Satterthwait (1933), Luckmann et  al. (1976), Archer et  al. (1980), and Beck (1988). Laboratory culture on artificial media has been described (Reese et al., 1972; Blenk et al., 1985). Rings et al. (1974b) published a bibliography. The larva was described, and included in a key, by Crumb (1929, 1956). Larvae are also included in keys published by Okumura (1962), Rings (1977b), Oliver and Chapin (1981), Capinera (1986), and Stehr (1987), and is included in a key to armyworms and cutworms in Appendix A. Moths are included in keys of Rings (1977a) and Capinera and Schaefer (1983).

Damage This species occurs frequently in many crops and is one of the best-known cutworms. Despite the frequency of occurrence, however, it tends not to appear in great abundance, as is known in some other cutworms and armyworms. Black cutworm is not considered to be a climbing cutworm, most of the feeding occurring at soil level. However, larvae feed above-ground until about the fourth instar. Larvae can consume over 400 cm2 of foliage during their development, but over 80% occurs during the terminal instar, and about 10% in the instar immediately preceding the last (Satterthwait, 1933). Thus, little foliage loss is possible during the early stages of development. Once the fourth instar is attained,

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l­arvae can do considerable damage by severing young plants, and a larva may cut several plants in a single night. Plants tend to outgrow their susceptibility to injury. Showers et al. (1983) demonstrated that corn at the one-leaf stage was very susceptible to damage, but that by the four- or five-leaf stage plant yield was not decreased by larval feeding. Levine et al. (1983) showed that leaf feeding and cutting above the soil line were less damaging to corn than cutting at the soil surface and that subterranean damage was very injurious.

Management Sampling. Adult populations can be monitored with both blacklight and sex pheromone traps. However, several authors have noted the inefficiency of light traps. Light traps are relatively effective in the summer and autumn, but the late-season generations generally pose little threat to crops. Pheromone traps are more effective during the spring flight when larvae present the greatest threat to young plants (Willson et al., 1981). Trap color affects moth capture rate, with white and yellow traps capturing more than green traps (Hendrix III and Showers, 1990). Large larvae burrow in the soil and are difficult to observe. However, larvae can be sampled with bait traps, and this is most effective before emergence or planting of seedlings. Various trap designs have been studied, but many employ a container sunk into the soil with the upper lip at the soil surface. The container is baited with fresh plant material and/or bran, and with vermiculite so that the larvae can attain shelter. Larvae are effectively captured in baited containers if the vermiculite is not very near the surface (Story and Keaster, 1983), and catches are enhanced if a screen cylinder, which provides a visual stimulus to the cutworms, is suspended above the baited container (Whitford and Showers, 1984). If plants are present in the field they compete with the bait in the traps and trap efficiency declines markedly. The distribution of larvae in the spring is random (Story and Keaster, 1982b). Insecticides. Persistent insecticides are commonly applied to plants and soil for black cutworm suppression, but surface rather than subsurface soil applications are desirable (Foster et al., 1990). Larvae readily accept insecticidetreated bran and other baits (Sechriest and Sherrod, 1977; Gholson and Showers, 1979). Application of systemic insecticides to seeds also provides some protection against larval injury (Levine and Felsot, 1985; Berry and Knake, 1987). Bacillus thuringiensis is not usually recommended for cutworm control. Cultural Practices. Black cutworm larvae feed readily on weeds, and destruction of weeds can force larvae to feed exclusively on crop plants, exacerbating the damage. Thus, it is often recommended that weeds need not be tilled or treated with herbicide until larvae are matured.

Timing is important, however, because prolonged competition between crop and weed plants can reduce crop yield (Engelken et al., 1990). Presence of flowering weeds also can be beneficial by supporting prolonged survival of parasitoids (Foster and Ruesink, 1984). In contrast, reduced tillage cropping practices, which often produce higher weed populations, seem to result in increased abundance of black cutworm and higher levels of cutting in corn (Johnson et al., 1984; Tonhasca Jr. and Stinner, 1991; Willson and Eisley, 1992). This may be due, in part, to the tendency of moths to oviposit on weeds; weedy fields tend to have higher cutworm populations (Sherrod et al., 1979). Black cutworm populations also tend to be higher in wet areas of fields, and in fields that are flooded. Black cutworm has been known, at times, as “overflow worm,” due to its tendency to become abundant and damaging in fields that are flooded by overflowing rivers (Rockwood, 1925). In the home garden, barriers are sometimes useful to prevent damage to seedlings by cutworms. Metal or waxedpaper containers with both the top and bottom removed can be placed around the plant stem to deter consumption. Aluminum foil can be wrapped around the stem to achieve a similar effect. As the larvae burrow and feed below the soil line it is necessary to extend to barrier below the soil surface. Because black cutworm moths, which easily circumvent such barriers, are active during the growing season; this procedure alone may have little value. Use of netting or row covers, in addition to larval barriers, can prove more effective. Biological Control. Entomopathogenic nematodes (Nematoda: Steinernematidae and Heterorhabditidae) infect and kill black cutworm larvae, but their populations normally need to be supplemented to realize high levels of parasitism (Capinera et  al., 1988; Levine and OloumiSadeghi, 1992). Their effectiveness is related to soil moisture conditions (Baur et al., 1997).

Bronzed Cutworm

Nephelodes minians Guenée (Lepidoptera: Noctuidae)

Life history Distribution. Bronzed cutworm is widespread in distribution, occurring throughout the United States except for the southernmost tier of states. It is also found in southern Canada, from the Maritime Provinces to British Columbia. Despite the broad distribution of this species, its economic impact is limited to the eastern portion of its range, as far west as the Rocky Mountains. Also, it is rarely known to be damaging south of Kansas, Missouri, and Virginia. It is a native species. Host Plants. Bronzed cutworm larvae feed on grasses and grain crops such as barley and wheat. It is most f­ requently

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considered a pest of pasture and lawn grasses, especially Poa spp., and occasionally it damages field crops such as clover and sugarbeet. It commonly damages corn in the midwestern states, and when preferred plants are exhausted it may feed on other vegetables. On occasion, larvae are also observed to climb fruit trees and feed on the buds and leaves. Natural Enemies. Parasitoids and predators, though observed, seem less significance as mortality factors than viral diseases. Wasps known to attack bronzed cutworm include Rogas terminalis (Cresson), Apanteles rufocoxalis Riley (both Hymenoptera: Braconidae), and Campoletis oxylus (Cresson) (Hymenoptera: Ichneumonidae). Among parasitic flies reared from bronzed cutworm are Aplomya trisetosa (Coquillett), Euexorista futilis (Osten Sacken), Phryxe pecosensis (Townsend), and Tachinomyia variata Curran (all Diptera: Tachinidae). Western yellowjacket, Vespula pensylvanica (Saussure), is reported to prey on bronzed cutworm moths (Warren, 1990). A polyhedrosis virus has long been considered to be an important mortality factor (Walkden, 1937), but a granulosis virus has also been reported (Steinhaus, 1957). Life Cycle and Description. There is a single generation per year over the entire range of this insect. Adults are present in the autumn. In New York, moth flights consistently occur in September (Chapman and Lienk, 1981), in Minnesota they occur in late August and early September (Knutson, 1944), and in the central Great Plains their flights occur in September and October (Walkden, 1950). The eggs overwinter; however, egg hatch occurs early in the year, often in January and February. Larvae complete their development in April or May and become quiescent until July or August when pupation occurs. Egg. Moths are reported to scatter eggs singly at the surface of the soil. In shape, the egg has a slightly compressed sphere. It measures about 0.93 mm wide and 0.77 mm high. The egg is marked with about 250-min ribs that radiate outward from the center. In color, the egg is initially grayish, but soon acquires a pinkish tint and then a hint of purple. Duration of the egg stage, which is the overwintering form, is quite variable, but Walkden (1937) reported a mean of 127 days (range 98–145 days).

FIG. 10.71  Bronzed cutworm larva. (Photo by J. Capinera.)

Larva. There are 6–7 instars. Head capsule widths are about 0.5, 0.8, 1.2, 2.1, 3.0, and 4.3 mm for instars 1–6, respectively. Mean development time for larvae with six instars is 30.0, 10.5, 9.0, 8.2, 13.5, and 158.3 days, respectively. For larvae exhibiting seven instars, the duration of the first five instars is the same as with six-instar larvae, but the duration of instar six is 15.6 days, and instar seven is 147.3  days. Total larval development time is estimated to be 230 days. Body length increases from 3 to 5 mm in the first instar to about 35–45 mm at maturity. The mature instar is very distinctive in appearance, with a shiny bronze body and five sharply defined, broad stripes running to the length of the body. The stripes are whitish to yellowish. The head is orangish brown. The first four instars differ in background color, in that they are green instead of bronze, but also are marked with longitudinal stripes as found in the latter instars. Pupa. Mature larvae form a small cell in the soil for pupation. The pupa is brown and measures 23–33 mm long and 8–11 mm wide. Duration of the pupal stage is reported to average 27.3 days (range 24–34 days).

FIG. 10.72  Bronzed cutworm moth. (Photo by J. Capinera.)

Adult. The moth is reddish brown, the front wings marked with an irregular dark-brown band crossing the wing centrally. Both the front and hind wings may display a reddish or violet tint. The moth measures about 35–50 mm in wingspan. Adults live for about 14 days and commence oviposition when about 2-day old. Based on the dissection of eggs from adults, oviposition potential of about 1000 eggs is estimated. A sex attractant for male moths is known (Steck et al., 1977). Detailed description and biology of bronzed cutworm were given by Crumb (1926) and Walkden (1937). A bibliography was published by Rings et  al. (1974a). A comprehensive key to larvae of the Noctuidae, including this species, was presented by Crumb (1956). It is also included in less inclusive keys for caterpillar pests in Nebraska (Whelan, 1935) and Colorado (Capinera, 1986), and in Stehr (1987), and is a key to armyworms and cutworms in

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Appendix A. Moths are included in pictorial keys by Rings (1977a) and Capinera and Schaefer (1983).

Damage Larvae are defoliators and consume the leaves and stems of young plants. As they are present early in the year they normally damage only early-season plants.

Management Larvae have been controlled successfully with applications of residual insecticides to the soil and foliage. Bacillus thuringiensis is not often recommended for cutworms. Although there seems to be no report of experimentation with baits, treated bran would likely prove effective. Entomopathogenic nematodes have been proven to kill bronzed cutworm in turfgrass, and would likely be effective in vegetables as well.

Cabbage Looper

Trichoplusia ni (Hübner) (Lepidoptera: Noctuidae)

Natural History Distribution. The origin of cabbage looper is uncertain, but it is now found in Africa, Asia, Europe, and in the Americas. In North America, during the summer months, it is found throughout Canada, Mexico, and the United States wherever crucifers are cultivated. Overwintering in the United States apparently occurs only in the southernmost states, however. The lower limit for development is about 10–12°C, and temperatures of 40°C or higher can also be lethal to some stages. Thus, in Florida continuous activity occurs south of Orlando, whereas the remainder of Florida, southern Georgia, and portions of South Carolina experience intermittent activity during the winter, depending on the weather. North of these areas has no winter activity. It is somewhat erratic in occurrence—typically very abundant 1 year, and then scarce for 2–3 years; this is likely due to NPV and differing success in dispersal from southern to northern areas. Cabbage looper is highly dispersive and is sometimes found at high altitudes and far from shore. Flight ranges of approximately 200 km have been estimated. Host Plants. Cabbage looper feeds on a large number of cultivated plants and weeds. As its common name implies, it feeds readily on crucifers and has been reported damaging broccoli, cabbage, cauliflower, Chinese cabbage, collards, kale, mustard, radish, rutabaga, turnip, and watercress. Other vegetable crops injured include beet, cantaloupe, celery, cucumber, lima bean, lettuce, parsnip, pea, pepper, potato, snap bean, spinach, squash, sweet potato, tomato, and watermelon. Additional hosts are flower crops such as chrysanthemum, hollyhock, snapdragon, and sweetpea, and field crops such as cotton and tobacco. With such a wide

range of crops serving as suitable hosts, it is not surprising that an extremely wide range of broadleaf weeds also serve as hosts (Eichlin and Cunningham, 1978; Soo Hoo et  al., 1984), though the weeds vary somewhat with geographic locale, depending on availability. Surprisingly few common agricultural weeds are frequent hosts; among those that are suitable are lambsquarters, Chenopodium album; wild lettuce, Lactuca spp.; dandelion, Taraxacum officinale; and curly dock, Rumex crispus. Despite the wide host range of cabbage looper, not all hosts are equivalent. For example, Elsey and Rabb (1967) compared the suitability of collards and tobacco and reported differing suitabilities. Adults preferred to oviposit on collards when given a choice, and early instar survival was higher on collards. However, once the third instar was attained, survival was equivalent on both hosts, and pupal weights were only slightly diminished when larvae were reared on tobacco. Sutherland (1966) studied the growth rate of cabbage looper fed on a wide variety of crops and weeds. He found relatively small differences among the different crucifer crops, and many other vegetable crops and some weeds were equally suitable for larval growth. Soo Hoo et al. (1984) conducted one of the most complete studies of relative suitability and reported that only about one-third of the plants tested were suitable for complete development of larvae. A survey of looper pests infesting crops in Alabama revealed that though cabbage looper could be recovered from numerous hosts (clover, cotton, crucifers, peanut, soybean, sweet potato, tomato), most were found on cotton and crucifer crops. Soybean looper, Chrysodeixis includens (Walker), an insect easily confused with cabbage looper and having a similar broad host range, occurred predominately on soybean (Canerday and Arant, 1966). The adults feed on nectar from a wide range of flowering plants, including clover, Trifolium spp.; goldenrod, Solidago canadensis; dogbane, Apocynum spp.; sunflower, Helianthus spp.; and others. Natural Enemies. Cabbage looper is attacked by numerous natural enemies, and the effectiveness of each seems to vary spatially, temporally, and with crop environment. Most studies noted the effectiveness of wasp and tachinid parasitoids, and a NPV. Predation has not been well studied except in cotton. In studies conducted on collards in North Carolina, Elsey and Rabb (1970a) observed considerable variation in the impact of natural enemies between years. They determined that Trichogramma (Hymenoptera: Trichogrammatidae) egg parasitoids were not very important; they were reared from less than 5% of the eggs. They identified no major mortality factors until the fifth instar, despite the presence of considerable ‘disappearance’ during this period. Either predators or weather could account for these larval deaths. During the latter instars, Voria ruralis (Fallén) (Diptera: Tachinidae),

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an endoparasite attacking the medium- or large-size larvae, was the dominant cause of death, accounting for an average of about 53% mortality. Elsey and Rabb (1970b) presented the biology of this important parasitoid. Trichoplusia ni NPV caused about 12% mortality, and undetermined fungi about 10%. Copidosoma truncatellum (Dalman) (Hymenoptera: Encyrtidae) was the other significant mortality factor but accounted for only 6%–7% mortality. Copidosoma truncatellum oviposits in cabbage looper eggs, emerging from and killing the mature larvae or prepupae. In studies conducted in California involving cabbage, Oatman and Platner (1969) reported that egg parasitism of cabbage looper by Trichogramma, while variable, could reach about 35%. Larval parasitism averaged 38.9% and tended to increase toward the end of the year. The tachinid V. ruralis was the dominant parasitoid and was especially abundant in the autumn and winter months. The other principal parasitoids, especially during summer and autumn, were C. truncatellum and Hyposoter exiguae (Viereck) (Hymenoptera: Ichneumonidae). The latter species is a solitary endoparasite that attacks small larvae. A total of 24 species of parasitoids were observed: 14 wasps and 10 flies. Despite the abundance of parasitoids, however, the authors concluded that T. ni NPV was the key factor affecting populations. In Ontario, Harcourt (1963a) indicated that C. truncatellum was the most important parasitoid. No data were provided, but cabbage looper populations were said to be “frequently destroyed” by NPV. One of the most complete studies of cabbage looper natural enemies was conducted in California by Ehler (1977a, b), on cotton. He determined that the egg and early larval stages experience most of the generational mortality, and that predators and C. truncatellum were the most important elements contributing to this mortality. During the early larval instars, the minute pirate bug, Orius tristicolor (White) (Hemiptera: Anthocoridae), the big-eyed bug, Geocoris palens Stål (Hemiptera: Lygaeidae), and the damsel bug, Nabis americoferis Carayon (Hemiptera: Lygaeidae) were the predators responsible for most of the mortality. Ehler documented several mortality factors during the middle larval instars. The parasitoid Microplitis brassicae Muesebeck (Hymenoptera: Braconidae), a solitary endoparasite attacking small larvae, was the dominant mortality factor, but rarely exceeded 20% mortality. Other factors included H. exiguae and T. ni NPV, but both at levels of less than 10% mortality. During the late larval instars C. truncatellum inflicted 40%–50% mortality, and V. ruralis and T. ni NPV each caused less than 10% mortality. Pupal mortality was insignificant. Predation is rarely studied as it is very difficult to measure accurately. Barry et  al. (1974) attempted to assess the potential of selected predators of cabbage looper by using caged populations in Missouri. They reported that

the ­ damsel bug Nabis alternatus Parshley (Hemiptera: Nabidae) was most effective, the big-eyed bug Geocoris punctipes (Say) (Hemiptera: Lygaeidae) was intermediate, and the green lacewing Chrysoperla carnea (Stephens) was relatively ineffective in predation of cabbage looper on soybean. Sutherland (1966) suggested that general predators, including yellow-jackets (Hymenoptera: Vespidae) and birds, could be important mortality factors. The T. ni NPV is well-studied. Larvae normally die within 5–7 days of consuming virus inclusion bodies. Early signs of larval infection are a faint mottling of the abdomen in the area of the third to the sixth abdominal segments. This is followed by a more generalized blotchy appearance, and the caterpillar eventually becomes creamy white swollen, and limp. Death usually follows within hours following the limp condition, and caterpillars are often found hanging by their prolegs. Dark blotches appear after death, and the integument becomes very fragile and eventually ruptures. The body contents, heavily contaminated with new inclusion bodies, drip onto foliage where they can be consumed by other larvae (Semel, 1956; Drake and McEwen, 1959). Hofmaster (1961) reported that looper populations in Virginia were highest during dry weather because rainfall assisted the spread of NPV, and that this virus greatly suppressed loopers. In New York, Sutherland (1966) indicated that though T. ni NPV was an important mortality factor, natural incidence did not appear adequate to protect crops from damage. Life Cycle and Description. The number of generations completed per year varies from two to three in Canada, five in North Carolina, from five to seven in California. The generations overlap considerably and therefore are indistinct. Development time (egg to adult) requires 18–25 days when insects are held at 32–21°C, respectively (Toba et al., 1973), so at least one generation per month could be completed successfully under favorable weather conditions. There is no diapause present in this insect, and though it is capable of spending considerable time as a pupa, it does not tolerate prolonged cold weather. It reinvades most of the United States and all of Canada annually after overwintering in southern latitudes. The lower limit for development is about 10–12°C, and 40°C is fatal to some stages. Cabbage looper is considered to be a warm-weather insect; even in areas where it successfully overwinters it rarely occurs in high numbers until there has been adequate time for two to three spring generations. Sutherland (1966) conducted a survey of entomologists along the Atlantic coast and reported that looper populations were present year-round as far north as coastal South Carolina, and that looper infestations commenced in North Carolina and Maryland in May, in New Jersey in June, and in New York in July. Pennsylvania was not infested until August. Subsequent research by Chalfant et  al. (1974) clarified the winter activity patterns of cabbage looper in the southeastern United States: continuous

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activity and reproduction occur only in the part of Florida south of Orlando; the part of Georgia south of Byron as well as southeast South Carolina have intermittent adult activity during the winter months, depending on weather; all points north of this have no winter activity. Egg. Cabbage looper eggs are hemispherical in shape, with the flat side affixed to foliage. They are deposited singly on either the upper or lower surface of the leaf, though clusters of 6–7 eggs are not uncommon. The eggs are yellowish white or greenish, bear longitudinal ridges, and measure about 0.6 mm wide and 0.4 mm high. They hatch in about 2, 3, and 5 days at 32°C, 27°C, and 20°C, respectively, but require nearly 10 days at 15°C (Jackson et al., 1969).

FIG. 10.73  Cabbage looper larva. (Photo by L. Buss.)

Larva. Young larvae initially are dusky white but become pale green as they commence feeding on foliage. They are somewhat hairy initially, but the number of hairs decreases rapidly as larvae mature. Larvae have three pairs of prolegs and crawl by arching their back to form a loop and then projecting the front section of the body forward. The mature larva is predominantly green but is usually marked with a distinct white stripe on each side. The thoracic legs and head capsule are usually pale green or brown. Dorsally, the larva bears several narrow, faint white stripes clustered into two broad white bands. In some cases, the mature larva is entirely green. The body is narrower at the anterior end and broadens toward the posterior. It measures 3–4 cm long at maturity. Cabbage looper is easily confused with other loopers but can be distinguished from most by the presence of small, nipple-like structures (vestigial prolegs) located ventrally on abdominal segments three and four. Soybean looper, Chrysodeixis includens (Walker), also bears these structures, but usually has dark thoracic legs. Also, under high magnification it is possible to observe microspines on the body of soybean looper—a feature lacking from cabbage looper.

FIG. 10.74  Vestigial prolegs on abdominal segments 3 and 4. (Drawing by J. Capinera.)

The number of instars was given as 4–7 by Shorey et al. (1962), but many authors indicated only five. McEwen and Hervey (1960) gave mean head capsule width measurements as 0.29, 0.47, 0.74, 1.15, and 1.79 mm, respectively, for instars 1–5. Larval development required 17.8 and 19.9 days when reared on bean and held at 23°C and 32°C, respectively. When reared on cabbage at the same two temperatures, larval development required 19.9 and 20.8 days, respectively (Shorey et al., 1962). Development was also studied by Toba et  al. (1973), who determined that the number of larval instars could be increased from five to six by exposing the larvae to cooler temperature. Cool temperature also resulted in lowered egg production by ensuing adults. Pupa. At pupation, a white, thin, fragile cocoon is formed on the underside of foliage, in plant debris, or among clods of soil. The pupa contained within is initially green but soon turns dark brown or black. The pupa measures about 2 cm long. Duration of the pupal stage is about 4, 6, and 13 days at 32°C, 27°C, and 20°C, respectively.

FIG. 10.75  Cabbage looper adult. (Photo by L. Buss.)

Adult. The front wings of the cabbage looper moth are mottled gray brown; the hind wings are light brown at the base, with the distal portions dark brown. The forewing

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bears silvery-white spots centrally: a “U”-shaped mark and a circle or dot that are often connected. The forewing spots, though slightly variable, serve to distinguish cabbage looper from the most other crop-feeding noctuid moths. The moths have a wingspan of 33–38 mm. After a preoviposition period of about 1–2 days, females begin depositing their eggs, initially at about 80 per night. During the adult stage, which averages 10–12  days, 300– 600 eggs are produced by females having access to food, and less than 100 when only water is provided (Shorey, 1963). Moths are considered seminocturnal because feeding and oviposition sometimes occur about dusk. They may become active on cloudy days or during cool weather, but are even more active during the nighttime hours. They oviposit readily at temperatures as low as 15.6°C (Henneberry and Kishaba, 1967), but flight activity is higher on warmer evenings (Sutherland, 1966). Rearing procedures was given by McEwen and Hervey (1960) for plant-based culture and Shorey and Hale (1965) for artificial diet. Cabbage looper larvae are included in the keys by Okumura (1962), Capinera (1986), and Stehr (1987), and are included in a key to common loopers in Appendix A. Adults are included in the keys of Rings (1977a) and Capinera and Schaefer (1983).

Damage Cabbage looper replaced imported cabbage worm, Pieris rapae (Linnaeus), as the dominant cabbage caterpillar in the 1950s, apparently due to greater susceptibility of the latter to most insecticides. In recent years, diamondback moth has emerged as a more important caterpillar pest than cabbage looper; nevertheless, T. ni can be a serious problem. In studies conducted in South Carolina, diamondback moth was the major caterpillar pest in the spring crucifer crop, whereas cabbage looper predominated in the autumn crop (Reid Jr. and Bare, 1952). In Florida and Texas, however, spring populations of cabbage loopers can be damaging. Cabbage loopers are leaf feeders, and in the first three instars, they confine their feeding to the lower leaf surface, leaving the upper surface intact. The fourth and fifth instars chew large holes and usually do not feed at the leaf margin. With cabbage, however, they feed not only on the wrapper leaves, but also may bore into the developing head. Larvae consume three times their weight in plant material daily (McEwen and Hervey, 1960). Feeding sites are marked by large accumulations of sticky, wet fecal material. Despite their voracious appetite, larvae are not always as destructive as presumed. In California studies, feeding on celery during the first one-half of the growing season did not constitute loss because these petioles were routinely stripped from the plant at harvest (van Steenwyk and Toscano, 1981). With cabbage, moderate defoliation before the head formation is similarly irrelevant. In Texas, average

population densities of 0.3 larvae per plant justify control (Kirby and Slosser, 1984). In New York, Ohio, and Ontario, a density of 0.5 larvae per plant has been used as a treatment threshold (Shelton et  al., 1983b). In Florida and Georgia, one new feeding site per head is considered the damage threshold (Workman et  al., 1980). More recent work in Canada suggested that an appropriate action threshold was 40% of plants infested (Dornan et al., 1995). Cabbage looper can be a serious contaminant of fresh market broccoli and processed peas.

Management Sampling. Various sampling strategies have been developed for cabbage looper, and many approaches include consideration of the other crucifer-feeding caterpillars. Fixed sample units of at least 40 plants are sometimes recommended. However, larva-based sequential sampling (Shepard, 1973a) and variable intensity sampling (Hoy et  al., 1983) protocols have been developed to minimize the amount of sampling required to make appropriate management decisions. Similarly, an egg-based sequential sampling program was published (Muegge, 2004). Dornan et  al. (1995) recommended a binomial (presenceabsence) approach because it eliminated counting and insect identification. Blacklight traps and pheromone traps have been used in an attempt to predict looper population densities. Moth catches are monitored effectively by light traps (Hofmaster, 1961), but NPV, spread by rain, affects larval abundance and damage, thereby reducing predictability. The female-produced cabbage looper sex pheromone has at least seven chemical components, with Z-7-dodecenyl acetate the major component, but not all components are required to elicit attraction (Linn et  al., 1984). In addition, this insect has a male-produced pheromone consisting of S-(+) linalool, p-cresol, and m-cresol, that attracts females (Landolt et  al., 2004). Pheromone releasers and blacklight traps can be combined to increase moth catches, an approach that has been studied for area-wide suppression of cabbage loopers (Gentry et  al., 1971). Although numerous moths have been trapped by such techniques, and insects significantly decreased, suppression has not proven to be adequate to protect lettuce from damage (Debolt et al., 1979). Insecticides. Insecticide resistance has become a problem in cabbage looper control, but susceptibility varies widely among locations (Shelton and Soderlund, 1983). Botanical insecticides such as rotenone are less effective against cabbage looper than other cabbage-feeding Lepidoptera (Dills and Odland, 1948), but Bacillus thuringiensis kurstacki can be used, and neem functions as both a feeding deterrent and growth regulator (Isman, 1993). Resistance to B. thuringiensis is reported from greenhouses but is less of a problem under field conditions.

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Lures such asphenylacetaldehyde can be used to attract moths to bait stations where they encounter sugar solutions containing insecticide. This combination of lures and baits containing toxicants can be used to reduce the abundance of both male and female moths (Landolt et al., 1991). Biological Control. Microbial insecticides currently play a role in cabbage looper management, and their potential role has yet to be fully realized. Bacillus thuringiensis has long been used for effective suppression of cabbage looper (Kennedy and Oatman, 1976; Gharib and Wyman, 1991; Leibee and Savage, 1992), and has the advantage of not disrupting populations of beneficial insects. However, resistance in commercial greenhouses is a common problem (Janmaat and Myers, 2003). Trichoplusia ni NPV is effective (Hall, 1957), but it has not been commercialized because of the narrow host range. Home gardeners sometimes collect loopers dying of T. ni NPV, grind up the larval cadavers, and concoct their own effective microbial control agent. A NPV from alfalfa looper, Autographa californica (Speyer), has a wide host range, including cabbage looper (Jaques, 1977; Vail et  al., 1980; Tompkins et  al., 1986); it likely will become a useful tool for cabbage looper management. Mass release of Trichogramma spp. has been investigated for cabbage looper suppression. Looper egg parasitism can be increased several-fold by careful timing of parasitoid release (Oatman and Platner, 1971). Effectiveness varies among crops, however. This approach was most suitable in tomato, but also effective in crucifers and pepper (Martin et al., 1976b). Cultural Practices. Some differences in crucifer susceptibility have been observed. In New York, Dickson and Eckenrode (1975) found few significant differences, but red cabbages tended to be more resistant than kale or Chinese cabbage. In Wisconsin, Chinese cabbage, mustard, rutabaga, and turnip were less preferred for oviposition, whereas cabbage, Brussels sprouts, and collards were highly preferred. Unfortunately, there was no correlation between crops and varieties resistant to cabbage looper, and resistance to imported cabbage worm (Radcliffe and Chapman, 1966). Among cabbage cultivars studied in North Carolina, mammoth red rock and savoy perfection drumhead cultivars are considered to be relatively resistant, but this resistance dissipated under heavy insect feeding pressure. Interestingly, in this case, the resistant varieties received high numbers of cabbage looper eggs, but larval survival was poor (Chalfant and Brett, 1967). In studies of broccoli susceptibility in Virginia, Vail et al. (1991) found that early maturing varieties were less subject to attack than late-maturing varieties. Row covers, where economically practical, are effective at preventing cabbage looper moths from depositing eggs on crops. Manipulation of cropping patterns can benefit biological control of cabbage looper. In a study conducted in

Colorado, interplanting cabbage and nectariferous plants (alyssum and dill) enhanced parasitism (Al-Doghairi and Cranshaw 2004). Hooks and Johnson (2002), working in Hawaii, also reported that interplanted crops had lower levels of cabbage looper abundance, though the plant species affected the outcome.

Celery Looper

Anagrapha falcifera (Kirby) (Lepidoptera: Noctuidae)

Natural History Distribution. Celery looper is found throughout the United States and southern Canada. There is some question whether this native species overwinters in the northern United States and Canada. Research conducted in Iowa suggested that celery looper did not overwinter successfully, but was carried into the area in the spring when the appropriate weather patterns developed (Peterson et  al., 1988). This is highly plausible, as many other noctuids similarly overwinter in the south and disperse northward annually. However, there are also reports of this insect overwintering in the north in the larval stage, and adult activity was reported in New York from April to November (Chapman and Lienk, 1981). Host Plants. The host range of this insect is poorly known, but it appears that celery looper feeds on a large number of plants. Among vegetables damaged are celery, beet, cabbage, carrot, lettuce, and pea. Corn has been reported to be a host plant, but this is questionable. It is an occasional pest of sugarbeet and has been reported to feed on cranberry and hollyhock. Weeds fed upon include dandelion, Taraxacum officinale; plantain, Plantago sp.; and burdock, Arctium lappa. Coquillett (1881) reported oviposition on grass, so larvae also may develop on unknown grasses. Adults have been observed taking nectar from clover and lilac blossoms. Moths have also been found to be contaminated with pollen from several plant taxa, including Quercus, Rosaceae, and Pyrus, indicating an extensive range of adult host plants (Lingren et al., 1993). Natural Enemies. Little is known concerning the insect enemies of celery looper. A NPV is widespread and highly pathogenic to celery looper and may be the key factor that limits the abundance of this insect. This virus is unusual in that it is also pathogenic to numerous other species of Lepidoptera, affecting over 30 species in 10 families (Hostetter and Puttler, 1991). Due to its pathogenicity and wide host range it is being considered for use as a microbial insecticide. Life Cycle and Description. Most reports suggest two to three generations annually in northern states (Knutson, 1944). However, the generations overlap and it is difficult

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to discern population dynamics solely from capture rates of adults. For example, Chapman and Lienk (1981) presented data from New York showing the continuous occurrence of moths in all but the coldest months. Peterson et al. (1988) conducted a study in Iowa that included determination of ovary development, which aids in assessing the age of insects. Based on these studies, there were four generations annually.

Pupa. The mature larva spins a thin, white silken cocoon on the underside of a leaf or amongst the debris, and pupates within. The pupa is blackish brown and measures 13–15 mm long. Duration of the pupal stage is 10–20 days.

Egg. The eggs are milky white and measure about 0.5 mm in diameter. They are somewhat flattened and bear vertical ridges. They may be deposited singly or in small groups on either the upper or lower surfaces of vegetation. Duration of the egg stage is 4–6 days.

FIG. 10.77  Celery looper adult. (Photo by J. Capinera.)

FIG. 10.76  Celery looper larva. (Photo by J. Capinera.)

Larva. There are likely five instars, with each of about 3–5 days duration. Total larval development time is usually about 21 days, and larvae attain a length of about 35 mm. The larvae are green and tend to have a weak dark longitudinal line dorsally, accompanied by three narrow whitish lines on each side of the dorsal line. The thoracic legs are pale colored, and the abdominal tubercles located above the lateral spiracles are not black. The most distinctive marking is a narrow white line on each side, running through the lateral spiracles. The skin bears numerous minute spines, called microspines, over most of its surface. The spiracles are ringed with black pigment. If there is a dark bar on the head, it is indistinct. The caterpillar is more robust posteriorly and bears three pairs of prolegs. It is easily confused with other common loopers. Celery looper can be distinguished from bilobed looper, Megalographa biloba (Stephens), and alfalfa looper, Autographa californica (Speyer) by its pale thoracic legs and the absence of pronounced dark, lateral bars on the head; the latter species generally have dark bars and dark thoracic legs. Celery looper can be distinguished from cabbage looper, Trichoplusia ni (Hübner) by the presence of small, nipple-like structures located ventrally on the third and fourth abdominal segments, and the absence of dark microspines on the abdomen of the latter.

Adult. The moth measures 3.5–5 cm in wingspan. The front wings are dark, usually purplish brown and reddish brown, but the wings are marked distally with a silvery band. Many members of the noctuid subfamily Plusinae bear a spot near the center of the forewing, and this species is no exception. For celery looper, however, the spot is silver and is drawn out into a curving line that terminates at the posterior margin of the wing. Although not completely unique to celery looper, this pattern can serve to differentiate this moth from the other common vegetable pests such as cabbage looper, alfalfa looper, and bilobed looper. The front wings are also abnormally widened distally, due to a conical projection on the trailing edge of the forewing. The hindwings are yellowish brown, with darker brown bands and a whitish border. The biology of celery looper is poorly documented. Some elements of the biology were given by Coquillett (1881), Chittenden (1902), and Peterson et  al. (1988). Information on rearing and artificial diet were found in Treat and Halfhill (1973). Keys for the differentiation of celery looper moths from related species were given by Rings (1977a), Eichlin and Cunningham (1978), and Capinera and Schaefer (1983). Larvae are included in the keys of Crumb (1956), Capinera (1986), and is a key to common vegetablefeeding loopers in Appendix A.

Damage The larva is a defoliator, eating holes in the leaves of lettuce, celery, and other crops. It has been known to be destructive in Florida (Ball et  al., 1932), but generally, this insect is considered to be a minor pest. It sometimes is a serious contaminant of peas harvested for canning and freezing.

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Management Moths of this species can be monitored with a black-light trap. Butler et al. (1977) provided information on trapping celery looper with a sex pheromone. Insecticides applied to commercial or home garden crops for other insect pests are adequate to keep celery looper at very low levels of abundance. The microbial insecticide Bacillus thuringiensis is effective.

Clover Cutworm

Discestra trifolii (Hufnagel) (Lepidoptera: Noctuidae)

Natural History Distribution. Clover cutworm occurs throughout the United States except for the southeastern states. It also occurs throughout southern Canada and in Alaska. Apparently, it is a native species, though it is also reported to occur in Europe and Asia. Host Plants. This species is an occasional pest of several crops, preferring crops and weeds in the plant family Chenopodiaceae. Among vegetables attacked are beet, cabbage, lettuce, onion, pea, spinach, tomato, and turnip. Field crops consumed include alfalfa, cotton clover, flax, canola (rape), sugarbeet, and sunflower. The incidence of clover cutworm on sugarbeet explains the alternative common name, “striped beet caterpillar.” However, weeds are the usual host. Weeds known to be suitable larval food plants include kochia, Kochia scoparia; lambsquarters, Chenopodium album; purslane, Portulaca sp.; and Russian thistle, Salsola kali. Natural Enemies. Walkden (1950) reported that of fieldcollected larvae, 12% died from parasitic Hymenoptera, 16% from parasitic Diptera, and 22% from pathogens. Santiago-Alvarez and Federici (1978) reported Euplectrus sp. (Hymenoptera: Eulophidae) and Euphorocera tachinomoides Townsend (Diptera: Tachinidae) attacking larvae in southern California. Other wasps parasitizing clover cutworm are Apanteles plathypenae Muesebeck, Meteorus leviventris (Wesmael) (both Hymenoptera: Braconidae), and Enicospilus merdarius (Gravenhorst) (Hymenoptera: Ichneumonidae). Other parasitic flies include Euphorocera claripennis (Macquart) and Lespesia archippivora (Riley) (both Diptera: Tachinidae) (Arnaud Jr., 1978). Both granulosis and a NPVs are known from clover cutworm larvae, and Federici (1978) reported the granulosis virus to be especially important in regulating insect density. Federici (1982) also described an unusual rickettsia-like organism in clover cutworm larvae, but there is no indication that this pathogen occurs frequently. Life Cycle and Description. There are three generations annually in Colorado and Kansas. Moth flights occur in late May, early July, and late August-early September. Pupae

from the third generation overwinter. Knutson (1944) and Ayre et al. (1982a) suggested only two flights of adults in Minnesota and Manitoba, respectively, though the “second” flight period was protracted and may represent two overlapping periods of adult activity. Ayre et al. (1982a) suggested that the level of diapause induction in pupae was a critical determinant in overwintering survival in Manitoba; they speculated that if the last generation develops early in the season, diapause is not induced and the insects proceed with a generation that is not completed before the onset of winter. Egg. The eggs are deposited singly or in small clusters on the underside of leaves. They are white to pale yellow. They resemble a slightly flattened sphere and are equipped with ribs that radiate out from the top of the egg. The number of eggs produced by females is not well documented but seems to be in excess of 500 eggs, and is likely much greater. When reared on artificial diet, they generally produced 650–1700 egg per female (Santiago-Alvarez et  al., 1979). Duration of the egg stage is 4–5 days.

FIG. 10.78  Clover cutworm larva. (Photo by J. Capinera.)

Larva. The larvae are dull green. There may be a weak white line dorsally along the length of the body. The most distinctive character, however, is the combination of a broad lateral yellowish or pinkish band below the spiracles, and black pigmentation surrounding the spiracles. The black pigmentation forms a series of black spots immediately above the lateral band and serves to distinguish this caterpillar from beet armyworm, Spodoptera exigua (Hübner), a species that is superficially quite similar (Capinera, 1986). Larvae attain a length of 35–40 mm at maturity. Duration of the larval stage is 16–22 days under warm conditions, but it may be extended to nearly 50 days by cool weather. Pupa. Mature larvae burrow into the soil to a depth of about 2–3 cm to pupate. The pupa is reddish brown and measures 13–14 mm long. Duration of the pupal stage is 10–20 days during the spring and summer generations, but about 150 days for the overwintering population.

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of little value. It is possible to deny access by ovipositing moths through the use of netting and row covers, however.

Corn Earworm

Helicoverpa zea (Boddie) (Lepidoptera: Noctuidae)

Natural History

FIG. 10.79  Adult clover cutworm. (Photo by J. Capinera.)

Adult. The adults have a wingspan of 31–35 mm. The front wings are yellowish brown, and heavily marked with darker and lighter spots. The hind wings are grayish basally with a diffuse darker brown band distally. The compound eyes of this moth bear hairs—a feature that is useful for distinguishing it from some similar species. Females produce a sex pheromone that has been identified and synthesized (Struble and Swailes, 1977a). This insect is not well studied. Marsh (1913) gave a brief account of its biology. Walkden (1950) gave additional notes and provided a key to noctuids in the central Great Plains. Crumb (1956) described the mature larva and provided a key to larvae. The larva was also included in a key by Capinera (1986), and Stehr (1987), and in a key to armyworms and cutworms in Appendix A. The moth was included in pictorial keys developed by Rings (1977a) and Capinera and Schaefer (1983). Santiago-Alvarez et al. (1979) reported a suitable artificial diet.

Damage The larval stage defoliates plants, though it appears to favor weeds over crops. It is not a ground-dwelling species and does not sever plants at the soil surface, but climbs plants to feed on leaves. During periods of great abundance, it has caused significant damage and has been reported to assume a gregarious, dispersive “armyworm” habit.

Management Natural enemies, especially pathogens, generally serve to keep the population in check. Adult populations can be monitored with pheromone traps (Struble and Swailes, 1977a; Swailes and Struble, 1979; Ayre et al., 1982a). This species is reported to be easy to control with foliar applications of insecticides, though Bacillus thuringiensis is not often recommended. This species is a plant-inhabiting, climbing cutworm; therefore, the mechanical barriers recommended for protection of seedlings against ground-dwelling species are

Distribution. Corn earworm is found throughout North America except for northern Canada and Alaska. It tends to be less abundant west of the Rocky Mountains and is infrequently a pest in Canada’s Prairie Provinces. It also occurs in Hawaii and the Caribbean islands. Corn earworm is common in South America, persisting to a southern latitude of about 40°. Its origin is uncertain but likely is native to North America. In the eastern United States, corn earworm does not normally overwinter successfully in the northern states. It is known to survive as far north as about 40° north latitude, or about Kansas, Ohio, Virginia, and southern New Jersey, depending on the severity of winter weather (Blanchard, 1942). However, it is highly dispersive and routinely spreads from southern states into northern states and Canada (Hardwick, 1965b; Fitt, 1989; Westbrook et  al., 1997). Thus, areas have overwintering, both overwintering and immigrant, or immigrant populations, depending on location and weather. In the relatively mild Pacific Northwest, corn earworm can overwinter at least as far north as southern Washington. Host Plants. Corn earworm has a wide host range; hence, it is also known as “tomato fruitworm,” “sorghum headworm,” “vetchworm,” and “cotton bollworm.” In addition to corn and tomato, perhaps its most favored vegetable hosts, corn earworm also attacks artichoke, asparagus, cabbage, cantaloupe, collards, cowpea, cucumber, eggplant, lettuce, lima bean, melon, okra, pea, pepper, potato, pumpkin, snap bean, spinach, squash, sweet potato, and watermelon. Not all are good hosts, however. Harding (1976a), for example, studied relative suitability of crops and weeds in Texas and reported that though corn and lettuce were excellent larval hosts, tomato was merely a good host, and broccoli and cantaloupe were poor. Other crops injured by corn earworm include alfalfa, clover, cotton, flax, oat, millet, rice, sorghum, soybean, sugarcane, sunflower, tobacco, vetch, and wheat. Among field crops, sorghum is particularly favored. Cotton is frequently reported to be injured, but this generally occurs only after more preferred crops have senesced. Fruit and ornamental plants may be attacked, including ripening avocado, grape, peaches, pear, plum, raspberry, strawberry, carnation, geranium, gladiolus, nasturtium, rose, snapdragon, and zinnia. In studies conducted in Florida, Martin et al. (1976a) found corn earworm larvae on all 17 vegetable and field crops studied, but corn

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and sorghum were most favored. In cage tests, earworm moths preferred to oviposit on tomato over a selection of several other vegetables that did not include corn. Weeds such as common mallow, Malva neglecta; crown vetch, Coronilla varia; fall panicum, Panicum dichotomiflorum; hemp, Cannabis sativa; horsenettle, Solanum spp.; lambsquarters, Chenopodium album; lupine, Lupinus spp.; morningglory, Ipomoea spp.; pigweed, Amaranthus sp.; prickly sida, Sida spinosa; purslane, Portulaca oleracea; ragweed, Ambrosia artemisiifolia; Spanish needles, Bidens bipinnata; sunflower, Helianthus spp.; toadflax, Linaria canadensis; and velvetleaf, Abutilon theophrasti have been reported to serve as larval hosts (Ditman and Cory, 1931; Roach, 1975; Sudbrink Jr. and Grant, 1995). However, Harding (1976a) rated only sunflower as a good weed host relative to 10 other species in a study conducted in Texas. Stadelbacher (1981) indicated that crimson clover and winter vetch, which may be both crops and weeds, were important early-season hosts in Mississippi. He also indicated that cranesbill, Geranium dissectum and G. carolinianum, were particularly important weed hosts in this area. In North Carolina, especially important wild hosts were toadflax and deergrass, Rhexia spp. (Neunzig, 1963). Gross Jr. and Young (1977) documented some of the differences in suitability among various natural hosts relative to development time, weight gain, and fecundity. Adults collect nectar or other plant exudates from numerous plants. Lingren et al. (1993) studied adult host associations in Texas and Oklahoma and reported that trees and shrub species were especially frequented. Among the hosts identified were Citrus, Salix, Pithecellobium, Quercus, Betula, Prunus, Pyrus and other Rosaceae, and Asteraceae. Callahan (1958) also presented a long list of adult hosts. The quality or quantity of nectar affects potential fecundity of moths, with plants such as alfalfa; red and white clover; milkweed, Asclepias syriaca; and Joe-Pye weed, Eupatorium purpureum, proving especially suitable in Virginia (Nuttycombe, 1930). Natural Enemies. Although numerous natural enemies have been identified, they are usually not effective at causing high levels of earworm mortality or preventing crop injury. For example, in a study conducted in Texas, Archer and Bynum Jr. (1994) reported that less than 1% of the larvae were parasitized or infected with disease. However, eggs may be heavily parasitized (Oatman 1966a; Campbell et al., 1991). Trichogramma spp. (Hymenoptera: Trichogrammatidae), and to a lesser extent Telenomus spp. (Hymenoptera: Platygastridae), are common egg parasitoids. Also, natural control agents can affect populations late in the season (Roach, 1975). Exotic parasitoids and predators have been introduced to North America in the hope of gaining better natural control of H. zea, but thus far the imported beneficials have failed to establish successfully (King and Coleman, 1989).

Common larval parasitoids include Cotesia spp., and Microplitis croceipes (Cresson) (all Hymenoptera: Braconidae); Campoletis spp. (Hymenoptera: Ichneumonidae); Eucelatoria armigera (Coquillett) and Archytas marmoratus (Townsend) (Diptera: Tachinidae). However, additional wasp and fly species have, on occasion, been reported from corn earworm (Arnaud Jr., 1978; Krombein et  al., 1979). In Mississippi, Lewis and Brazzel (1968) observed only M. croceipes to be abundant regularly. General predators often feed on eggs and larvae of corn earworm; over 100 insect species have been observed to feed on H. zea. Among the common predators are lady beetles such as convergent lady beetle, Hippodamia convergens Guerin-Meneville, and Coleomegilla maculata De Geer (both Coleoptera: Coccinellidae); softwinged flower beetles, Collops spp. (Coleoptera: Melyridae); green lacewings, Chrysopa and Chrysoperla spp. (Neuroptera: Chrysopidae); minute pirate bug, Orius tristicolor (White) (Hemiptera: Anthocoridae); and big-eyed bugs, Geocoris spp. (Hemiptera: Lygaeidae) (King and Coleman, 1989), plant bugs (Hemiptera: Miridae), nabids (Hemiptera: Nabidae), and spiders (Pfannenstiel and Yeargan, 2002). In Kentucky, Pfannenstiel and Yeargan (2002) also noted that the ladybird beetle C. maculata accounted for almost half of the predation occurring in sweet corn. Birds can also feed on earworms, but rarely are adequately abundant to be effective (Barber, 1942). Within season mortality during the pupal stage seems to be slight (Kring et  al., 1993), and though overwintering mortality is often very high the mortality is due to adverse weather and collapse of emergence tunnels rather than to natural enemies. A nematode, Chroniodiplogaster aerivora (Nematoda: Diplogasteridae), occurs naturally in midwestern states, but it is a weak pathogen (Steinkraus et al., 1993b). In Texas, Steinernema riobrave (Nematoda: Steinernematidae) has been found to be an important mortality factor of prepupae and pupae, but this parasitoid is not yet generally distributed. Similarly, Khan et al. (1976) found Heterorhabditis heliothidis (Nematoda: Heterorhabditidae) parasitizing corn earwom in North Carolina, but it has not been found widely. Both of the latter species are being redistributed, and can be produced commercially, so in the future they may assume greater importance in natural regulation of earworm populations. Epizootics caused by pathogens may erupt when larval densities are high. The fungal pathogen Nomuraea rileyi and the Helicoverpa zea NPV are commonly involved in outbreaks of disease, but the protozoan Nosema heliothidis and other fungi and viruses also have also been observed. Life Cycle and Description. This species is active throughout the year in tropical and subtropical climates but becomes progressively more restricted to the summer months with increasing latitude. In northeastern states dispersing adults may arrive as early as May or as late as

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August due to the vagaries associated with the weather; thus, their population biology is variable. The number of generations is usually reported to be one in northern areas such as most of Canada, Minnesota, and western New York (Knutson, 1944; Beirne, 1971; Chapman and Lienk, 1981), two in northeastern states (Prostak, 1995), two to three in Maryland (Ditman and Cory, 1931), three in the central Great Plains (Walkden, 1950) and northern California (Okumura, 1962), four to five in Louisiana (Oliver and Chapin, 1981) and southern California (Okumura, 1962), and perhaps seven in southern Florida and southern Texas. The life cycle can be completed in about 30 days. Egg. The eggs are deposited singly, usually on leaf hairs and corn silk. The egg is pale green when first deposited, becomes yellowish and then gray with time. The shape varies from slightly dome-shaped to a flattened sphere, and measures about 0.5–0.6 mm in diameter and 0.5 mm high. They bear 21–31 ridges radiating from the center. Fecundity ranges from 500 to 3000 eggs per female. The eggs hatch in about 3–4 days.

reared at 25°C. Butler Jr. (1976) reported the cultured earworm on corn at several temperatures, reporting total larval development times of 31.8, 28.9, 22.4, 15.3, 13.6, and 12.6 days at 20°C, 22.5°C, 25°C, 30°C, 32°C, and 34°C, respectively. The larva is variable in appearance. Overall, the head tends to be orange or light brown with a white net-like pattern. The thoracic plate is black, and the body brown, green, pink, or sometimes yellow or mostly black. The larva usually bears a broad dark band laterally above the spiracles and a light yellow to white stripe below the spiracles. A pair of narrow dark stripes often occurs along the center of the back. The close examination reveals that the body bears numerous black thorn-like microspines. These spines give the body a rough feel when touched. The presence of spines and the light-colored head serve to distinguish corn earworm from fall armyworm, Spodoptera frugiperda (J.E. Smith), and European corn borer, Ostrinia nubilalis (Hübner). These other common corn-infesting species lack the spines and bear dark heads. Tobacco budworm, Chloridea virescens (Fabricius), is a closely related species in which the late instar larvae also bear microspines. Although it is easily confused with corn earworm, it is rarely a vegetable pest and never feeds on corn. Close examination reveals that in tobacco budworm larvae the spines on the tubercles of the first, second, and eighth abdominal segments are about half the height of the tubercles, but in corn earworm the spines are absent or up to one-fourth the height of the tubercle. Younger larvae of these two species are difficult to distinguish, but Neunzig (1964) gave a key to aid in separation.

FIG. 10.80  Corn earworm larva. (Photo by L. Buss.)

Larva. Upon hatching, larvae wander about the plant until they encounter a suitable feeding site, normally the reproductive structure of the plant. Young larvae are not cannibalistic, so several larvae may feed together initially. However, as larvae mature they become very aggressive, killing and cannibalizing other larvae. Consequently, only mature larva normally are found in each ear of corn. Normally, corn earworm displays six instars, but five is not uncommon and seven to eight have been reported. Mean head capsule widths are 0.29, 0.47, 0.77, 1.30, 2.12, and 3.10 mm, respectively, for instars 1–6. Larval lengths are estimated at 1.5, 3.4, 7.0, 11.4, 17.9, and 24.8 mm, respectively. Development time averaged 3.7, 2.8, 2.2, 2.2, 2.4, and 2.9 days, respectively, for instars 1–6 when

FIG.  10.81  Spinules associated with tubercles on the abdomen; the spines are less than one-fourth the height of the tubercle. (Photo by J. Capinera)

FIG. 10.82  Head capsule of corn earworm larva. (Photo by USDA.)

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Pupa. Mature larvae leave the feeding site and drop to the ground, where they burrow into the soil and pupate. The larva prepares a pupal chamber 5–10 cm below the surface of the soil. The pupa is mahogany brown, and measures 17– 22 mm long and 5.5 mm wide. Duration of the pupal stage is about 13 days (range 10–25 days) during the summer, but 250 days or more for the overwintering pupae.

including Whelan (1935), Walkden (1950), Frost (1955), Okumura (1962), Oliver and Chapin (1981), Capinera (1986), Stehr (1987), and Sparks and Liu (2001). Corn earworm is also included in a key to armyworms and cutworms in Appendix A. Keys to moths can be found in Rings (1977a) and Capinera and Schaefer (1983). Artificial diet and rearing procedures have been developed (Burton, 1970; Singh and Moore, 1985).

Damage

FIG. 10.83  Adult of corn earworm. (Photo by L. Buss.)

Adult. Because pupation occurs rather deep in the soil, the moths have difficulty digging to the surface unless the tunnel created by the larvae as they dig into the soil remains intact. As with the larval stage, adults are quite variable. The front wings of the moths usually are yellowish brown, and often bear a small dark spot centrally. The small dark spot is especially distinct when viewed from below. The forewing may also bear a broad dark transverse band distally, but the margin of the wing is not darkened. The hind wings are creamy white basally and blackish distally and usually bear a small dark spot centrally. The moth measures 32–45 mm in wingspan. Adults are reported to live for 5–15 days but may survive for over 30 days under optimal conditions. The moths are principally nocturnal and remain active throughout the dark period. During the daylight hours they usually hide in vegetation, but sometimes they can be seen feeding on nectar. Oviposition commences about 3  days after emergence and continues until death. Fresh-silking corn is highly attractive for oviposition but even ears with dry silk can receive eggs. Fecundity varies from about 500 to 3000 eggs, though feeding is a prerequisite for high levels of egg production. Females may deposit up to 35 eggs per day. The biology of corn earworm was presented by several authors; among the most complete were Quaintance and Brues (1905), Ditman and Cory (1931), Brazzel et al. (1953), Hardwick (1965b), and Neunzig (1969). An extensive bibliography was published by Kogan et  al. (1978). Keys to Helicoverpa adults and larvae were provided by Hardwick (1965b). Keys to differentiate earworm from similar crop-infesting larvae were given by many authors,

Some consider corn earworm is the most costly crop pest in North America. It is more damaging in areas where it successfully overwinters, however, because in northern areas it may arrive too late to inflict extensive damage. It often attacks harvested portions of valuable crops. Thus, larvae often are found associated with such plant structures as blossoms, buds, and fruits. When feeding on lettuce, larvae may burrow into the head. On corn, its most common host, young larvae tend to feed on silks initially and interfere with pollination, but eventually, they usually gain access to the kernels. They may feed only at the tip, or injury may extend half the length of the ear before larval development is completed. Such feeding also enhances development of plant pathogenic fungi and is attractive to sap beetles (Coleoptera: Nitidulidae). If the ears have not yet produced silk, larvae may burrow directly into the ear. They usually remain feeding within a single ear of corn, but occasionally they abandon the feeding site and begin search for another. Larvae also can damage whorl-stage corn by feeding on the young, developing leaf tissue. Survival is better on more advanced stages of development, however (Gross Jr. et al., 1976). Young fields adjacent to favored, more mature plants are likely to experience low rates of egg deposition (Weisenborn and Trumble, 1988). On tomato, larvae may feed on foliage and burrow in the stem, but most feeding occurs on the tomato fruit. Larvae commonly begin to burrow into a fruit, feed only for a short time, and then move on to attack another fruit. Tomato is more susceptible to injury when corn is not silking; in the presence of corn, moths will preferentially oviposit on fresh corn silk. Other crops such as bean, cantaloupe, cucumber, squash, and pumpkin may be injured in a manner similar to tomato, and also are less likely to be injured if silking corn is nearby.

Management Sampling. Eggs and larvae are often not sampled on corn because eggs are very difficult to detect, and larvae burrow down into the silks, where they are out of sight, soon after hatching. Sampling protocols have been developed for larvae on corn, however (Hoffmann et al., 1996b). On tomato, eggs tend to be placed on the leaves immediately below the highest flower cluster (Zalom et al., 1983). Thus, sampling protocols, including both fixed and sequential sampling

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procedures, have been developed for this crop (Hoffmann et al., 1991a). A common procedure is to examine the leaves beneath all flower clusters on 20–30 plants per field, but the sampling effort can be reduced significantly using sequential sampling. Similarly, sampling protocols for fruit damage have been developed (Wilson et al., 1983b). Moths can be monitored with blacklight and pheromone traps. Both sexes are captured in light traps whereas only males are attracted to the sex pheromone. Both trap types give an estimate of when moths invade or emerge, and relative densities, but pheromone traps are easier to use because they are selective. The pheromone is usually used in conjunction with an inverted cone-type trap. Trap designs and commercial source of pheromone lures affect trap catches (Gauthier et  al., 1991; Lopez Jr. et  al., 1994). Generally, the presence of 5–10 moths per night is sufficient to stimulate pest control practices (Foster and Flood, 1995). Light traps have also been investigated for the removal of moths from cropping areas. Some protection of small plantings can be attained, but this approach is ineffective for large areas or when moth densities are high (Barrett et al., 1971). Pheromone components can also be released in an area to confuse the moths and disrupt mating; this has been demonstrated experimentally (Mitchell et al., 1975a, b) but has not yet come into commercial practice. Insecticides. Cornfields with more than 5% of the plants bearing new silk are susceptible to injury if moths are active. Insecticides are usually applied to foliage in a liquid formulation, with particular attention to the ear zone, because it is important to apply insecticide to the silk, where the larvae will contact it before they burrow into the ear. Systemic insecticides are not effective because the plant does not translocate insecticide effectively to reproductive tissues (Russell et al., 1993). Considerable economic benefit has been documented for chemical suppression of earworm on tomato in North Carolina (Walgenbach and Estes, 1992). Insecticide applications are often done at 2- to 6-day intervals, sometimes as frequently as daily in Florida. Because it is treated frequently and over a wide geographic area, corn earworm has become resistant to many insecticides (Fitt, 1989; Kanga et al., 1996). Susceptibility to Bacillus thuringiensis also varies, but the basis for this variation in susceptibility is uncertain (Stone and Sims, 1993). Mineral oil, applied to the corn silk soon after pollination, has insecticidal effects. Application of about 0.75–1.0 mL of oil 5–7 days after silking can provide good control in the home garden (Carruth, 1942; Barber, 1942). Corn earworm moths, like many moths, feed readily on baits containing sweet material such as sucrose. Some work has been conducted to demonstrate that baits containing insecticide can attract and kill moths (Ditman, 1937; Creighton and McFadden, 1976), but this has yet to be developed into a practical technique. Cultural Practices. Cultural procedures have some application for corn earworm management. This dispersive

species moves readily from weeds to crops, and among crops, as their host plants become more or less suitable. Thus, effective management is best considered on an areawide basis (Graham et  al., 1972). Trap cropping is often suggested for this insect; the high degree of preference by ovipositing moths for corn in the green silk stage can be used to lure moths from less preferred crops. Lima beans are also relatively attractive to moths, at least as compared to tomato (Pepper, 1943). However, it is difficult to maintain attractant crops in an attractive stage for protracted periods. In southern areas where populations develop first on weed hosts and then disperse to crops, treatment of the weeds through mowing, herbicides, or application of insecticides can greatly ameliorate damage on nearby crops (Snodgrass and Stadelbacher, 1994). In northern areas, it is sometimes possible to plant or harvest early enough to escape injury. Throughout the range of this insect, population densities are highest, and most damaging, late in the growing season. Tillage, especially in the autumn, can significantly reduce overwintering success of pupae in southern locations (Barber and Dicke, 1937). Biological Control. Several insect pathogens have been evaluated for suppression of corn earworm. A NPV isolated from corn earworm is efficacious and has been sold commercially for larval suppression on nonfood crops (Young and McNew, 1994). Application of virus to weed hosts over an area-wide basis early in the season has been shown to reduce earworm population increase (Bell and Hayes, 1994; Hayes and Bell, 1994). Several other viruses, isolated from other caterpillars, are possible biological control agents if they attain commercial development (Young and McNew, 1994). In addition to the NPV, the fungus Nomuraea rileyi, the bacterium Bacillus thuringiensis, and steinernematid nematodes all provide some suppression (Oatman et  al., 1970; Ignoffo et al., 1978; Mohamed et al., 1978; Bartels and Hutchison, 1995). Entomopathogenic nematodes, which are available commercially, provide good suppression of developing larvae if they are applied to corn silk; this has application for home garden production of corn but not commercial production (Purcell et  al., 1992). Soil surface and subsurface applications of nematodes can also affect earworm populations because larvae drop to the soil to pupate (Cabanillas and Raulston, 1994, 1995, 1996). This approach may have application for commercial crop protection, but larvae must complete their development before they are killed, so some crop damage ensues. Trichogramma spp. (Hymenoptera: Trichogrammatidae) egg parasitoids have been reared and released for suppression of H. zea in several crops. Levels of parasitism averaging 40%–80% have been attained by such releases in California and Florida, resulting in fruit damage levels of about 3% (Oatman and Platner, 1971). The host crop seems to affect parasitism rates, with tomato being an especially suitable crop for parasitoid releases (Martin et al., 1976b).

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Host-Plant Resistance. Numerous varieties of corn have been evaluated for resistance to earworm, and some resistance has been identified in commercially available corn varieties (McMillian et  al., 1977; Story et  al., 1983; Archer et  al., 1994). Resistance is derived from physical characteristics such as husk tightness and ear length, which impede access by larvae to the ear kernels, or chemical factors such as maysin, which inhibit larval growth (Douglas, 1947). Hostplant resistance is a valuable component of multifaceted pest management programs. Varieties of sweet corn are now available that incorporate Bacillus thuringiensis (Bt) toxin, which reduces damage by H. zea, and other Lepidoptera (Benedict et  al., 1996; Burkness et  al., 2001; Horner et  al., 2003). Different Bt cultivars provide more or less protection, often causing 35%–75% reduction in damage. It is not necessarily desirable to obtain complete suppression of earworm, as this practice will likely lead to rapid selection for Bt-resistant insects. Indeed, resistance to Bt corn is increasingly a problem (Tabashnik et al., 2013). The resistance of corn to earworm is usually expressed by frequent sampling of plant tissue by larvae, prolonged larval development, smaller pupae, and reduced fecundity (Horner et al., 2003).

Darksided Cutworm

Euxoa messoria (Harris) (Lepidoptera: Noctuidae)

Natural History Distribution. Darksided cutworm, a native species, is found throughout most of Canada and the northern United States. Although it occurs as far south as South Carolina, Oklahoma, and southern California, it is a pest only in northern climates. Host Plants. This species has been reported to feed on numerous plants including such vegetables as bean, broccoli, Brussels sprouts, cabbage, cauliflower, Chinese cabbage, corn, cucumber, onion, pea, pepper, potato, radish, rutabaga, sweet potato, tomato, and turnip. In Canada, it is sometimes known as a field crop pest, injuring barley, flax, oat, rye, sugarbeet, tobacco, and wheat. The larvae also climb readily and damage the blossom and leaf buds of apple, currant, grape, peach, and other trees and shrubs. Natural Enemies. Bucher and Cheng (1971) studied mortality factors associated with darksided cutworm. They reported that bacterial, fungal (microsporidian), and other diseases collectively accounted for about 30% mortality, whereas NPV and other fungal pathogens occurred only at low levels. The other major mortality factor was parasitism, which caused about 15% mortality. Over 35% of cutworm larvae successfully pupated in these studies, which were conducted in Ontario. In another study, Cheng (1977) noted 10 species of parasitoids, including Meteorus

leviventris (Wesmael) (Hymenoptera: Braconidae); Eutanyacra suturalis (Say), Arenetra rufipes vernalis Walley, Campoletis flavicinctus (Ashmead), Enicospilus sp. (all Hymenoptera: Ichneumonidae); Copidosoma bakeri (Howard) (Hymenoptera: Encyrtidae); Muscina stabulans (Fallén) (Diptera: Muscidae); Winthemia rufopicta (Bigot), and W. deilephilae (Osten Sacken) (both Diptera: Tachinidae). However, most of the parasitism was due to a single species, C. bakeri. Other parasitoids known to affect darksided cutworm include Apanteles laeviceps Ashmead, A. militaris Walsh, Meteorus communis (Cresson) (all Hymenoptera: Braconidae); Spilichneumon superbus (Provancher), Diphyus euxoae Heinrich, Campoletis sp. (all Hymenoptera: Ichneumonidae); Sarcophaga cimbicis Town (Diptera: Sarcophagidae); Bonnetia comta (Fallén), and Aphria ocypterata Townsend (both Diptera: Tachinidae) (Cheng, 1977, 1981). Cheng (1973a) observed predation of larvae by ground beetles (Coleoptera: Carabidae) and by birds and suggested that rodents were important predators. Life Cycle and Description. There is a single generation throughout the geographic range of this species. Overwintering occurs in the egg stage. In Tennessee the eggs hatch in late January-late March, larvae complete their development in spring and early summer, and pupation occurs in June-August. Moths emerge beginning in September until about mid-October, and deposit overwintering eggs. In Ontario, the life cycle is similar but compressed owing to the shorter period of favorable weather. Thus, eggs hatch in late March-early May, pupation occurs in July-September, and adults are present in August-October. Egg. The eggs are deposited singly or in small clusters of up to 30 in the soil at a depth of 6–12 mm. They are difficult to locate in the soil because soil particles adhere to the chorion. Females provided with adequate nutrition (honey water) produced a mean of 1300 eggs per female over the course of its life. Even when provided only water, they produced a mean of 260 eggs (Cheng, 1972). The eggs initially are iridescent white, become yellow and then brown as the embryo develops. They are elliptical in shape, and measure about 0.55–0.63 mm in diameter and 0.38–0.45 mm in height, and bear slight ridges radiating from the center. Duration of the egg stage is 5–7 months. Larva. There are 6–8 instars. Crumb (1929) reported only six instars, but Cheng (1973a) reported mostly seven instars. Mean head capsule widths are 0.28, 0.39, 0.64, 0.95, 1.46, 2.18, and 3.10 mm for instars 1–7, respectively. The range of body lengths during the larval instars is about 2.2–3.0, 3.5–4.8, 5.5–7.8, 9.0–14.1, 16.2–23.0, 21.2–33.2, and 37.2–43.7 mm, respectively. Duration of the instars is about 17, 11, 10, 10, 10, 11, and 47 days, respectively, for a total larval development time of about 117 days (the last instar includes a lengthy nonfeeding prepupal period of

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about 30  days). Larvae are grayish, with irregular brownish longitudinal lines subdorsally and laterally. The common name “dark sided” is not particularly appropriate, and refers to the fairly indistinct, wavy dark band above the spiracles. Laterally, below the spiracles, is found a whitish band. The head is orange brown with darker spots. During the early instars larvae tend to feed on the upper surface of the leaves, leaving the lower epidermis intact. By the fourth instar, however, larvae consume leaves entirely. Large larvae, instars 6–7, may cut plants off at the soil surface. Throughout development, larvae tend to feed at night. When not feeding they may rest on the lower parts of plants or under leaves, but in the final instar they tend to hide in the soil when not eating. When larvae seek shelter in the soil the depth at which they bury themselves varies and is related to soil moisture. If the soil is moist they are buried just beneath the surface; under dry conditions they may dig to depths of 7–10 cm. Pupa. After larvae complete the feeding period they dig into the soil and create a pupal cell at a depth of 7.5–10 cm. As noted above, the larvae remain quiescent for about 30 days, then pupate. The pupa is initially yellowish brown but becomes dark brown toward maturity. The pupa measures about 16 mm long and 5 mm wide. Duration of the pupal stage is about 22 (range 18–28 days) days. Adult. The adult, like the larva, is not distinctly marked. The front wings are grayish brown and sometimes marked with dark transverse lines. A bean-shaped spot is evident on the forewing. The hind wings are grayish white, but darker distally and along the veins. The moth measures 30–40 mm in wingspan. Moths are most active 1–2 h after sunset, but mating apparently occurs shortly after midnight and oviposition occurs in the early morning. The female inserts her ovipositor into the soil to deposit eggs. In the laboratory, females produce about 1300 eggs. Adult longevity is usually 13–14 days. The biology of darksided cutworm was most completely described by Cheng (1973a), though Crumb (1929) gave a detailed description and some biological observations. Hinks and Byers (1976) and Belloncik et  al. (1985) gave information on rearing. Sex pheromone components were identified by Struble et  al. (1977) and Struble and Byers (1987). A bibliography was published by Rings et  al. (1975b). Larvae are included in keys by Crumb (1929), Whelan (1935), Rings (1977b), and Capinera (1986), and are included in a key to armyworms and cutworms in Appendix A. Adults are included in keys by Rings (1977a) and Capinera and Schaefer (1983).

Damage Larvae feed on the leaves and stems of young plants, sometimes causing complete defoliation and death of the plant.

Although widespread, this insect historically has been principally a pest of tobacco in southern Ontario. Elsewhere it is only an occasional pest.

Management Sampling. Larvae are difficult to sample, especially when they are young. Bucher and Cheng (1970) recommended transplanting a few small attractive plants such as tobacco into a field to serve as bait because larvae quickly accumulate in the area of such favored food plants. Both blacklight and pheromone traps can be used to monitor population density of adults. However, though both types of traps show similar population trends, pheromone traps are much more attractive (Cheng and Struble, 1982). There is some indication that the sex pheromone can also be used to disrupt mating, but this has not been fully evaluated (Palaniswamy et al., 1984). Insecticides. Insecticides are often used when this insect becomes troublesome. Insecticides may be applied to young plants or directly to the soil; in the latter case, it may be applied only to the surface or may be incorporated (Cheng, 1971, 1973b, 1980, 1984). Also, it is sometimes possible to apply the insecticide to the cover crop grown before vegetables, or to apply the insecticide postplanting as a rescue treatment following unexpected invasion by cutworms (Harris et  al., 1975b). Darksided cutworm is tolerant of some soil insecticides (Harris et al., 1962a). The microbial insecticide Bacillus thuringiensis was evaluated as a control agent for darksided cutworm by Cheng (1973c). Although young larvae are susceptible, by the time they attain the fourth instar susceptibility drops. Bacillus thuringiensis is not usually recommended for suppression of cutworms. Cultural Practices. Barriers are sometimes used to reduce access by cutworms to seedlings grown in the home garden. Metal or waxed-paper containers, with both the top and bottom removed, can be placed around the plant stem to deter consumption. Aluminum foil can be wrapped around the stem to achieve a similar effect. The barrier should be extended below the soil surface because larvae burrow and feed below the soil line.

Dingy Cutworm

Feltia jaculifera (Guenée) Feltia subgothica (Haworth) (Lepidoptera: Noctuidae)

Natural History Distribution. The name “dingy cutworm” is applied to at least two, and possibly four, species in North America. Possibly the most important is F. jaculifera, also known as F. ducens Walker. Second in importance usually is F. subgothica, but F. tricosa Lintner and F. herilis (Grote) are very

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similar in appearance and biology. The literature on these species is terribly confused, and they usually are treated as a complex of co-occurring species (Chapman and Lienk, 1981). The distribution of this cutworm complex is most of the United States and southern Canada. They are absent only from southern Florida and from California and adjacent desert areas (Rings et  al., 1975a, b). They are most damaging in the midwestern and eastern states, and eastern Canada. Host Plants. Dingy cutworm is commonly known as a corn pest, but also feeds on such vegetables as bean, cabbage, celery, cucumber, lettuce, onion, pea, squash, sweet potato, and tomato. Among other crops injured are alfalfa, blue grass, clover, flax, horseradish, raspberry, sweetclover, tobacco, and wheat. A wide variety of weeds are suitable food for dingy cutworm, including aster, Aster ericoides; chickweed, Stellaria sp.; goldenrod, Solidago sp.; mullein, Verbascum sp.; plantain, Plantago sp.; and yellow dock, Rumex crispus. Natural Enemies. Collection of larvae in spring from the central Great Plains has shown that natural enemies may account for 28% mortality, with most attributable to parasitoids (Walkden, 1950). Among the wasps parasitizing dingy cutworm are Aleiodes aciculatus Cresson, Chelonus sericeus (Say), Apanteles griffini Viereck, Microplitis feltiae Muesbeck, Meteorus leviventris (Wesmael), Spilichneumon superbus (Provancher) (all Hymenoptera: Braconidae), and Copidosoma bakeri (Howard) (Hymenoptera: Encyrtidae). Flies known to parasitize dingy cutworms include Euphorocera claripennis (Macquart), Gonia frontosa Say, G. fuscicollis Tothill, Triachora omissa (Aldrich), and Winthemia quadripustulata (Fabricius). A nematode, Hexamermis arvalis (Nematoda: Mermithidae) infects young dingy cutworms in the autumn and emerges in the spring, killing the larvae (Puttler and Thewke, 1971). Diseases known to affect dingy cutworm include the fungi Beauveria sp. and Metarhizium anisopliae and an unspecified virus (Crumb, 1929). Life Cycle and Description. There is a single generation of dingy cutworm annually. Moth flight occurs in JulySeptember followed immediately by oviposition. Larvae hatch and become partly grown before the onset of winter. Dingy cutworm overwinters in the larval stage, with larval development normally completed in March-May. The larvae remain quiescent in the soil until August, when pupation occurs, followed immediately by the emergence of the adults. Feltia jaculifera usually lags behind the other species in appearing as an adult. In New York, F. jaculifera normally first occurs in early August, whereas the other species comprising the dingy cutworm complex first appear in midJuly (Chapman and Lienk, 1981). Egg. The eggs reportedly are deposited on vegetation and on the soil surface. However, Balduf (1931) described

the deposition of eggs into flower heads of sunflowers. The eggs are oval, whitish to light brown, and the surface is marked by about 36 (F. jaculifera) or 56 (F. subgothica) narrow ridges. They measure about 0.60–0.65 mm long, 0.5 mm wide, and 0.36–0.38 mm in height. Duration of the egg stage is 5–21  days, depending on weather conditions, but normally 6–11 days. Based on dissections, females appear to be capable of producing about 800 eggs (range 500–1220 eggs). However, when Stanley (1936) captured moths feeding at flowers and confined them, he observed egg production of only about 100 eggs per female.

FIG. 10.84  Larva of dingy cutworm. (Photo by J. Capinera.)

Larva. Larvae display 6–7 instars, with 6 being normal. Head capsule widths are 0.30, 0.4–0.5, 0.55–0.80, 0.80–1.25, 1.30–1.90, 1.90–2.50, and 2.30–2.80 mm for instars 1–7, respectively. Duration of the instars is normally 3–12, 4–12, 4–12, 5–17, 8–119, 15–160, and 18–222 days, respectively. Total larval development time is 250–350 days. Body size increases from about 3 to 22–32 mm over the course of development. The body color is grayish brown. An interrupted blackish stripe is found dorsolaterally, bounding a broad lighter dorsal band. The head is brownish gray and marked with darker regions. The larvae of dingy cutworm are extremely hardy, able to tolerate long periods without food. They rarely display a tendency to climb, preferring to feed along the soil surface. Larvae are normally considered to be foliage feeders, and in the early spring, this is certainly the case. However, Balduf (1931) described pollen consumption by first instar larvae, and Duffus et al. (1983) indicated that larvae may feed until the fourth instar on sunflower heads. Pupa. Larvae prepare a cell in the upper 1–2 cm of the soil, and pupate within. The pupa is brown, and measures about 18 mm long and 6 mm wide. Duration of the pupal stage is 12–35 days.

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(Hufnagel) in abundance (Story et al., 1984). On occasion, larvae have been observed to ascend plants, including trees, to feed on buds and young foliage.

Management

FIG. 10.85  Dingy cutworm adult. (Photo by J. Capinera.)

Adult. The moths are brownish and measure 35–42 mm in wingspan. The front wings are grayish brown marked with darker brown and a hint of purple. The hind wings may be whitish basally and brown distally, or uniformly brown. The thorax and abdomen are gray to brown. Adults live for 5–10  days. Moths reportedly are affected by weather, with significant effects on reproductive success. Specifically, F. jaculifera is reported to be favored by dry weather during autumn, especially in October, because moths are most likely to fly and oviposit. In the case of F. subgothica, dry weather during September is favorable (Stanley, 1936). Sex pheromones have been described for F. jaculifera (Byers and Struble 1990). It appears, based on pheromone studies, that there is a genetic substructuring of the nominal species known as F. jaculifera. The ‘pheromonal strains’ respond to at least four slightly different pheromones (Byers et al., 1990), but the genetic differences among strains are, as yet, too small to be considered different species (Gooding et al., 1992). A detailed description of dingy cutworm is provided by Crumb (1929), and it was included in keys by Whelan (1935), Crumb (1956), Rings (1977b), Capinera (1986), and in a key to armyworms and cutworms in Appendix A. The moths were included in keys by Rings (1977a), Oliver and Chapin (1981), and Capinera and Schaefer (1983). Developmental data were provided by Walkden (1950). Chapman and Lienk (1981) made valuable observations on the species complex comprising dingy cutworm. A bibliography on dingy cutworm was published by Rings et al. (1975a).

Damage Larvae damage young plants in the spring, usually by cutting the seedlings off at the soil surface. In a survey of midwestern cornfields conducted between 1979 and 1981, dingy cutworm was the second most abundant cutworm encountered, following only black cutworm, Agrotis ipsilon

Sampling. The adult populations may be monitored with light or pheromone, but because plant damage does not occur for several months after adult activity, larval monitoring is needed. Duffus et  al. (1983) compared five types of larval sampling protocols and recommended sack trapping—the collection of larvae from beneath squares of plastic or burlap. Insecticides. Cutworms can be controlled by the application of persistent insecticides to soil or plants, or by application of baits such as bran that have been treated with insecticide. Bacillus thuringiensis is not usually recommended for control of this insect. Cultural Practices. If seedlings are to be transplanted into the home garden, larger plants are preferred, because they are less likely to be irreparably damaged by cutworms. Transplanted plants can be protected if surrounded by a barrier such as a can or waxed-paper container with the bottom removed. Aluminum foil wrapped around the base of the seedling also deters cutting by larvae.

Fall Armyworm

Spodoptera frugiperda (J.E. Smith) (Lepidoptera: Noctuidae)

Natural History Distribution. Fall armyworm is a native to the tropical regions of the western hemisphere from the United States to Argentina. It normally overwinters successfully in the United States only in southern Florida and southern Texas, but during warm winters it may survive along the Gulf Coast and in southern Arizona. It is commonly found in the Caribbean, including Puerto Rico. Fall armyworm is a strong flier and disperses to long distances annually during the summer months. It is recorded from virtually all states east of the Rocky Mountains, from Arizona and California, and from southern Ontario. As a regular and serious pest, its range tends to be mostly the southeastern states, though it is feared by sweet corn growers as far north as the New England states. In 2016 it was reported for the first time in West and Central Africa (Goergen et al., 2016), so it now threatens most of Africa and southern Europe. Host Plants. This species seemingly displays a very wide host range, with over 80 plants recorded, but clearly prefers grasses. The most frequently consumed plants are field corn and sweet corn, sorghum, Bermudagrass, and grass weeds such as crabgrass, Digitaria spp. When the larvae are very numerous they defoliate the preferred

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plants, acquire an “armyworm” habit and disperse in large ­numbers, consuming nearly all vegetation in their path. Many host records reflect such periods of abundance and are not truly indicative of oviposition and feeding b­ ehavior ­under normal conditions. Vegetables, other than sweet corn which is frequently at risk, are only occasionally damaged, but include a wide range of crops such as asparagus, bean, beet, cabbage, chickpea, cowpea, corn, cucumber, kale, onion, pea, pepper, potato, rutabaga, spinach, sweet potato, tomato, turnip, and watermelon. Field crops are frequently injured, including alfalfa, barley, Bermudagrass, buckwheat, cotton, clover, corn, oat, millet, peanut, rice, ryegrass, sorghum, sugarbeet, sudangrass, soybean, sugarcane, timothy, tobacco, and wheat. Other crops occasionally injured are apple, grape, orange, papaya, peach, strawberry and many flowers. Among the weeds known to serve as hosts are bentgrass, Agrostis sp.; crabgrass, Digitaria spp.; Johnsongrass, Sorghum halepense; morningglory, Ipomoea spp.; nutsedge, Cyperus spp.; pigweed, Amaranthus spp.; and sandspur, Cenchrus tribuloides. In studies conducted in Honduras, fall armyworm larvae preferred Amaranthus foliage over both corn and sorghum (Portillo et al., 1996b). Pencoe and Martin (1981) measured development on several grasses found in Georgia, and determined that large crabgrass, Digitaria sanguinalis; goosegrass, Eleusine indica; vaseygrass, Paspalum urvillei; and coastal Bermudagrass, Cynodon dactylon were very suitable hosts whereas yellow nutsedge, Cyperus esculentus; the sedge, Cyperus globulosus, and Texas panicum, Panicum texanum, were relatively poor hosts. There is some evidence that fall armyworm strains exist, based primarily on their host plant preference. One strain feeds principally on corn, but also on sorghum, cotton, and a few other hosts if they are found growing near the primary hosts. The other strain feeds principally on rice, Bermudagrass, and Johnsongrass. Some reproductive isolation exists between the strains, even when both occur in the same area (Pashley, 1988; Nagoshi and Meagher, 2004, 2008). Pannuti et al. (2016) studied larval feeding behavior, and reported that although young (vegetative stage) corn leaf tissue is suitable for growth and survival, on more mature plants the leaf tissue is unsuitable, and the larvae tend to settle and feed in the ear zone, and particularly on the silk tissue. However, silk is not very suitable for growth. Larvae feeding on corn kernels display the fastest rate of development. Also, although the closed tassel is suitable with respect to survival, it results in poor growth. Thus, tassel tissue may be suitable for initial feeding, perhaps until the larvae locate the silk and ears, but feeding only on tassel tissue is suboptimal. Natural Enemies. Cool, wet springs followed by warm, humid weather in the overwintering areas favor survival and reproduction of fall armyworm, allowing it to escape

suppression by natural enemies. Once dispersal northward begins, the natural enemies are left behind. Therefore, ­ though fall armyworm has many natural enemies, a few act effectively enough to prevent crop injury. Numerous species of parasitoids are known from throughout the range of fall armyworm, mostly in the families Braconidae and Ichneumonidae (both Hymenoptera), and Tachinidae (Diptera), but not all occur in North America. The known parasitoids, nearly all of which attack the larval stage, were listed by Ashley (1979) and Molina-Ochoa et  al. (2003). The latter authors list 95 species of parasitoids from North America, 86 from Central America and the Caribbean Region, and 86 from South America. The parasitoids most frequently reared from larvae in the United States are Cotesia marginiventris (Cresson) and Chelonus texanus (Cresson) (both Braconidae), species that are also associated with other noctuid species. In a study conducted in Georgia, for example, C. marginiventris was the most abundant parasitoid collected in 1990, attaining up to 34% parasitism early in the season, but Chelonus insularis Cresson (Hymenoptera: Braconidae) assumed dominance late in the season (Riggin et  al., 1992). During the second year of the study, however, the most abundant parasitoid was Archytas marmoratus (Townsend) (Diptera: Tachinidae), followed closely by Ophion flavidus Brulle and Aleiodes laphygmae (Gahan) (both Hymenoptera: Ichneumonidae). Also, in a subset of the study, the same authors found Aleiodes laphygmae and Ophion flavidus to be the dominant parasitoids (Riggin et al., 1993). Luginbill (1928) and Vickery (1929) described and pictured many of the fall armyworm parasitoids. An ectoparasitic nematode, Noctuidonema guyanense, parasitizes the adult and is thought to be debilitating (Simmons and Rogers, 1996). The predators of fall armyworm are general predators that attack many other caterpillars. Among the predators noted as important are various ground beetles (Coleoptera: Carabidae); the striped earwig, Labidura riparia (Pallas) (Dermaptera: Labiduridae); the spined soldier bug, Podisus maculiventris (Say) (Hemiptera: Pentatomidae); and the insidious flower bug, Orius insidiosus (Say) (Hemiptera: Anthocoridae). Vertebrates such as birds, skunks, and rodents also consume larvae and pupae readily. Predation may be quite important, as Pair and Gross Jr. (1984) demonstrated loss of pupae to predators at 60%–90% in Georgia. Numerous entomopathogens, including viruses, fungi, protozoa, nematodes, and a bacterium are associated with fall armyworm (Gardner et al., 1984), but only a few cause epizootics. Among the most important are the S. frugiperda NPV, and the fungi Entomophaga aulicae, Nomuraea rileyi, and Erynia radicans. Incidence of NPV reached 50%–60% in Louisiana, but disease typically appears too

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late to ­alleviate high levels of defoliation. A most interesting ­pathogen is the ectoparasitic nematode Noctuidonema guyanense (Nematoda: Aphelenchoididae). This nematode is a weak pathogen, h­ aving a debilitating effect on its host. Although fall armyworm is the principal host, it is associated with many other Lepidoptera (Rogers et  al., 1991; Simmons and Rogers, 1996). Life Cycle and Description. The life cycle is completed in about 30 days during the summer, but 60 days in the spring and autumn, and 80–90 days during the ­winter. The number of generations occurring in an area varies with the appearance of the dispersing adults. The ability to ­diapause is not present in this species. The population often spreads northward at about 480 km (300 miles) per generation. In Minnesota and New York, where fall armyworm moths do not appear until August (Knutson, 1944; Chapman and Lienk, 1981), there may be but a single generation. The number of generations is reported to be one to two in Kansas (Walkden, 1950), three in South Carolina (Luginbill, 1928), and four in Louisiana (Oliver and Chapin, 1981). In coastal areas of north Florida, moths are abundant from April to December, but some are found even during the winter months (Tingle and Mitchell, 1977). With the aid of certain weather patterns, dispersal northward may be much faster than just described. For example, during 1973 a redistribution of moths from Mississippi to Ontario occurred in about 2  days, a distance of 1600 km (1000 miles), with the aid of strong surface winds (Rose et al., 1975). Egg. The egg of fall armyworm is dome-shaped; the base is flattened and the egg curves upward to a broadly rounded point at the apex. It is well-marked with 47–50 ridges that radiate outward from the apex. The egg measures about 0.4 mm in diameter and 0.3 mm in height. They are deposited on hosts and nonhosts; in the latter case the larvae disperse, often with the help of a strand of silk, which allows them to be blown a considerable distance by wind. The female typically produces several egg masses during her oviposition period, with deposition occurring at night, and on larger plants if provided a choice between large and small. The number of eggs per mass varies considerably but it is often 100–200, and total egg production per female averages about 1500, with a maximum of over 2000. They are sometimes deposited in layers, but most are spread over a single layer and are attached to foliage. The female also deposits a layer of grayish scales between the eggs and over the egg mass, imparting a furry or moldy appearance. Initially the eggs are grayish green, but they soon turn brown. Egg masses are deposited beneath leaves when the moth density is low, but oviposition becomes indiscriminate at high densities. The period of incubation is only 2–3 days during the summer months.

FIG. 10.86  Fall armyworm larva. (Photo by P. Choate.)

Larva. There are usually six instars in fall armyworm. Head capsule widths are about 0.35, 0.45, 0.75, 1.3, 2.0, and 2.6 mm, respectively, for instars 1–6. Larvae attain lengths of about 1.7, 3.5, 6.4, 10.0, 17.2, and 34.2 mm, respectively, during these instars. Young larvae are greenish with a black head, the head turning orangish in the second instar. In the second, but particularly the third instar, the dorsal surface of the body becomes brownish, and lateral white lines begin to form. In the fourth to the sixth instars the head is reddish brown, mottled with white, and the brownish body bears white subdorsal and lateral lines. Elevated spots occur dorsally on the body; they are usually dark and bear spines. The face of the mature larva is also marked with a white inverted “Y” and the epidermis of the larva is rough or granular in texture when examined closely. However, this larva does not feel rough touch, as does corn earworm, Helicoverpa zea (Boddie), because it lacks the microspines found on the similar-appearing corn earworm. In addition to the typical brownish form, its brown dorsal coloration may be replaced with green. In the green form, the dorsal elevated spots are pale rather than dark. Larvae are most active in the morning, late afternoon, and evening, and tend to conceal themselves during the brightest time of the day. Duration of the larval stage tends to be about 14 days during the summer and 30 days during cool weather. Mean development time was determined to be 3.3, 1.7, 1.5, 1.5, 2.0, and 3.7 days for instars 1–6, respectively, when larvae were reared on corn at 25°C (Pitre and Hogg, 1983). However, total larval development time was extended when larvae were fed less suitable hosts: from 13.5 days on corn to 18.9 days on soybean, and 22.3 days on cotton. As development time increased, pupal weights and survival rates decreased. Pupa. Pupation normally takes place in the soil at a depth of 2–8 cm. The larva constructs a loose cocoon, oval in shape and 20–30 mm long, tying together particles of soil with silk. If the soil is too hard, larvae may web

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together leaf debris and other material to form a cocoon on the soil surface. The pupa is reddish brown and measures 14–18 mm long and about 4.5 mm wide. Duration of the pupal stage is about 8–9 days during the summer but reaches 20–30 days during the winter in Florida and Texas. Unlike many noctuids, the pupal stage of fall armyworm cannot withstand protracted periods of cold weather. For example, Wood et al. (1979) studied winter survival of the pupal stage in Florida and found 51% survival in southern Florida, but only 27.5% in central Florida, and 11.6% in northern Florida.

the keys by Whelan (1935), Walkden (1950), and Crumb (1956), and the pictorial keys by Okumura (1962), Rings (1977b), Oliver and Chapin (1981), Capinera (1986), and Stehr (1987). It also was included in a key to armyworms and cutworms in Appendix A. Fall armyworm was included in the keys to moths by Rings (1977a) and Capinera and Schaefer (1983), and pictured by Chapman and Lienk (1981). Heppner (1998) provided an excellent key to the adults of North American Spodoptera. A sex pheromone has been described (Tumlinson et  al., 1986). Culture of this insect was easily accomplished with a bean-based diet (Perkins, 1979).

Damage

FIG. 10.87  Fall armyworm moth. (Photo by L. Buss.)

Adult. The moths, with a wingspan of 32–40 mm, are quite variable in appearance. In the male moth, the forewing is shaded gray and brown, with triangular white spots at the tip and near the center of the wing. The front wings of females are less distinctly marked, ranging from a uniform grayish brown to a fine mottling of gray and brown. The hind wing is iridescent silver white with a narrow dark border in both sexes. Adults are nocturnal and are most active during warm, humid evenings. On the first night of emergence they may feed, but do not mate. They feed on nectar from many plants, usually during early evening. Females may mate repeatedly, but only once per night. After a preoviposition period of 3–4 days, the female normally deposits most of her eggs during the first 4–5  days of life, but some oviposition occurs for up to 3 weeks. The oviposition period tends to be shorter under warm conditions, sometimes as short as 1 day, and longer under cool conditions. Duration of adult life is estimated to average about 10 days (range about 7–21 days). A comprehensive account of the biology of fall armyworm was published by Luginbill (1928), and an informative synopsis by Sparks (1979). Ashley et  al. (1989) presented an annotated bibliography. Fall armyworm was included in many larval identification guides, such as

A highly visible form of damage by larvae is consumption of foliage. Young larvae initially consume leaf tissue from one side, leaving the opposite epidermal layer intact. By the second or third instar, larvae begin to make holes in leaves and eat from the edge of the leaves inward. Feeding in the whorl of corn often produces a characteristic row of perforations in the leaves, though the larvae quickly produce a ragged appearance as they grow and feed. Larval densities are usually reduced to 1–2 per plant when larvae feed in close proximity to one another, due to cannibalistic behavior. Older larvae cause extensive defoliation, often leaving only the ribs and stalks of corn plants, or a ragged, torn appearance. Total leaf consumption by larvae exceeds 100 cm. The proportion of defoliation is estimated at 0.1%, 0.6%, 1.1%, 4.7%, 16.3%, and 77.2% during instars 1–6, respectively. As over threefourths of the defoliation occurs during the last instar, the presence of larvae is easily overlooked and damage occurs suddenly. Marenco et  al. (1992) studied the effects of fall armyworm injury to early vegetative growth of sweet corn in Florida. They reported that the early whorl stage was least sensitive to injury, the mid-whorl stage intermediate, and the late whorl stage was most sensitive to injury. Further, they noted that mean densities of 0.2–0.8 larvae per plant during the late whorl stage could reduce yield by 5%–20%. Larvae also burrow into the growing point (bud, whorl, etc.), destroying the growth potential of plants, or clipping the leaves. In corn, they sometimes burrow into the ear, feeding on kernels in the same manner as corn earworm, Helicoverpa zea. Unlike corn earworm, which tends to feed down through the silk before attacking the kernels at the tip of the ear, fall armyworm feed by burrowing through the husk on the side of the ear. Ear damage is of greater concern to sweet corn growers than foliage feeding.

Management Sampling. Moth populations can be sampled with blacklight and pheromone traps; the latter is more efficient (Starratt and McLeod, 1982). Pheromone traps should be

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suspended at canopy height, preferably in corn during the whorl stage. The type of trap selected for population monitoring can have significant effect on moth catches; plastic canister styles are most desirable based on both the number of moths captured and the ease of trap servicing (Adams et  al., 1989). Such catches are not necessarily good indicators of density but indicate the presence of moths in the area. Once moths are detected, it is advisable to search for eggs and larvae. A search of 20 plants in five locations, or 10 plants in ten locations, is generally considered to be adequate to assess the proportion of plants infested. Hoffmann et al. (1996b) compared fixed sample size with sequential sampling protocols for caterpillar pests of corn in New York and reported substantial savings in time by using this technique for classification of infestation, relative to fixed samples of 100 plants. Sampling to determine larval density often requires large sample sizes, especially when larval densities are low or larvae are young (Mitchell and Fuxa, 1987), so it is not often used. Insecticides. Insecticides are usually applied to sweet corn in the southeastern states to protect against damage by fall armyworm, as frequently as daily during the silking stage. In Florida, fall armyworm is the most important pest of corn. It is often necessary to protect both the early vegetative stages and reproductive stage of corn. As larvae feed deep in the whorl of young corn plants, a high volume of liquid insecticide may be required to obtain adequate penetration. Insecticides may be applied in the irrigation water if it is applied from overhead sprinklers (Sumner et  al., 1991). Granular insecticides are also applied over the young plants because the particles fall deep into the whorl. Baits and ultralow volume techniques are less frequently used. Some resistance to insecticides has been noted, with resistance varying regionally (Harrell et al., 1977; Young, 1979; All et al., 1986). Foster (1989) reported that keeping the plants free of larvae during the vegetative period reduced the number of sprays needed during the silking period. The grower practice of concentrating the sprays at the beginning of the silking period instead of spacing the sprays evenly provided little benefit. Cultural Techniques. The most important cultural practice, employed widely in southern states, is early planting and/or early maturing varieties. Early harvest allows many corn ears to escape the higher armyworm densities that develop later in the season (Mitchell, 1978). Reduced tillage seems to have little effect on fall armyworm populations (All, 1988), though delayed invasion by moths of fields with extensive crop residue has been observed, thus delaying and reducing the need for chemical suppression (Roberts and All, 1993). Host-Plant Resistance. Partial resistance is present in some sweet corn varieties, but it is inadequate for complete protection. Resistance is largely due to nonpreference by larvae, but some antibiosis is present (Wiseman et al., 1981; Wiseman and Widstrom, 1986). Transgenic plant resistance

due to Bt hybids expressing Cry1A(b) endotoxin can dissipate quickly, within a few generations. Biological Control. Although several pathogens have been shown experimentally to reduce the abundance of fall armyworm larvae in corn, only Bacillus thuringiensis now is feasible, and success depends on having the product on the foliage when the larvae first appear. Natural strains of Bacillus thuringiensis tend not to be very potent, and genetically modified strains improve performance (All et al., 1996). An interesting and unusual approach to biological control involves the application of mass-produced parasitoid larvae, Archytas marmoratus (Diptera: Tachinidae). The fly larvae are mechanically extracted from the female flies, suspended in aqueous solution, and sprayed onto plants (Gross Jr. and Johnson, 1985; Gross Jr. et al., 1985).

Glassy Cutworm

Apamea devastator (Brace) (Lepidoptera: Noctuidae)

Natural History Distribution. This native species is found throughout the United States and southern Canada except for the southeastern states. Its range also includes South America. Host Plants. Glassy cutworm is principally a grassfeeding species, and crop damage is most likely done when crops follow sod or are planted into fields heavily infested with grassy weeds. In Oregon, Kamm (1990) reported that bentgrass, Agrostis tenus; ryegrass, Lolium perenne; and wild oats, Avena fatua; were attractive to ovipositing females. Vegetables reported injured by glassy cutworm include beet, bean, cabbage, corn, lettuce, and radish. Among other crops injured are alfalfa, barley, bluegrass, fescue, oat, strawberry, timothy, tobacco, and wheat. Natural Enemies. Several parasitoids of glassy cutworm larvae are known, though in general there is little information about natural population regulation of this species. Kamm (1990) reported that Lissonota montana (Cresson) (Hymenoptera: Ichneumonidae) and Nowickia latianulum (Tothill) (Diptera: Tachinidae) collectively caused 30%–48% parasitism in Oregon. Other wasp parasitoids include Macrocentrus crassipes Muesebeck (Hymenoptera: Braconidae), Pterocormus ambulatorius (Fabricius), and Spilichneumon inconstans (Cresson) (both Hymenoptera: Ichneumonidae). Among other fly parasitoids known from glassy cutworm are Gonia aldrichi Tothill and G. frontosa Say (both Diptera: Tachinidae). Life Cycle and Description. There is one generation annually. The winter is passed in the larval stage, with pupation beginning in May. Moth emergence begins in June, but peak abundance usually is during late July or August. Moths may be present until October, and produce eggs that hatch into the overwintering larval stage.

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Egg. The egg stage of this poorly known species seems to be undescribed. In Minnesota, Knutson (1944) reported that egg laying was completed by the end of August, but Kamm (1990) inferred from adult and larval data that oviposition occurred over several months in Oregon. Duration of the egg stage is about 12–21 days.

Adult. The adult is light gray to brownish gray in general, with extensive amounts of dark brown mottling. A narrow white transverse line is usually present distally on the forewing, with a series of dark triangles located along the inner margin of the transverse line. The hind wing are brownish, and darker distally. The wingspan is 35–45 mm. A sex pheromone produced by females has been identified (Steck et al., 1977, 1980b). The most complete description of this insect is found in Crumb (1929). Knutson (1944) and Kamm (1990) also provided useful observations. A bibliography was published by Rings and Arnold (1974). Keys including the larva of this species were given by Crumb (1929, 1956), Whelan (1935), Rings (1977b), Capinera (1986), and Stehr (1987). It is also included in a key to armyworms and cutworms in Appendix A. The moth was included in pictorial keys by Rings (1977a) and Capinera and Schaefer (1983).

Damage FIG. 10.88  Glassy cutworm larva. (Photo by J. Capinera.)

Larva. The larva feeds entirely below-ground, or at least below the plant litter on the soil surface. The body of this cutworm is largely unpigmented, and many authors noted that this grayish larva resembles a white grub (Coleoptera: Scarabaeidae) in general appearance. The mature larva measures about 35–40 mm long. The head is reddish brown, and measures 4.5 mm wide. A large prothoracic plate is also present, and is reddish brown but with a darker margin. Duration of the larval stage is several months, depending on weather. Pupa. The larva prepares a pupal cell several centimeters below the surface of the soil. The reddish-brown pupa is about 18–20 mm long and 5 mm wide. Duration of pupation is not well-documented but there are reports of 15–60 days in the literature, with the lower value more typical.

The larva lives below-ground, feeding on roots and the base of plant stems. Plants are readily killed by this type of injury, and the first sign of injury usually is wilting plants.

Management Glassy cutworm is not a common pest unless crops are planted into fields that previously had been pasture or grass sod. The problem normally dissipates within 2–3  years after the destruction of the grass. Population monitoring is most easily accomplished with pheromone traps because the other stages are associated with the soil and difficult to detect. Food-based lures, especially acetic acid and 3-methyl-1-butanol, are also attractive (Landolt and Hammond, 2001). Chemical insecticides are useful for prevention of injury but are most effective when placed in the furrow at planting. Baits are not very effective because larvae remain below-ground and have little contact with bait. Mechanical barriers such as metal cans with the top and bottom removed are often recommended for prevention of cutworm damage in home gardens. For glassy cutworm, a burrowing species, the lower edge of the barrier must be sunk well below the soil surface to become an effective deterrent to feeding.

Granulate Cutworm

Agrotis subterranea (Fabricius) (Lepidoptera: Noctuidae)

Natural History

FIG. 10.89  Glassy cutworm moth. (Photo by J. Capinera.)

Distribution. This species is native to the western hemisphere and principally tropical in distribution. Although occasionally found as far north as Nova Scotia and Minnesota,

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it appears not to breed at these latitudes and is not known from the northwestern states. It is commonly found south of the Ohio River and regularly damaging in the southernmost states from Georgia to California. It also occurs in Central and South America and the Caribbean. Host Plants. Granulate cutworm feeds on a wide range of plants. Among vegetables attacked are bean, beet, broccoli, Brussels sprouts, cabbage, carrot, cauliflower, celery, corn, cowpea, eggplant, kale, lettuce, onion, pea, pepper, potato, radish, spinach, sweet potato, tomato, turnip, and watermelon. Other crops reported injured include alfalfa, clover, cotton, lespedeza, peach, peanut, sorghum, soybean, strawberry, tobacco, vetch, and wheat. Some of the weeds observed to support larvae include thorny amaranth, Amaranthus spinosus; cocklebur, Xanthium sp.; dandelion, Taraxacum sp.; passion vine, Passiflora incarnata; plantain, Plantago sp.; and shepherdspurse, Capsella bursa-pastoris. Natural Enemies. Considering the importance of this cutworm in southern states, surprisingly little is known about natural enemies. Among the wasps known to parasitize granulate cutworm are Apanteles griffini Viereck, Chelonus insularis Cresson, Meteorus laeventris (Wesmael), M. laphygmae Viereck, Microgasterfeltiae Meusebeck, Zele mellea (Cresson) (all Hymenoptera: Braconidae), Campoletis flavicincta (Ashmead) and Simphion merdarius (Gravenhorst) (both Hymenoptera: Ichneumonidae). Fly parasitoids known from this cutworm include Bonnetia comta (Fallén), Gonia crassicornis (Fabricius), G. longipulvilli Tothill, Lespesia archippovora (Riley), and Spallanzania hebes (Fallén) (all Diptera: Tachinidae). A fungus (microsporidian) disease was reported from Florida (Adlerz, 1975) and a granulosis virus is known (Hamm and Lynch, 1982), but the importance of natural pathogens is uncertain. An ectoparasitic nematode, Noctuidonema guyanense, parasitizes the adult and is thought to be debilitating (Simmons and Rogers, 1996). Life Cycle and Description. Granulate cutworm is active continuously in the south; adults, eggs and larvae have been collected during all months in Louisiana. Nevertheless, there seems to be a seasonality to reproduction, as unmated females are found mostly from May to -November. Total abundance similarly is greatest in June-November. In Tennessee, three complete generations are reported, with overwintering insects emerging in March. In addition to egg production about March, peaks in egg production occur in May, July, and September. The pupae from the September generation overwinter. The complete life cycle requires 50–70 days. Egg. The eggs are deposited singly or in small clusters on the upper surface of foliage. Females produce about

800–1600 eggs. They are hemispherical, with 36–40 narrow ridges radiating from the apex, and measure 0.60–0.71 mm in diameter and about 0.50 mm in height. Initially, they are white but darken with age. Normally they hatch in 3–5 days.

FIG. 10.90  Granulate cutworm larva. (Photo by L. Buss.)

Larva. Young larvae initially remain on the foliage during both day and night, but after a few days, they begin to hide beneath plant debris or soil during the daylight hours, feeding only at night. The larva buries itself very shallowly, even remaining partially exposed during the day. The number of instars varies from 5 to 7, but six instars are most common. Mean (range) head capsule widths are 0.31 (0.30–0.32), 0.48 (0.45–0.50), 0.81 (0.73–0.86), 1.31 (1.17–1.57), 1.98 (1.62–2.26), and 2.93 (2.70–3.19) mm for instars 1–6, respectively. Mean (range) duration of the instars is 3.4 (3–5), 2.5 (2–4), 3.1 (3–4), 3.1 (2–5), 4.8 (3–7), and 7.7 (5–10) days, respectively. Total larval development time is about 25  days for larvae with six instars, but 22  days for five-instar larvae and 32  days for seveninstar larvae. The body length measures about 2.0–3.5, 5–6, 12, 17, 22, and 30–37 mm long during instars 1–6, respectively. Larvae with six instars consume about 150 cm2 of foliage but the longer-lived larvae that undergo seven instars consume considerably more, up to 240 cm2. This cutworm is grayish to reddish brown, with each abdominal segment bearing dull yellowish oblique marks subdorsally. A weak gray line occurs laterally below the spiracles, accompanied by spots of white or yellow. The head is yellowish to brownish. Pupa. Pupation occurs in the soil, usually at a depth of 3–12 cm. The pupae are dark brown or mahogany, and measure 15–21 mm long and 5–6 mm wide. Duration of the pupal stage is 10–20 days.

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delivered as liquid, granule, or bait formulation. Baits, particularly bran-based, seem particularly effective (Morgan and French, 1971). Bacillus thuringiensis is not usually recommended for cutworms. Many cutworm species have a great affinity for weedy fields, but granulate cutworm seems to lack this association. Mechanical barriers can provide some protection from dispersing larvae to seedlings in the home garden. However, moths often are active during the growing season and easily circumvent such barriers. Therefore, it may also be necessary to use netting or row cover material to deny access to plants by ovipositing moths. FIG. 10.91  Granulate cutworm moth. (Photo by L. Buss.)

Adult. Moths begin mating about 1 day after emergence, and peak oviposition occurs 2–3 nights after mating (Cline and Habeck, 1977). The longevity of adults is 10–20  days, averaging about 14  days. The moth is medium in size, with a wingspan of 31–43 mm. The color of the forewing varies considerably in its shades of brown and gray, but it is often yellowish brown and distinctly lighter distally. The forewing bears distinct bean-shaped and round spots centrally, and these spots are linked by a small but sharply defined black bar. The hind wings are white, but dusky marginally and along the veins. Biology was described by Jones (1918b), and Snow and Callahan (1968), with the most complete morphological description by Crumb (1929). Culture techniques were described by Lee and Bass (1969). Keys including the larva of this species were given by Crumb (1929, 1956), Whelan (1935), Okumura (1962), Oliver and Chapin (1981), and Stehr (1987), and in a key to armyworms and cutworms in Appendix A. The moth was included in a pictorial key by Capinera and Schaefer (1983).

Damage This is the most important cutworm pest of vegetables in the Gulf Coast region, and also quite important in California. It damages seedlings by cutting off the stem at the soil surface, older plants by climbing and feeding on foliage, and injures such plants as tomato, watermelon, and eggplant by feeding on, or burrowing into, the fruit. Due to its surface-feeding behavior, granulate cutworm is sometimes a major component of the “rindworm” complex affecting cucurbit fruit. This type of damage usually occurs when fruit is in contact with soil, a common habitat of the larva. Young larvae, through about the second instar, feed on the lower leaf surface and skeletonize leaves. Thereafter, they consume entire leaves.

Management Moth populations can be monitored with blacklight traps. Larvae can be controlled by the application of insecticides

Old World Bollworm

Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae)

Natural History Distribution. Helicoverpa armigera has long been known from Africa, Asia, and Australia (Tay et al., 2013). More recently, it has become a pest in Europe, though mostly in southern Europe and in greenhouses. In cooler environments, they may occur but are unlikely to become very abundant. Recently, H. armigera established in South America, was recognized in Brazil only in 2013. It is frequently detected at ports of entry into the United States, so there is a great likelihood of it becoming established and a pest in warm-weather states of the southeastern and southwestern United States, and possibly in cooler areas as well. Host Plants. The host range of H. armigera is quite extensive, including over 180 plant hosts in 45 plant families. Among the vegetable crops consumed are onion, garlic, leek, okra, cabbage, broccoli, Brussels sprouts, cauliflower, bell pepper, squash, chickpea, cowpea, green bean, tomato, eggplant, potato, asparagus, and corn. Field crops are also commonly consumed, including peanut, oat, pigeon pea, safflower, millet, soybean, cotton, sunflower, barley, flax, alfalfa, tobacco, sorghum, and wheat. Flowers and tree fruit can also be attacked. Not all hosts are equally preferred, of course. For example, Jallow and Zalucki (1996) found that corn, sorghum, and tobacco were preferred over cotton. Similarly, Firempong and Zalucki (1990) found tobacco, corn, and sunflower to be preferred; soybean, cotton, and alfalfa were intermediate; and cabbage, pigweed, and flax (linseed) were least preferred. In another study, pigeon pea was most preferred by adults for oviposition (and suitable for larval growth), whereas sowthistle (Sonchus oleraceus L.) [Asteraceae] and cotton were not as attractive for oviposition but equally suitable as pigeon pea for growth, and mungbean was neither attractive for oviposition nor suitable for larvae (Rajapakse and Walter, 2007). Liu et al. (2004) compared development from the egg to the adult stage when

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reared at 27°C and fed one of several crops, and reported that mean total development times were similar: 29.7, 26.6, 27.9, 35.1, 33.8, and 32.9 days when fed cotton, corn, bean, tomato, pepper, and tobacco, respectively. However, survival was much better on cotton than on vegetable crops. Natural Enemies. Natural enemies are numerous but vary in importance in different crops and different locations around the world. Among important predators are minute pirate bugs (Anthocoridae), green lacewings (Chrysopidae), lady beetles (Coccinellidae), and ants (Formicidae). Among important groups of parasitoids are Trichogrammatidae, Braconidae, Ichneumonidae, and Tachinidae. Fungi, bacteria, Helicoverpa armigera nuclear polyhedrois virus, and nematodes also can be important in some environments. In general, however, quantitative data on population regulation by natural enemies are lacking. Natural enemies have been introduced to new parts of the world where H. armigera has become established, but the effects of the introductions have not had too great a beneficial effect. Life Cycle and Description. The number of generations is variable, and because they overlap, it is difficult to distinguish the number in most areas. The number of annual generations normally is 2–5, but under ideal (tropical) conditions 11 generations can be attained. Diapause is facultative, but if present it occurs in the pupal stage. After overwintering in the pupal stage, the moths oviposit on plants, usually on blossoms or leaves located near blossoms or fruit. Pubescent vegetation is preferred for oviposition. There are 5–7 instars, with 6 instars most frequent. Then, larvae drop from the plant to pupate in the soil. Diapause is induced by short day lengths (11–14 h per day). Hot, dry conditions have also been reported to induce summer diapause, though this occurs infrequently. Egg. Eggs are deposited 1–4 days after mating. The eggs are yellowish white initially, but turn into brown before hatching. They are hemispherical, rounded above but with a flat base. They measure about 0.4–0.6 mm in diameter. Eggs require 2–4 days for hatching under warm (summer) conditions and about 7  days under cooler (spring and autumn) conditions. Females produce on average, about 750 eggs, but some produce over 2000 eggs. Larva. The number of larval instars varies from 5 to 7, but typically is six. The appearance of the larvae is quite variable, and affected by both diet and maturity. They tend to become darker with age, initially being yellowish but with a dark head, prothoracic and supra-anal shield, spiracles, and the base of the tubercles. They transitioned to greenish or brownish caterpillars with brown lateral stripes and a dorsal stripe. They eventually attain a length of about 3.5–4 cm and a mean weight of 377 mg. The larvae are very

difficult to distinguish from corn earworm, Helicoverpa zea (Boddie) (Lepidoptera: Noctuidae), even for experts. Like other ­holometabous insects, the instars can be distinguished fairly reliably based on maximum head capsule widths. The mean, standard error, and range of the head capsule widths were reported to be 0.298 ± 0.007, 0.2–0.4 mm for instar 1; 0.495 ± 0.007, 0.4–0.7 mm for instar 2; 0.818 ± 0.015, 0.6–1.2 mm for instar 3; 1.424 ± 0.055, 0.9–3.3 for instar 4; 2.017 ± 0.062, 1.4–3.5 for instar 5; and 2.992 ± 0.044, 1.8–4.5 mm for instar 6, respectively (Stavridis et al., 2003–2004). Development time varies with temperature and diet, though the pattern across the different developmental periods is similar. For example, when Liu et al. (2004) reared H. armigera on cotton at 27°C, the mean number of days required was 3.0, 1.6, 2.1, 2.3, 2.6, 2.9, 3.5, 4.8, 2.5, and 10.1 for the egg stage, instars 1–7, prepupa, and pupal stage, respectively. For a less suitable host (tomato) reared in the same manner, the mean duration was 3.0, 3.1, 2.2, 2.6, 2.5, 2.8, 4.2, 5.6, 2.3, and 9.3  days for the egg stage, instars 1–7, prepupa, and pupal stage, respectively. Although development time was not much affected by larval diet, pupal weight was greatly affected. Thus, in Liu et al. (2004) study, mean female pupal weight was 280 mg on a diet of cotton, but only 167 mg on tomato. This potentially translates into marked differences in fecundity. Pupa. The pupae are brownish, to dark brown in color. They are 14–22 mm long and 4.5–6.5 mm wide. The anterior end of the pupa is broadly rounded, whereas the posterior end tapers, and terminates with two parallel spines. They are found in the soil, usually at a depth of 3–18 cm, though this varies with the hardness of the soil. Duration of the nondiapausing pupae is about 6 days at 35°C but about 30 days at 15°C. When diapausing, the pupal stage can persist for several months. Adult. The adult emerges from the pupal cell in the soil during the evening hours, but usually before midnight. Moths feed on nectar. Mating occurs about 4  days after emergence from the soil. Like most other noctuids, the moth is stout bodied. The wing span is 3.0–4.0 cm. The body is 12–20 mm long. The overall color of males varies greatly, from yellowish to greenish or brown, whereas the female is usually orange brown. The forewings bear a dark brown or black kidney-shaped (reniform) spot near the center, and a weak to strong dark band crossing the wing in the distal third. The hind wings are creamy white with a broad dark band at the distal margin. As with the larval stage, identification is difficult, and normally requires dissection of the adult male genitalia or molecular assessment for accurate species determination. Details on differentiation of H. armigera and H. zea are provided by Pogue (2004).

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Damage Wherever it occurs, H. armigera is a pest of major importance for several reasons: it is highly polyphagous, attacking many economic plants; it can diapause during inclement weather, or continue to flourish if the weather is favorable; it has a high rate of reproduction; and it is resistant to many insecticides. Also, it is quite mobile (though a facultative migrant), dispersing long distances when weather or food conditions are unsuitable, but otherwise not flying far. This species attacks most stages of growth, including the vegetative, flowering, and fruiting stages. Like most caterpillars, the larvae of H. armigera consume a portion of the leaves, and can completely defoliate small plants. Like its wellknown relative Helicoverpa zea (Boddie), H. armigera also bores into blossoms and fruit or pods. The damaged fruit often drop from the plant.

hosts such as maize (corn) and sorghum, though attractive to the insects, have not successfully reduced the abundance of the insects. Similarly, although tillage can destroy pupae in the soil, the effects are inadequate for protection. A few varieties of major crops have been produced that suppress insect growth and survival.

Pale Western Cutworm

Agrotis orthogonia Morrison (Lepidoptera: Noctuidae)

Natural History

Distribution. This native species is found in semiarid regions of the western United States and Canada. Its distribution is largely restricted to the western edge of the Great Plains and eastern portions of the Rocky Mountains, including southern Alberta and Saskatchewan, most of Montana, Management Wyoming, Colorado and Utah, and northern New Mexico. Sampling. Because this species is capable of long-­ It also occurs in portions of adjoining states and has been distance dispersal (several hundred kilometers), it can some- particularly troublesome in western Kansas. times attain areas where it does not normally occur. Thus Host Plants. Larvae generally feed on grasses and grain vigilance is necessary so as not to be surprised by infestation. crops, and also on some weeds. However, they occasionally A sex pheromone is known and is commercially avail- have been known to damage vegetables, including bean, able; it is used to bait several forms of traps for popula- beet, carrot, corn, onion, potato, and tomato. Most comtion monitoring. Funnel traps and Texas (Hartstack) traps monly damaged are the small grain crops such as wheat, are superior to sticky traps and water-pan traps. Traps barley, rye, oats, and millet, but other field crops such as normally are suspended 1.5–1.8 m above the soil surface. alfalfa, flax, Sudan grass, sugarbeet, and sweet clover may Unfortunately, some other species (especially Spodoptera be fed upon. Among weeds known to be consumed are sunspp.) can be attracted to H. armigera traps, thus requiring flower, Helianthus annuus; tumble mustard, Sisymbrium aladditional sorting. tissimum; Russian thistle, Salsola kali; wild lettuce, Lactuca The presence and abundance of insects in a crop can be scariola, mallow, Malva sp.; and dandelion, Taraxacum spp. confirmed by visual examination. The underside (abaxial) The adults feed on nectar from flowers, preferring goldof the leaf is a preferred oviposition site. Eggs hatch quickly, enrod, Solidago spp.; sunflower, Helianthus spp.; and rabbit so examination of plant organs is advisable for signs of bor- brush, Chrysothamnus spp. In the absence of the preferred ing, such as holes and extruded frass. Insects are usually nectar sources other flowers are used, including snakeweed, found in the upper one-third of the plant. Gutierresia sp.; Canada thistle, Cirsium arvense; fleabane, Insecticides. Growers often depend on insecticides for Erigeron spp.; and Russian thistle, Salsola kali. control of H. armigera. However, steps should be taken to Natural Enemies. Natural enemies are believed to play reduce the rate of development of insecticide resistance. In an important role in the occurrence of this species, but it Australia, for example, dependence on insecticides led to is the interaction of weather and natural enemies that is the rapid evolution of resistance to DDT, organophosphates, critical. Larvae of this cutworm normally spend most of and pyrethroids in cotton, necessitating rotation of insecti- their time below-ground. However, wet weather in the cides (Downes et al., 2016). Use of genetically modified Bt spring months causes larvae to move to the soil surface, cotton was also implemented, but this approach is not an where they can be attacked by parasitoids and predators. option for many food crops. The proportion of the population lost to natural enemies Cultural Practices. The H. armigera population tends varies from about 20% to 70%. Among the wasps known to increase over the course of the growing season, so early to parasitize pale western cutworm are Meteorus levivenplanting and cultivation of early maturing crops can be ad- tris Wesmael, Chelonus sp., Zele sp. (all Hymenoptera: vantageous by reducing exposure to high pest population Braconidae); Apanteles griffini Viereck, Paniscus sp. (both densities later in the season. Not all crops are equally suscep- Hymenoptera: Ichneumonidae); and Copidosoma bakeri tible, so planting less susceptible crops is commonly prac- (Howard) (Hymenoptera: Chalcididae). Other parasitticed. Use of Bt cotton can greatly reduce damage to cotton, oids include Bonnetia comta (Fallén), Mericia sp., Gonia and also to other nearby susceptible crops. Diversionary aldrichi Tothill, G. longiforceps Tothill, G. longipulvilli

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Tothill, Peleteria texensis Curran, and Periscepsia rohweri (Townsend) (all Diptera: Tachinidae), Anthrax molitor Lowen, Villa alternata (Say), V. willistoni (Coquillett), and Poecilanthrax sackenii (Coquillett) (all Diptera Bombyliidae). Numerous avian and insect predators have been observed to feed on cutworm larvae, among the insects are leaf-footed bugs (Hemiptera: Coreidae), assassin bugs (Hemiptera: Reduviidae), and ambush bugs (Hemiptera: Phymatidae), ground beetles (Coleoptera: Carabidae), and predatory wasps (Hymenoptera: Sphecidae and others). The role of pathogens is uncertain, fungi and viruses seem to be unimportant, but bacterial diseases are sometimes suggested to be a significant mortality factor. Weather. The abundance of pale western cutworm is directly related to precipitation patterns. Outbreaks of pale western cutworm rarely occur in areas with more than 12 cm of precipitation during the period of May-July, when pale western cutworm is in the larval stage. Within the semiarid area of western North America inhabited by this cutworm, spring periods with fewer than 10 precipitation events exceeding 6.4 mm are followed by increases in cutworm number during the following year. Similarly, spring periods with more than 15 such precipitation events are followed by population decreases. The effect of the precipitation is to drive the larvae to the surface of the soil, where they are susceptible to attack by predators and parasitoids. Higher moisture levels may also favor the spread of disease among the insects, but this is less certain. Life Cycle and Description. There is only a single generation per year throughout the range of this insect. Eggs are laid in the autumn and hatch in the winter or early spring. Larvae feed until early June and then enter a quiescent prepupal period that may last for 40–50 days if the weather is warm. Pupation occurs in July or August, with moths common in late August-October. Not surprisingly, the active period of this species is shorter in the north, approximately April-September in Alberta, whereas in New Mexico the insects are active from February to October. Egg. Eggs are deposited in the soil at a depth of 6–12 mm, apparently singly or in small clusters. They are white, turn yellowish gray and then slightly bluish as the embryo matures. In shape the egg is a slightly flattened sphere, measuring about 1 mm in diameter and 0.8 mm in height. The egg bears 27–32 ridges radiating from the apex. Incubation requires 30–50 days in the field and embryos require a cold period before hatching. Under laboratory conditions, embryo development requires 11, 14, 21, and 33 days at 30°C, 25°C, 20°C, and 15°C, respectively. They must have contact with moisture or high humidity in order to hatch. As noted above, hatching occurs early in the spring.

FIG. 10.92  Pale western cutworm larva. (Photo by J. Capinera.)

Larva. The larvae feed below-ground for their entire life. They normally display 6–8 instars. Jacobson (1971) gave mean development times of 8, 6, 7, 7, 8, and 14 days (excluding the prepupal part of the terminal instar of another 13 days) for instars 1–6, respectively, when reared at 20°C. In contrast, Parker et  al. (1921) gave mean development times of 11.2, 8.0, 9.4, 9.8, 11.3, 14.4, 22.6, and 29.6 days (excluding the prepupal period) for instars 1–8, respectively, under unspecified insectary conditions. Thus, larval development time in the latter study, which averaged 118  days, was more than twice in the former study, where development required only 50 days. The difference is even greater if development times at warmer temperature are compared. Jacobson (1971) gave mean larval development times of only 29 and 24  days at 25°C and 30°C, respectively. Head capsule widths are 0.27–0.31, 0.35–0.45, 0.48–0.67, 0.70–0.98, 1.00–1.50, 1.60–2.20, 2.30–2.80, and 2.90–4.00 mm, respectively, for instars 1–8 (Sutter et  al., 1972). Mean body lengths are reported to be 1.4, 4.0, 5.9, 9.3, 15.9, 24.6, 31.1, and 36.6 mm, respectively. The larva is gray or bluish gray, and is largely free from distinctive markings. The head capsule, which is yellowish brown, bears two dark vertical bars. The larva attains a length of about 35–40 mm at maturity. At maturity the larva digs deeper into the soil, usually 5–15 cm, and forms an earthen cell with the aid of salivary secretions. While in this cell, the prepupa shrivels to a length of about 20–25 mm and its color changes to yellowish white. A period of inactivity follows, 30– 75 days in duration, that is quite long compared to similar species, and likely is an adaptation allowing the insect to escape the heat and dryness of the summer in its hostile environment. Pupa. Pupation occurs within the cell formed by the larva. The pupa is yellowish initially, turning brown with time. The pupa measures about 13–19 mm long. Duration of the pupal period is 20–40 days.

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Management

FIG. 10.93  Pale western cutworm moth. (Photo by J. Capinera.)

Adult. The adult is an attractive moth, gray with yellowish and brownish spots on the forewing, and white on some of the veins. The hind wings are whitish, but darker distally. The body is robust and clothed with long scales. The wingspan of the moth is 25–40 mm. Although the moths are considered to be principally nocturnal, females fly and oviposit in the late afternoon, and both sexes begin feeding at flowers about sunset. Females select loose soil for oviposition, avoiding hard or crusted surfaces. Fecundity is not well documented, but apparently females can produce 300–400 eggs. Adult activity usually ceases by midnight, principally because it becomes cool, but there are reports of mating occurring at this time. Length of adult life in the field is uncertain but appears to be 7–14 days. The biology of pale western cutworm was described by Parker et al. (1921), Cook (1930), Sorenson and Thornley (1941), and Jacobson (1971). Rearing was described by Sutter et  al. (1972). A sex pheromone was described by Struble and Swailes (1978). Keys to the larval stage were found in Whelan (1935), Walkden (1950), Capinera (1986), Stehr (1987), and in a key to armyworms and cutworms in Appendix A. Adults were included in a key by Capinera and Schaefer (1983).

Damage The larvae feed below-ground near the growing point of plants. Young larvae are not large enough to completely sever the plant, but this is accomplished routinely once the third instar is attained. This feeding usually results in the death of the plant. In grain crops, entire fields may be killed. The larvae are reported to move underground from plant to plant, often feeding on just the small below-ground section of each seedling. They rarely move above the soil and are not known to disperse long distances in an “armyworm” fashion in search of food.

Sampling. The eggs and small larvae are very difficult to detect. Large larvae can be recovered from soil, but their densities are usually assessed indirectly from plant damage. Adult densities can be monitored easily with light traps or pheromone traps. Damaging populations are predicted on the basis of the pattern of precipitation during the larval stage. The days of the larval stage when at least 6.4 mm of precipitation ­occurs—termed “wet days”—are tabulated, and if they are less than 10, cutworm densities are expected to increase during the next year. In contrast, if the number of “wet days” exceeds 15, then cutworm populations are expected to decrease (Seamans, 1935). Two successive years of dry weather are required to cause high and damaging population densities. Insecticides. Persistent insecticides applied to the foliage or soil in a liquid formulation provide good control of pale western cutworm. Larval mortality in fields treated with insecticide is not rapid, often requiring several days. The use of persistent insecticides is very important because larvae are inactive and remain below-ground for 3–5 days during molting cycles. Thus, if insecticides are to be effective, they must be persistent enough to remain until the larvae resume activity and come into contact with insecticide (Byers et al., 1992; Hill et al., 1992). Bait formulations generally provide only partial control because larvae generally feed below the soil surface and therefore have little contact with baits (DePew, 1980; McDonald, 1981a). Bacillus thuringiensis is not usually recommended for control of cutworms. Cultural Practices. Cultural methods of management have been developed for grain production systems, but they may have applicability for other cropping systems. Moths infrequently deposit eggs in soil that has a crust, apparently because females cannot penetrate hard soil with the tip of their abdomen. Therefore, a widely practiced technique is to allow a crust to form on the soil in the autumn, before oviposition. Another common practice is to till the soil 10–14 days before planting. This destroys weeds and other alternate hosts on which larvae may be feeding and causes them to starve before the new crop germinates. A practice that is often suitable for vegetable production is to increase irrigation frequency during periods of cutworm abundance; frequent irrigation in the spring is deleterious to larval survival. Barriers are sometimes used to reduce access by cutworms to plants grown in the home garden. Metal or waxedpaper containers with both the top and bottom removed can be placed around the plant stem to deter consumption. The barrier should be extended below the soil surface because larvae may burrow below the soil surface. For pale western cutworm, the requirement that the lower edge of the ­barrier

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is deeply recessed in the soil is especially important, because larvae burrow actively in the soil, rarely coming to the soil surface except during wet weather.

Plantain Looper

Autographa precationis (Guenée) (Lepidoptera: Noctuidae)

Natural History Distribution. This native species is eastern in distribution. It is found widely in the United States east of the Great Plains, though it is infrequent in the southernmost states. Occasionally, plantain looper is found as far west as Kansas, Nebraska, and Wyoming. Similarly, in Canada, it is known from Nova Scotia to Manitoba. Host Plants. This insect sometimes feeds on such vegetables as bean, cabbage, and parsnip, but is more commonly associated with such weeds as burdock, Arctium lappa; common morningglory, Ipomoea purpurea; dandelion, Taraxacum officinale; lambsquarters, Chenopodium album; plantain, Plantago spp.; thistle, Carduus sp. and Cirsium sp.; wild lettuce, Lactuca sp.; and wild sunflower, Helianthus sp. It has also been found feeding on hollyhock. Natural Enemies. Natural enemies of this insect are unknown. Life Cycle and Description. Larvae are the overwintering stage, and apparently, there are two to three generations annually (Knutson, 1944; Chapman and Lienk, 1981). Moths are present in New York from May until November with a reduction in abundance near the end of June, which probably signifies the completion of the first generation (Chapman and Lienk, 1981). A complete life cycle requires about 30–37 days for completion (Khalsa et al., 1979). Egg. The egg stage of this little-known insect seems to be undescribed. Larva. The number of instars usually is 6, but occasionally 7 are observed. Head capsule widths are about 0.3, 0.5, 0.7, 1.0, 1.3, 2.0, and 2.7 mm for instars 1–7, respectively. The total duration of the larval stage is 17–20 days, with the length of individual instars being about 3.0, 3.0, 2.0, 3.5, 2.8, and 2.3 days, respectively, for instars 1–6. The mature larva is green and, like most of the other loopers, bears three white lines on each side of the back and a white lateral line slightly above the lateral spiracles. The white lines are pale in overwintering larvae but distinct in summer larvae. There tends to be dark shading above the lateral line. The body appears to lack microspines, but there is at least a subdorsal strip of these minute structures. The thoracic legs generally are black. The head bears a broad black line on each side; sometimes the lines are broad and cover the entire head. The larva measures about 30 mm long at maturity. This insect closely resembles alfalfa looper, Autographa ­californica,

and bilobed looper, Megalographa biloba, in appearance, and is reliably distinguished by examination of the larval mandibles (see Crumb, 1956, or Eichlin and Cunningham, 1978 for a key). However, the geographic range is generally adequate for differentiation from alfalfa looper, and if microspines are readily apparent there is great likelihood that it is bilobed looper. Pupa. Duration of the pupal stage is about 6–7 days. Adult. The moth is similar to alfalfa looper in general appearance, with the forewing bearing a silvery-white central spot shaped roughly like a “dog leg.” However, the “foot” is weakly connected to the “leg,” or even disconnected, in plantain looper. The background color of the forewing varies from gray to dark-brown. The hind wing is light brown basally and light to dark brown distally. The wingspan of this moth is about 35 mm. Mating typically occurs about 2 days after emergence from the pupa. Females oviposit over a period of about 14 days and produce over 2000 eggs per female. Total adult longevity is estimated at about 19 days. Key elements of the biology of this insect can be found in Khalsa et  al. (1979), with additional information and keys in Crumb (1956), Eichlin and Cunningham (1978), Chapman and Lienk (1981), and Stehr (1987). A key to some common vegetable-feeding loopers, including plantain looper, can be found in Appendix A.

Damage This insect feeds on the underside of leaves during the first three instars, causing a skeletonizing effect. Thereafter, larvae eat large, irregular holes in leaves. Khalsa et al. (1979) demonstrated that this insect consumed almost as much as the more damaging soybean looper, Chrysodeixis includens. However, few plantain loopers are usually found attacking crops, so ovipositional preference keeps this insect from becoming a serious pest.

Management Moths of this species are attracted to light traps. They also can be captured in traps baited with phenylacetaldehyde (Cantelo et al., 1982). This insect normally does not warrant suppression. Foliar insecticides are effective if needed.

Potato Stem Borer

Hydraecia micacea (Esper)

Hop Vine Borer

Hydraecia immanis (Guenée) (Lepidoptera: Noctuidae)

Natural History Distribution. Potato stem borer is native to Europe, northern Asia, and Japan. It was first found in North America

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in 1905 in Nova Scotia, but has since spread through eastern Canada west to Manitoba, and is occasionally damaging throughout this geographic range. In the United States, potato stem borer is known from the northeastern and midwestern states. Hop vine borer, in contrast, is a native species, found across southern Canada and northern United States from coast to coast. Whereas potato stem borer tends to be more common in Canada, hop vine borer occurs frequently in the United States. Both species assumed greater importance as crop pests starting in the 1970s and 1980s, though the cause is unknown. Host Plants. Potato stem borer is polyphagous, but it is known principally as a pest of potato, corn, and rhubarb. Crops attacked include barley, corn, hops, onion, potato, raspberry, rhubarb, strawberry, sugarbeet, tomato, and wheat. Several grasses can serve as hosts, including bromegrass, Bromus sp.; barnyardgrass, Echinochloa crusgalli; reed canary grass, Phalaris arundinacea; orchardgrass, Dactylis glomerata; and quackgrass, Agropyron repens. Hemp nettle, Galeopsis tetrahit; curly dock, Rumex crispus; and possibly other swamp or marsh dwelling plants are suitable broadleaf hosts (Giebink et al., 1992, 1999). The major hosts of hop vine borer are hops and native perennial grasses, and this insect is a serious pest principally in areas where cultivated or wild hops grow. Increasingly, however, it has become a pest of corn. Among weed hosts preferred by ovipositing females are foxtail, Setaria spp.; quackgrass, A. repens; wild proso millet, Panicum milliaceum; and to a lesser extent large crabgrass, Digitaria sanguinalis; barnyardgrass, Echinochloa crusgalli; and fall panicum, Panicum dichotomiflorum. Larvae can survive on curly dock, but growth is poor (Giebink et al., 1992, 1999). Natural Enemies. Several native parasitoids of potato stem borer are known. Egg parasitoids include Telenomus sp. (Hymenoptera: Scelionidae), Trichogramma retorridum (Girault) (Hymenoptera: Tichogrammatidae), and Centrodora sp. (Hymenoptera: Mymaridae). Parasitoids reared from larvae include Lydella radicus Townsend (Diptera: Tachinidae), Diadegma sp., Campoletis sp., Ectopimorpha luperinae Cushman, and Glypta sp. (all Hymenoptera: Ichneumonidae). Reared from pupae are Therion sp. and Pterocormus sp. (both Hymenoptera: Ichneumonidae), but these species likely attack the larval stage. The most effective parasitoid in Ontario is Lydella radicus, and parasitism levels of 25%–60% have been reported (West et  al., 1983), but the other species seem to contribute little to the overall level of parasitism. Additional parasitoids have been imported from Europe and released in Canada, including Macrocentrus blandus Eady and Clark (Hymenoptera: Braconidae) and Lydella stabulans Fallén (Diptera: Tachinidae). Natural enemies of hop vine borer are less well known, but several predators and parasitoids were identified in New York. Among the ground beetle predators are

Calosoma calidum Fabricius, Harpalus pensylvanicus De Geer, Pterostichus lucublandus Say, Pterostichus stygicus Say, and Amara impuncticollis Say (all Coleoptera: Carabidae). Hawley (1918) suggested that the ground beetles consumed the egg, larval, and pupal stages of hop vine borer. Parasitoids identified from New York included Microplitis gortynae Riley, Aenoplex sp., and Synaldis sp. (all Hymenoptera: Braconidae), and Lespesia frenchii Williston (Diptera: Tachinidae). Life Cycle and Description. Potato stem borer and hop vine borer are similar in biology and appearance. They display one generation per year, with the egg serving as the overwintering stage. Eggs hatch in April-May, pupation typically occurs in July, and adults are found from late July to September. Egg. The eggs are laid in two to three parallel rows between the stem and leaf sheath of grasses with a split leaf sheath. The number of eggs ranges from about 30 to 300 per clutch (Levine, 1986a). In shape, the eggs are a flattened sphere. They measure 0.64–0.82 mm in diameter and 0.31– 0.51 mm in height. The edges of the eggs are marked with about 100 narrow, branching ridges, but ridges are absent from the center of the egg. They are white when first deposited, but turn reddish brown, and then black just before hatching. The egg cluster is covered with a transparent film. They are often deposited in August-September and hatch in April-May—a duration of about 8 months. Larva. There are six instars. Mean (range) of head capsule widths is about 0.33 (0.30–0.38), 0.60 (0.41–0.76), 0.97 (0.89–1.11), 1.43 (1.25–1.75), 2.24 (2.00–2.50), and 3.50 (2.75–4.06) mm, respectively, for instars 1–6. Larval development time is about 30–70 days when reared at 24– 27°C. Deedat et al. (1983), for example, gave mean instarspecific development times of potato stem borer as 4.9, 3.8, 3.8, 3.2, 4.4, and 11.5 for instars 1–6, respectively. In the field, however, larval periods are reported to be longer, about 6–9  weeks in Wisconsin (Giebink et  al., 1984) and 9–12 weeks in New York (Hawley, 1918). The developmental threshold of hop vine borer is about 5°C, slightly lower than that of potato stem borer, which is about 7°C (Giebink et al., 1985). The larva is whitish, but the early instars bear rose to purplish bands on the thoracic and abdominal segments. Thereafter the bands fade in hop vine borer, whereas potato stem borer larvae bands tend the remain evident. The head capsule is yellow in potato stem borer but brown in hop vine borer. The larvae measure about 2–3 mm at hatch but eventually attain a length of 30–50 mm. Pupa. Pupation occurs in the soil at a depth of 24 cm. Pupation normally occurs in July, with a duration of 4–6 weeks in the field in New York, but only 20–30 days under laboratory conditions. The pupa is dark brown and measures 15–28 mm long. The tip of the abdomen bears two short spines.

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Adult. The adult is light brown, with pinkish or greenish tints. In hop vine borer the front wings bear broad light-colored transverse lines bordered with brown and ­ some darker gray to olive-brown shaded areas. In potato stem borer the overall color and pattern are similar, but the transverse lines on the front wings are narrow and dark. The hind wings are grayish with dark veins. The wingspan is 40–50 mm. The moths were pictured by Rings and Metzler (1982). Moths emerge in August-September, mate and begin egg production within a few days of emergence. Fecundity is estimated at 400–1200 eggs. Life history of potato stem borer was given by Deedat et al. (1983). Life history of hop vine borer was given by Hawley (1918) and Giebink et  al. (1984). Developmental biology of both species was described by Giebink et  al. (1985). Larvae are included in a larval key in Stehr (1987). Culture of Hydraecia spp. on artificial diet was described by West et al. (1985) and Giebink et al. (1985). A sex pheromone has been described for potato stem borer (Teal et al., 1983; Burns and Teal, 1989).

Damage Larvae feed initially on grasses growing as weeds among or near crop plants, then usually switch to larger grasses such as corn, or broadleaf plants such as hops, potato, or curly dock. On perennial grasses, the feeding occurs aboveground, but after the feeding switch at about the fourth instar larvae tunnel below-ground into the base of the stem and roots. Some of the perennial grasses and other plants may have sizable underground rhizomes, roots, or stems that allow complete larval development, but this aspect of larval biology is poorly known. Damage to corn by potato stem borer was described by Deedat and Ellis (1983), who reported that over 90% of seedling corn plants were infested in some fields. Small plants, such as two-leaf stage seedlings, may be completely severed by the entry of potato stem borer, and this damage resembles cutworm injury. With larger seedlings, however, larvae may burrow within the stem, feeding just above the roots, until larvae attain the fifth or sixth instar. Such mature larvae tend to remain below-ground, outside the stem, entering only to feed. Early signs of larval feeding are leaf or plant wilting; later signs are death and disintegration of the plant. Young plants perish within a few days of larval attack. Plants that have attained the eight-leaf stage are slow to wilt and die, but eventually, perish. Larvae often destroy 3–4 plants during the course of their development.

Management Sampling. Adults can be sampled with blacklight traps, and the sex pheromone of these species may eventually prove useful. Levine (1989) used temperature summation, about 1700 degree-days above a threshold of 5.3°C, to estimate peak moth flight by hop vine borer. Larvae can be

sampled by dissecting seedlings, but the below-ground portion should also be included in the sample. Wilting plants are a good indication of infestation by larvae. Insecticides. Persistent insecticides applied to crop plants and soil can provide some suppression of larvae, including those that disperse into crop fields from nearby weedy vegetation. However, better crop protection can be attained by applying insecticide directly to the source of many larvae, weedy fence rows (Deedat et  al., 1982). Insecticidal control alone is often inadequate if the borers are abundant in the proximity of a susceptible crop. Cultural Practices. Cropping practices can help alleviate injury by Hydraecia spp. Of foremost importance is weed management. Larvae often invade crop fields from weedy fence rows, resulting in considerable damage along field margins. Thus, insecticidal treatment of the crop periphery, or destruction of grasses and weeds by burning or application of herbicides, can reduce injury. The critical period for weed management is early in the season, typically April or May before susceptible annual crops are available. The presence of wild hops is often related to the occurrence of hop vine borer, whereas potato stem borer is positively affected by the presence of marshy areas where several alternate host plants may occur. Potato stem borer is most likely to be widespread in fields that are heavily infested with grasses (Deedat et al., 1982).

Redbacked Cutworm Euxoa ochrogaster (Guenée) (Lepidoptera: Noctuidae)

Natural History Distribution. Redbacked cutworm is widely distributed in northern climates, occurring in Asia as well as Canada and northern parts of the United States. It is found south to New Mexico in the Rocky Mountain region, but elsewhere is restricted to more northern latitudes, extending only as far south as Missouri. Although redbacked cutworm has damaged crops throughout Canada, it is most abundant, and damaging, in the northern Great Plains and westward, including British Columbia, Washington, and Oregon. Host Plants. Redbacked cutworm feeds on numerous crops. Among vegetables injured are asparagus, bean, beet, broccoli, cabbage, cantaloupe, cauliflower, cucumber, lettuce, onion, pea, radish, squash, tomato, and turnip. Many other vegetables probably are unreported. Other crops injured include alfalfa, alsike, barley, canola, flax, mustard, oat, sugarbeet, sunflower, sweetclover, and wheat. As might be expected from an insect with such a varied diet, there are occasional reports of injury to flowers and fruit trees. Natural Enemies. Natural enemies, particularly diseases, seem to play a significant role in redbacked cutworm biology. This cutworm displays periods of abundance l­ asting

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2–4 years, followed by periods of scarcity persisting for at least 2 years. Duration of abundance typically is preceded by warm, dry weather in late summer and followed by similar warm weather in spring. Warm and dry weather apparently allows the moths to feed freely in the summer, and minimizes the impact of fungal disease in the spring. Diseases and parasitoids apparently contribute to the c­ utworm-free period that typically follows population increase. King and Atkinson (1928) suggested that diseases were the most effective factor in reducing outbreaks, but diseases of redbacked cutworm apparently have escaped serious study. Among the parasitoids known to affect redbacked cutworm are Gonia aldrichi Tothill, G. fuscicollis Tothill, Bonnetia comta (Fallén), Periscepsia helymus (Walker) (all Diptera: Tachinidae); Villa alternata (Say), V. fulviana (Say), V. lateralis (Say), Poecilanthrax alcyon (Say), P. willistonii (Coquillet) (all Diptera: Bombyliidae); Eutanyacra suturalis (Say), Diphyus euxoa Heinrich, D. apiculatus (Walkley), Exetastes obscurus Cresson, Spilichneumon superbus (Provancher), Campoletis atkinsoni (Viereck), C. australis (Viereck), Netelia sp., Gravenhorstia propinqua (Cresson) (all Hymenoptera: Ichneumonidae); Apanteles lalticola (Ashmead), A. aeviceps (Ashmead), A. griffini Viereck, Microplitus kewleyi Muesebeck, Meteorus vulgaris Cresson, M. laeviventris (Wesmael) (Hymenoptera: Braconidae); Copidosoma bakeri (Howard) (Hymenoptera: Encyrtidae); and Agamermis sp. (Nematoda: Mermithidae). Not all species are abundant and important in population regulation, and their relative importance varies among locations and years. Nevertheless, Canadian studies indicate that the larval parasitoids Gonia aldrichi, Meteorus vulgaris, and Campoletis atkinsoni, and the egg parasitoid Copidosoma bakeri, are among the more important species (Schaaf, 1972). Predators also affect redbacked cutworm populations. In Alberta, Frank (1971) reported 21 species of ground beetles (Coleoptera: Carabidae), and several species of rove beetles (Coleoptera: Staphylinidae) feeding on eggs, larvae, or pupae. Spiders also are thought to feed on this cutworm. Life Cycle and Description. There is one generation per year. Adults are active in late July and throughout August. They deposit eggs just beneath the soil surface, and the eggs overwinter. They hatch in April, larvae develop over a 6–8 week period, and pupation normally occurs in June. Egg. The eggs are deposited in August and September in loose soil. Heavy, crusted, and wet soil tends to be avoided. They are whitish initially, become bluish as the embryo completes its development. Embryonic development is completed in 8, 10, 14, and 20 days at 30°C, 25°C, 20°C, and 15°C, respectively. Eggs do not hatch upon completion of embryonic development, the embryos remaining in diapause until spring. When held at 5°C for 3–4 months the eggs can hatch. They are quite resistant to desiccation (Jacobson, 1970).

FIG. 10.94  Redbacked cutworm moth. (Photo by J. Capinera.)

Larva. Larvae tend to live below-ground during the daylight hours, but come to the surface, and even climb plants, at night. The grayish larvae are distinguished by the presence of brick-red stripes along the back, separated by a narrow pale stripe centrally. Laterally, a black stripe borders the red stripes. The head and prothoracic plate are yellowish brown, though the head bears dark brown submedial arcs. Normally there are six instars, but when reared at 30°C only five instars develop. Mean duration (range) of the instars is estimated at 5.6 (5–7), 4.0 (3–7), 3.7 (3–8), 5.8 (3–8), 6.1 (4–11), and 14.0 (6–22) days, respectively, at 20°C. The total larval development time (including the prepupal portion of the last instar) is reported to be about 72, 39, 32, and 22 days at 15°C, 20°C, 25°C, and 30°C, respectively. Larvae attain a length of about 38 mm at maturity. Pupa. Larvae pupate in the soil at depths of 2.5–5.0 cm. The pupa is reddish brown and measures about 2 cm long. Pupal development times are about 36.8 (33–42), 21.5 (18– 23), 14.3 (13–16), and 12.6 (12–14) days at 15°C, 20°C, 25°C, and 30°C, respectively. Adult. The moths measure about 35–40 mm in wingspan. The front wings are variable and tend toward four basic types. One common wing pattern is uniformly dark reddish brown with bean-shaped and round spots on the front wings bearing a light border. Another common wing pattern is much lighter in color, often bearing a grayish bar along the leading edge of the wing. This latter color form also tends to have a black bar connecting the two spots; the spots similarly have a light margin. The hind wing in both forms is grayish basally, with brown distally and along the major veins. The major color forms of the adult were pictured by Hardwick (1965a). Females produce a sex pheromone (Struble 1981a; Palaniswamy et al., 1983) beginning about 7 days after adult emergence. Males respond to the pheromone soon after sunset (Struble and Jacobson, 1970). The preoviposition period is 6–13  days. Adults normally survive for about 20 days. A brief summary of redbacked cutworm, including management options, was given by King (1926). The ­importance of redbacked cutworm in Canada was described by

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Beirne (1971). Developmental biology was given by Jacobson (1970). A pictorial key that included redbacked cutworm adults was published by Capinera and Schaefer (1983). It was also included in a comprehensive treatment of North American noctuids (LaFontaine, 1987).

Damage Larvae generally feed at the soil surface or slightly below the surface, though on fruit crops they may climb the plant to feed on leaf and blossom buds. On herbaceous plants, young larvae eat holes and notch the leaf tissues, whereas older larvae sever the plant stem at the soil surface. The spotty distribution of damage is easily confused with poor seed germination, but the presence of severed, shriveled plants indicates cutworms. Larvae are most damaging during hot, dry weather because the plants tend to be stressed. As their distribution in the field tends to be aggregated, larvae usually damage most plants in an area, leaving few individual plants available to compensate for damage. Thus, damage tends to be almost directly proportional to abundance (Ayre, 1990). This species is considered by some to be the most destructive cutworm in Canada.

Management Sampling. Both blacklight and pheromone traps give good estimates of moth activity, though blacklight traps capture moths earlier, and pheromone traps capture moths longer (Steck et al., 1980a; Ayre et al., 1982b; Gerber and Walkof, 1992). Floral lures (acetic acid plus 3-methyl-­1-butanol, or a combination of floral odor chemicals) can also be used to trap these moths (Landolt et al., 2007). The sex pheromone, when applied in large quantities to an area, can also be used to disrupt mating. The distribution of larvae in peppermint fields is generally clumped (Danielson and Berry, 1978), and this is likely the condition in other crops. In peppermint, 20–40 soil samples were necessary to determine whether or not a crop required treatment. Insecticides. Persistent insecticides are used for redbacked cutworm suppression. Most products are effective when applied to foliage, and some when applied to soil (McDonald, 1981b). Some populations of redbacked cutworm have been known to develop measurable resistance to insecticides (Harris, 1976). As redbacked cutworm comes to the soil surface at night to feed, baits containing insecticide are very effective. Cultural Practices. Tamaki et  al. (1975) documented the abundance of cutworm larvae in weedy fields and weedy sections of fields in Washington. Perennial weeds such as field bindweed, Convolvulus arvensis, and Canada thistle, Cirsium arvense, either favor larval survival or attract ovipositing moths, leading to increased larval abundance. Tillage of fallow fields during the oviposition period deters egg laying because moths seem to select weedy patches.

Also, clean cultivation early in the season, before planting, can cause young larvae to starve. Mechanical barriers are sometimes recommended for protection of plants in the home garden. Metal or waxedpaper containers with the top and bottom removed may be placed over susceptible seedlings to deter feeding by larvae. It is helpful to extend the barrier well below the soil surface to minimize access by burrowing larvae. Biological Control. Steinernematid and heterorhabditid nematodes can infect larvae in the soil, but redbacked cutworm larvae appear to be less susceptible than some other species (Morris and Converse, 1991). Bacillus thuringiensis is not usually recommended for suppression of cutworms.

Southern Armyworm

Spodoptera eridania (Cramer) (Lepidoptera: Noctuidae)

Natural History Distribution. This insect is native to the Americas, occurring widely in North, Central and South America, and the Caribbean. In the United States, southern armyworm is found principally in the southeastern states. Although its range extends west to Kansas, New Mexico, and California, this species is of little consequence in western states. Host Plants. Southern armyworm has a very broad host range. Some of the vegetables injured are beet, cabbage, carrot, collards, cowpea, eggplant, okra, pepper, potato, squash, sweet potato, tomato, and watermelon. Other crops injured include avocado, citrus, peanut, soybean, sunflower, velvet bean, tobacco, and many flowers, especially annuals. Numerous weeds are consumed, but pigweed, Amaranthus spp.; and pokeweed, Phytolacca americana; are especially favored, and grasses are rarely eaten. There are several reports of armyworm infestations beginning with these two weeds, and adjacent crops experiencing damage only after the more favored weeds were consumed. Although reported to feed on over 200 plants from several plant families (Montezano et  al., 2014), flora is not equally suitable for larval growth and survival. For example, Santos et  al. (2005) studied growth and survival on cotton, soybean, and morning glory. Larval development on cotton and morning glory were similar, but development required longer on soybean. Also, pupal weights were higher on cotton and morning glory than on soybean. This pattern indicates that soybean is less suitable for southern armyworm. Natural Enemies. Several wasp parasitoids commonly associated with caterpillars of other species, including Cotesia marginiventris (Cresson), Chelonus insularis Cresson, Meteorus autographae Muesebeck, and M. laphygmae Viereck (all Hymenoptera: Braconidae) also

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attack southern armyworm. Meteorus autographae was the dominant parasitoid in a Florida study (Tingle et al., 1978). Also reared from southern armyworm are Campoletis flavicincta (Ashmead) and Ophion flavidus Brulle (both Hymenoptera: Ichneumonidae); Euplectrus platyhypenae Howard (Hymenoptera: Eulophidae); Choeteprosopa hedemanni Braeur and Bergenstamm, Euphorocera claripennis (Macquart), Gonia crassicornis (Fabricius), Winthemia quadripustulata (Fabricius), and W. rufopicta (Bigot) (all Diptera: Tachinidae). Predators certainly must be an important factor in southern armyworm biology, but this aspect seems undocumented. Larvae are susceptible to infection by the fungus Beauveria bassiana (Gardner and Noblet, 1978). An ectoparasitic nematode, Noctuidonema guyanense, parasitizes the adult, and is thought to be debilitating (Simmons and Rogers, 1996). Life Cycle and Description. The number of generations is estimated at four annually in Florida. Insects may be present throughout the insect’s range from March until October, but they are most commonly observed in late summer and autumn. In northern Florida, moths can be found throughout the year, withstanding several days of freezing weather (Mitchell and Tumlinson, 1994). About 30–40 days are required for a complete generation. When reared at 25°C, the mean duration of the egg, larval, prepupal, and pupal stages were reported to be 4.0, 16.2, 1.6, and 9.2 days, respectively (Montezano et al., 2014).

FIG. 10.96  Larva of southern armyworm. (Photo by L. Buss.)

Larva. The larvae display six or seven instars, although most have six, as they grow to attain a length of about 35 mm. The head capsule widths are about 0.25–0.30, 0.40–0.50, 0.60–0.80, 0.95–1.15, 1.35–1.85, and 2.35–2.85 mm, respectively (Redfern, 1967). Larvae are green or blackish green with a uniform light brown or reddish-brown head throughout their development. Larger larvae bear a narrow white line dorsally, and additional stripes laterally. Each side normally bears a broad yellowish or whitish stripe that is interrupted by a dark spot on the first abdominal segment, though in some cases this spot is weak. The lateral spot on the first abdominal segment is an important diagnostic character in this species, and much more reliable than other color-based characteristics. A series of dark triangles is usually present dorsolaterally along the length of the body. Larvae are usually found on the lower surface of leaves and are most active at night. Duration of the larval stage is normally 14–20 days. Pupa. The larvae pupate in the soil, usually burrowing to a depth of 5–10 cm. The pupa is mahogany brown and measures about 16–18 mm long and 56 mm wide. Duration of the pupal period is 11–13 days.

FIG. 10.95  Eggs of southern armyworm. (Photo by L. Buss.)

Egg. The shape of the egg is a flattened sphere. Eggs measure about 0.45 mm in diameter and 0.35 mm in height. They bear about 50 slender ribs which radiate outward from the center. The eggs are greenish initially, turning tan as they age. They are laid in clusters and covered with scales from the body of the moth. Duration of the egg stage is 4–6 days. Under laboratory conditions a mean of nearly 1400 eggs can be produced (range of 634–1900) (Montezano et al., 2013).

FIG. 10.97  Moth of southern armyworm. (Photo by L. Buss.)

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Adult. The moth measures 33–38 mm in wingspan. The front wings are gray and brown, with irregular dark brown and black markings. The wing pattern is highly variable. Some individuals bear a pronounced bean-shaped spot (reniform spot) near the center of the wing, whereas others lack the spot or instead bear a broad black band extending from the center of the wing to the margin. Usually, it is the adult males that have a pronounced band or streak that extends into a streak attaining the tip of the wing, whereas in adult females the reniform spot is weak or absent. The hind wings are opalescent white. Male and female longevity, when held at 25°C, is reported to be 9.3 and 10.8 days, respectively (Montezano et al., 2014). The biology of southern armyworm is not fully documented, but Chittenden and Russell (1910) gave key features. Additional description was given by Crumb (1929). The larval keys developed by Levy and Habeck (1976), Passoa (1991), and Heppner (1998) are useful to distinguish southern armyworm from related species. It was also included in a key by Okumura (1962), Oliver and Chapin (1981), and Stehr (1987). Heppner (1998) provided a useful key to the adults of North American Spodoptera. Rearing techniques were provided by Redfern and Raulston (1970). A sex pheromone has been identified and evaluated in the field (Mitchell and Tumlinson, 1994).

Damage This is one of the most common pests of southeastern vegetable gardens. Larvae are defoliators and feed gregariously while young, often skeletonizing leaves. As they mature they become solitary, and also bore readily into fruit, often damaging tomato. When stressed by lack of food they eat the apical portions of branches, bore into stem tissue, and attack tubers near the surface of the soil. High densities and lack of food sometimes prompt larvae to move in great number, the basis of the “armyworm” designation, feeding on all vegetation in their path. Southern armyworm is more damaging to cassava than black cutworm, Agrotis ipsilon (Hufnagel), given equal numbers of insects (Pena and Waddill, 1981).

Management Insecticides. Southern armyworm is best controlled with foliar insecticides when larvae are small. Insecticides vary considerably in their toxicity to larvae (Aziz, 1973). This insect is difficult to control with botanical insecticides (Valles and Capinera, 1993). Berger (1920) reported some success at southern armyworm suppression by application of bran bait containing insecticide. However, this is useful principally for large, mobile larvae that have left the plant and are on the soil surface. Liburd et al. (2000) evaluated several biological insecticides on southern armyworm abundance and reported that although several Bacillus

thuringiensis products, azadirachtin, and a Steinernema carpocapsae product could reduce armyworm abundance significantly below untreated tomatoes, conventional chemical insecticides generally were more effective. Host-Plant Resistance. There is limited information that suggested differences in susceptibility among sweet potato cultivars to armyworm damage (Habeck, 1976). Conventional sources of insect resistance in corn have little effect on southern armyworm (Manuwoto and Scriber, 1982).

Soybean Looper

Chrysodeixis includens (Walker) (Lepidoptera: Noctuidae)

Natural History Distribution. Formerly known as Pseudoplusia includens (Walker), soybean looper is native to the western hemisphere, where it is found from the United States to Argentina. It is a tropical or subtropical insect and overwinters successfully only in warm climates. In the United States, it survives the winter in southern Florida and southern Texas. Not surprisingly, damaging populations are largely restricted to the southeastern and south-central states. The moths are highly dispersive, however, and this insect has been captured throughout the United States, except for the Pacific Northwest. It is rare in Canada but has been recovered from southern Ontario and Quebec, and from Nova Scotia. Host Plants. This insect is known principally as a defoliator of soybean, but it reportedly has a fairly wide host range. Vegetables reported to be consumed are asparagus, bean, broccoli, celery, collards, corn, cowpea, garlic, lettuce, mustard, okra, pea, pepper, potato, sweet potato, tomato, watercress, and watermelon. Other host plants include field crops such as alfalfa, cotton, peanut, sunflower, and tobacco, and flower crops such as aster, begonia, calendula, carnation, chrysanthemum, geranium, and poinsettia. Weeds are also suitable hosts, including such common species as pigweed, Amaranthus sp.; cocklebur, Xanthium pennsylvanicum; horseweed, Egeron canadensis; wild sunflower, Helianthus spp.; pepperweed, Lepidium virginicum; and dock, Rumex spp. Herzog (1980) provided a long list of host plants. Soybean looper has long been confused with cabbage looper, Trichoplusia ni; the larvae are difficult to separate accurately. Some of the aforementioned records may reflect such misidentifications, though this insect certainly does occur on vegetables, particularly tomato. It is invariably the dominant looper on soybean in the southeast, but cabbage looper is the dominant looper species on vegetables. Martin et al. (1976a) presented interesting data from Florida, collected both from field cages and crop fields, demonstrating the preference of soybean looper for field crops.

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Natural Enemies. Soybean looper is host to numerous parasitoids, pathogens, and predators. In Georgia, it is common for 40% of larvae to succumb to a parasitoid or disease (Beach and Todd, 1985). In Louisiana, such natural mortality seems to be even higher, but the effectiveness of these biotic control agents varies considerably with time, location, and cropping system (Burleigh, 1972). Nearly all research has emphasized soybean, so results of some studies may not be applicable to vegetables. It is of interest to note that Martin et al. (1981b) reported higher levels of parasitism on soybean looper in vegetables than in soybean during studies conducted in northern Florida. The most important parasitoids tend to be Copidosoma truncatellum (Dalman) (Hymenoptera: Encyrtidae), Meteorus autographae Muesebeck, Apanteles scitulus Riley, and Cotesia marginiventris (Cresson) (all Hymenoptera: Braconidae). Several other wasp and fly (Diptera: Tachinidae) parasitoids exert only low levels of mortality (Burleigh, 1971, 1972; Beach and Todd, 1985; Daigle et al., 1990). Fungi are important mortality factors for soybean looper. Entomophthora gammae, Massospora sp., and Metarhizium (Nomuraea) rileyi are especially common. A NPV is also known. Often the pathogens are not effective until late in the season, or until looper densities are high (Burleigh, 1972; Harper and Carner, 1973). Many general predators such as the lady beetle Coleomegilla maculata (Coleoptera: Coccinellidae); bigeyed bug, Geocoris spp. (Hemiptera: Lygaeidae); the damsel bug Nabis roseipennis Reuter (Hemiptera: Nabidae); and the ground beetle Calosoma spp. (Coleoptera: Carabidae) feed on soybean looper. All stages of development are subject to predation (Richman et al., 1980; Brown and Goyer, 1984). Life Cycle and Description. Total generation time for soybean looper is estimated at about 27–34 days during the summer months. In most of Florida, moths remain active throughout the year, but in the north the period of adult activity is restricted (Mitchell et al., 1975a, b). Moths disperse northward annually, attaining Georgia in June and July, and South and North Carolina in August and September. There are likely three to five generations annually in the southern areas of soybean looper’s range, and two per year in most northern latitudes such as North Carolina. Egg. The eggs are slightly flattened spheres, laid singly with a flattened side affixed to the foliage. They measure about 0.6 mm in diameter and 0.4 mm in height, and their close examination reveals numerous ridges radiating from the top of the egg. There are about 34 ridges per egg, but unlike many other noctuid eggs, the ridges are not sharply defined and easy to count. Initially, the egg is white but turns light brown with maturity. Duration of the egg stage averages 4.1 days at 27°C. Females deposit about 300–600 eggs. Egg deposition is affected by adult food sources,

c­otton nectar being especially favorable (Jensen et  al., 1974). Maximum egg production occurs at about 29°C (Mason and Mack, 1984). Larva. The larvae are green, and eventually, attain a length of about 30 mm. The body is marked with thin white stripes dorsally and along the sides. In most respects soybean looper closely resembles cabbage looper, Trichoplusia ni, including the presence of only three pairs of prolegs and the presence of nipple-like vestigial prolegs on abdominal segments three and four. Soybean looper differs in having minute microspines, but this character is of marginal diagnostic value because the microspines are difficult to detect. The thoracic legs of soybean looper are often dark, and sometimes can be used to differentiate a specimen from cabbage looper. A reliable character for differentiation of these species is structure of the mandible [see Eichlin and Cunningham (1978) for an illustration of this character]. The head is grayish green, and may be marked with dark bands laterally. Generally there are six larval instars. According to Mitchell (1967), mean duration (range) of the instars is 3.7 (3–5), 2.1 (2–3), 2.2 (1–5), 2.6 (1–5), 2.9 (1–5), and 6.2 (4–11) days, respectively, for instars 1–6 when reared at 27°C. Head capsule widths for the six instars are about 0.25, 0.39, 0.60, 0.96, 1.45, and 2.18 mm, respectively. However, Shour and Sparks (1981) noted that the total number of instars could vary, and that total larval duration was 15.4, 16.8, or 20.7 days depending on whether there were 5, 6, or 7 instars, respectively. Strand (1990) described the basis for instar variation in soybean looper. Pupa. Larvae spin a loose silk cocoon on the foliage and pupate within. Duration of the pupal stage is about 7 days (range 3–11 days). Adult. The moth’s front wings are marked with various shades of brown and gray, but the overall effect is dark. The silvery-white marking at the center of the forewing, which is sometimes said to resemble a “dog leg,” usually has a detached “foot.” The hind wings are light brown basally and dark brown distally. The moth is readily confused with plantain looper, Autographa precationis, but the forewing of plantain looper is rusty red, whereas that of soybean looper has a brassy reflection. The wingspan of soybean looper moths is 30–39 mm. Moths begin oviposition 3–4 days after emergence, and adult longevity is typically 5–10 days. Moth activity is highest between 10 p.m. and 2 a.m. (Mitchell 1973). A review of soybean looper biology was given by Herzog (1980). Rearing procedures were given by Hartley (1990). A multicomponent sex pheromone was described by Linn Jr. et al. (1987). Keys to adults and larvae can be found in LaFontaine and Poole (1991). Larvae were also included

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in the key of Crumb (1956), Stehr (1987), and Sparks and Liu (2001), and adults in Eichlin and Cunningham (1978). Soybean looper is included in the key to vegetable-feeding loopers found in Appendix A.

Damage Larvae feed principally on leaf tissue. Foliage consumption of soybean, which probably is similar for green bean, is 0.2, 0.8, 1.8, 7.9, 15.5, and 55.7 cm2 for the six instars, respectively (Reid and Greene, 1973). Larvae also sometimes feed on the pods of legumes, and the silk and kernels of corn (Janes and Greene, 1970).

Management Sampling. Soybean looper moths can be captured with light and pheromone traps (Tumlinson et  al., 1972), or a combination of light and pheromone traps (Mitchell et al., 1975a, b). Larval sampling has been studied extensively in soybean, but methodology for vegetables has not been developed. Herzog (1980) reviewed soybean looper sampling. Insecticides. Damage is usually prevented by the application of insecticide to foliage. However, insecticide resistance has been reported to be a serious problem (Leonard et al., 1990). Population increases of soybean looper have been reported following foliar application of certain insecticides; it seems likely that destruction of beneficial insects by insecticides allows the loopers to attain elevated levels of abundance (Shepard et al., 1977). Soil-applied systemic insecticides have little effect on parasitism (McCutcheon et al., 1990). Biological Control. Natural enemies are often adequate to maintain soybean looper at nondamaging densities on vegetable crops. If this level of control is not adequate, B. thuringiensis can be applied. A NPV has been demonstrated to be effective under field conditions, but it is not available commercially (McLeod et al., 1982). Cultural Control. After overwintering in southernmost areas of United States, soybean looper must migrate north annually. Thus, early-planted crops are less susceptible to injury than late-planted crops.

Spotted Cutworm

Xestia c-nigrum (Linnaeus) Xestia dolosa Franclemont (Lepidoptera: Noctuidae)

Natural History Distribution. Spotted cutworm was long known as Xestia (Amathes) c-nigrum (Linnaeus), a Eurasian species. However, Franclemont (1980) differentiated the insects found in North America into two other species, Xestia adela and Xestia dolosa. Later, Lafontaine (1998) determined

that the two species in North America were X. c-nigrum and X. dolosa; X. adela is not a valid name. Based on the lack of different pheromone responses, Landolt et al. (2011a) reported that there was only one species in Washington State, X. c-nigrum, and X. dolosa has long been known to be restricted to eastern North America. Hudson (1982) suggested that these species might be derived from separate introductions of X. c-nigrum, but in any case, they respond differentially to pheromones and if cross mated no fertile eggs are obtained. Host Plants. “Spotted cutworm” a is a general feeder, consuming various flowers, vegetables, fruit trees, and other plants. Among vegetables attacked are asparagus, beet, cabbage, carrot, cauliflower, celery, corn, lettuce, onion, pea, potato, rhubarb, tomato, and turnip. Fruits consumed include apple, cranberry, currant, gooseberry, raspberry, and pear. Occasional damage is incurred by barley, clover, flax, oat, rye, sugarbeet, tobacco, and wheat. Various weeds are consumed, including Canada thistle, Cirsium arvense; chickweed, Stellaria sp.; goldenrod, Solidago sp.; lambsquarters, Chenopodium album; morningglory, Ipomoea sp.; redroot pigweed, Amaranthus retroflexus; smartweed, Persicaria sp.; sunflower, Helianthus sp.; ferns and grasses. Natural Enemies. Flies known to parasitize spotted cutworm include Euphorocera claripennis (Macquart), Winthemia quadripustulata (Fabricius), and W. rufopicta (Bigot) (all Diptera: Tachinidae). Among the wasp parasitoids of spotted cutworm are Apanteles yakutatonsis Ashmead (Hymenoptera: Braconidae), Eutanyacra succincta (Brulle) (Hymenoptera: Ichneumonidae), Dibrachys carus (Walker) (Hymenoptera: Pteromalidae), and Euplectrus frontalis Howard (Hymenoptera: Eucharitidae). Life Cycle and Description. The number of generations is variable. There are reportedly three generations annually in Tennessee, with moths present in April-May, July, and September-October (Crumb, 1929), though evidence for the middle generation is weak. In Washington, two flight generally are observed, but a small third flight late in the year is possible (Howell, 1979). In New York, Minnesota, and elsewhere, flights of moths occur in JuneJuly and in August-October (Chapman and Lienk, 1981; Knutson, 1944); this seems to be the most common pattern of occurrence. Overwintering occurs in the larval stage, with pupation in early spring. Most damage to crops results from feeding by larvae of the summer generation, which normally pupate in August. Egg. The eggs are normally deposited on the underside of foliage. They are pinkish and in shape resemble a slightly flattened sphere. They measure 0.60–0.65 mm in diameter and 0.50–0.61 mm in height. Eggs bear about 30 ridges, of which about 13–14 extend from the base to

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the center; the remainder terminates before they reach the top. They reportedly are deposited singly, in rows, or in clusters of up to 200 eggs. Duration of the egg stage is 6–12 days. Larva. Spotted cutworm displays seven instars. Head capsule widths are about 0.35, 0.55, 0.8, 1.3, 1.7, 2.1, and 3.0 mm for instars 1–7, respectively. Body length is about 2–4, 6, 0.9–1.5, 12–16, 14–20, 28, and 30–36 mm, respectively. The general body color is brown or gray, and it lacks distinct stripes. The most diagnostic character is a series of subdorsal triangles or wedges along the length of the body. The triangles are distinct posteriorly but decrease in size toward the head. The head is light brown with dark markings. Duration of the larval stage in the winter period is 5–6  months, whereas it is only about 1 month during the summer. The larvae of X. adela and X. dolosa are structurally indistinguishable. However, X. dolosa has a longer developmental period and attains a greater size at larval maturity. When X. dolosa is reared at 24°C, the larval, pupal, and egg-adult periods were 27.6, 15.8, and 43.3  days, respectively. In contrast, the corresponding periods were 21.0, 12.1, and 32.4 days for X. adela. Maximum larval weights when reared at 24°C were 0.99 g for X. dolosa but only 0.56 g for X. adela. Not surprisingly, head capsule widths of X. dolosa tend to be slightly larger than X. adela. Although head size is about the same for both species through instar four, the widths are about 1.8 and 1.7 mm in instar five, and 3.0 and 2.4 in instar six for X. dolosa and X. adela, respectively (Hudson, 1983). Larvae of all ages are prone to climb, but older larvae climb and feed only at night, dropping to the ground and seeking shelter during the daylight hours (Olson and Rings, 1969). Pupa. Pupation occurs in the soil. The pupa is about 18 mm long and 6 mm wide. It is brown. Duration of the pupal stage is 17–41 days.

Adult. The adult is a medium-sized moth, measuring 29–43 mm in wingspan. The moth of X. dolosa, which has a wingspan of 37–43 mm, is slightly larger than X. adela, with a wingspan of 29–38 mm. The front wings are purplish or reddish brown, with dark brown to black basally. At about the midpoint of the leading edge of the forewing is a light brown or tan-colored triangle. On average, X. adela tends to be slightly darker than X. dolosa. The hind wings are white to light gray, often darker along the outer margin. Moths of X. dolosa and X. adela fly at about the same periods each year. However, as might be expected from the longer developmental period of X. dolosa, the flight period of this species begins at a slightly later date than X. adela. Adults live 2–3 weeks in the field. These species are poorly known. Crumb (1929) gave the most complete account, but Howell (1979) provided valuable data from the west coast. Hudson (1982) supplied developmental data. Franclemont (1980) and Hudson (1982) clarified the status of the North American species comprising dingy cutworm. Rings and Johnson (1977) published a bibliography. Spotted cutworm larvae were included in keys by Whelan (1935), Crumb (1956), Okumura (1962), Rings (1977b), Capinera (1986), and Stehr (1987), and are included in a key to armyworms and cutworms in Appendix A. Moths were included in keys by Rings (1977a) and Capinera and Schaefer (1983).

Damage Damage to crops results from feeding by both the overwintering larvae in the spring and from the summer generation later in the growing season. The overwintering larvae are perhaps best known for their damage to fruit because larvae readily climb vines and trees to feed on buds during the spring months, when vegetation is scarce. Although the larvae can sever young plants at ground level, they also feed on foliage and burrow into tomato fruits. At high densities, larvae may assume a gregarious, dispersive “armyworm” habit. Spotted cutworm has been reported to be an early-season rangeland pest, destroying nearly all forbs and grasses in some areas (Launchbaugh and Owensby, 1982).

Management

FIG. 10.98  Spotted cutworm moth. (Photo by J. Capinera.)

Sampling. Light traps can be used to monitor populations of adults. A pheromone-based lure has also been identified, though it is not completely specific (Landolt, 2000a, b). Although these are considered to be climbing cutworms, and larvae spend a good deal of time searching or feeding on foliar tissue, they also hide beneath plant debris and beneath the soil surface. Thus, it is important to rake the surface of the soil to search for larvae if cutworm injury is suspected. Insecticides. Insecticides are commonly used to control cutworms. Application of persistent insecticides to the base of the plant can be effective, as can foliar treatment

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of leafy vegetables. Bran and other organic baits treated with insecticide are effective methods of cutworm suppression. Bacillus thuringiensis is not usually recommended for these pests. Cultural Practices. Barriers have been used to prevent access by cutworms to buds and foliar tissues (Wright and Cone, 1983b), but in commercial crop production, this has more applicability to perennial crops such as fruit than to annual crops such as vegetables. In the home garden, however, transplanted plants can be protected by a barrier such as a can or waxed paper container with the top and bottom removed. Aluminum foil wrapped around the base of the seedling also deters cutting by larvae. Ditches with steep sides and metal barriers can be effective impediments to prevent dispersal of cutworm larvae into crops. Cutworm problems often develop in weedy fields or portions of fields infested with weeds. It is advisable to till, or otherwise destroy weeds, 10–14 days in advance of planting, as this should cause small larvae to starve. If seedlings are to be transplanted into a field or garden, larger plants are preferred because they are less likely to be irreparably damaged by cutworms.

Stalk Borer

Papaipema nebris (Guenée) (Lepidoptera: Noctuidae)

Natural History Distribution. This native insect occurs throughout the eastern United States and southern Canada east of the Rocky Mountains. It is rarely abundant in the southern states and along the western margin of its distribution, occurring as a pest principally in the midwestern states. There are nearly 50 species of Papaipema found in northeast and northcentral United States and adjacent Canada. Although, P. nebris is the dominant pest, on occasion related species such as P. cataphracta (Grote), the burdock borer, have been reported to damage crop plants. Accurate identification of these insects is difficult in the larval stage owing to their similarity in appearances and habits. Based on light trap collections in Iowa, Peterson et al. (1990) concluded that P. nebris was by far the most abundant species in this group. Host Plants. Stalk borer has a very wide host range, with almost 200 plant species recorded as hosts. In the spring the larvae burrow into grass stems, but as they grow the larvae move to nearby plants with thicker stems. Among the vegetable crops injured are asparagus, bean, cantaloupe, cauliflower, celery, corn, eggplant, parsnip, pea, pepper, potato, rhubarb, spinach, and tomato. Other crops sometimes injured include alfalfa, barley, cotton, oat, red clover, rye, sugarbeet, sweet clover, timothy, and wheat. Fruit damaged by stalk borer include apple, blackberry, currant, g­ ooseberry,

strawberry, peach, and plum. Shade trees can also be injured, including catalpa, elm, maple, poplar, and willow (Solomon, 1988). Among the numerous flower crops that have been reported to be damaged are anemone, canna, carnation, cosmos, daisy, gladiolus, hollyhock, iris, larkspur, lily, peony, phlox, purple coneflower, rose, and rose mallow. Some of the common weeds supporting stalk borer larvae are cattail, Typha spp.; dock, Rumex spp.; Kentucky bluegrass, Poa pratensis; switchgrass, Panicum virgatum L.; dogbane, Apocynum androsaemifolium; groundcherry, Physalis spp.; goldenrod, Solidago spp.; lambsquarters, Chenopodium album; quackgrass, Agropyron repens; ragweed, Ambrosia spp.; smartweed, Polygonum spp.; sunflower, Helianthus spp.; thistle, Cirsium spp.; wildrye, Elymus canadensis, and many others. Ragweed is often suggested as the favored host. Some common weeds such as milkweed, Asclepias syriaca; and velvetleaf, Abutilon theophrasti; are not suitable. Small-stemmed grasses often induce larval wandering (Alvarado et al., 1989). Natural Enemies. Stalk borer is attacked by many of the general predators that are found attacking other caterpillars. Because stalk borers often move among host plants, they are likely more susceptible to predation than some borers. Among the known predators are ground beetles (Coleoptera: Carabidae), lady beetles (Coleoptera: Coccinellidae), minute pirate bugs (Hemiptera: Anthocoridae), stink bugs (Hemiptera: Pentatomidae), and damsel bugs (Hemiptera: Nabidae). Parasitoids may be more important natural enemies than predators, but the significance of individual parasitoids varies among localities and habitats. Of more than 20 parasitoids known to attack stalk borer, Lydella radicis (Townsend) (Diptera Tachinidae) was reported to be the most important in Iowa (Decker, 1931), parasitizing up to 70% of larvae. The most important wasp parasitoid in Iowa, and probably the second most important parasitoid was Apanteles papaipemae Muesebeck (Hymenoptera: Braconidae); this parasitoid attained levels of parasitism up to 38% but averaged about 10%. In contrast, the important parasitoids in Ohio were Lixophaga thoracica (Curran) (Diptera: Tachinidae) in corn, Sympiesis viridula (Thompson) (Hymenoptera: Eulophidae) in potato and ragweed, and Lissonata ­brunnea (Cresson) (Hymenoptera: Ichneumonidae) and Gymnochaeta ruficornis Williston (Diptera: Tachinidae) in ragweed (Felland, 1990). Lasack et al. (1987) reported only low levels of parasitism in Iowa. Weather. Weather also influences the abundance of stalk borer. Both excessive rainfall and hot, dry weather during the spring when larvae are hatching and moving from host to host are reported to reduce larval survival markedly (Decker, 1931; Lasack et al., 1987). Life Cycle and Description. There is a single generation annually. The egg is the overwintering stage. Hatching occurs in April-June, followed by a larval development

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period of 60–90 days. Pupation occurs in late summer and autumn, with moths present and oviposition occurring during August-October. Population monitoring in Iowa demonstrated that nearly all moths were found during September (Bailey et al., 1985). Egg. The eggs are deposited singly or in clusters of up to 100 on the stems and leaves of dead grasses and weeds. The preferred site of oviposition is within rolled leaves, leaf sheaths, or cracks and crevices, particularly of narrow-leaved perennial grasses (Levine, 1985; Highland and Roberts, 1989). In shape, the egg is a slightly flattened sphere. It measures about 0.6 mm in diameter and 0.45 mm in height. It is white when first deposited, but turns grayish or amber with age. Narrow ridges, about 50 in number, radiate outward from the center of the egg. They normally hatch after a period of 7–9 months, usually in April or May. Under controlled conditions, postdiapause eggs require a mean of 12.4, 14.6, 18.8, 27.5, and 41.7 days for development when held at 24°C, 21°C, 18°C, 1°C, and 13°C, respectively (Levine, 1983).

FIG. 10.99  Stalk borer larva. (Photo by L. Buss.)

Larva. Larvae initially mine leaves, or if feeding on grass, then the stem may be attacked. However, larvae relocate to plants with large-diameter stems such as ragweed as these hosts become available. The shift among host plants occurs principally during instars 4–6. Depending on food availability, larvae may be forced to move repeatedly before completing their development. The larva is cylindrical in form, but tapering toward both the anterior and posterior ends. The head and thoracic shield are dark brown or black during the first two instars, and yellowish thereafter, though marked with a dark narrow band laterally. The larva bears pairs of prolegs on the third-sixth abdominal segments in addition to anal prolegs. The body color is brown with a broad white stripe dorsally and on each side. The lateral stripes are interrupted by a large brown spot in the region

behind the thoracic legs. The stripes fade at maturity, the larva assuming a whitish or purplish color. At maturity, the larva attains a length of about 27 mm. Larval development entails 6–17 instars, with larvae usually displaying 7–9 instars. Head capsule widths for larvae with seven instars are 0.25, 0.38, 0.57, 0.87, 1.3, 2.0, and 2.9 mm, respectively, for instars 1–7. In contrast, head capsule widths for larvae with nine instars are 0.25, 0.34, 0.46, 0.63, 0.87, 1.2, 1.6, 2.1, and 2.9 mm, respectively, for instars 1–9. The development time for larvae with seven instars when reared at 27°C is 4, 3.6, 3.7, 4.5, 10, 15, and 28 days for instars 1–7, respectively, producing a mean larval development time of 68  days. Development time for larvae with nine instars when reared at 27°C is 4, 3.4, 3.9, 4.2, 5.3, 6.2, 9.0, 14.1, and 26 days for instars 1–9, respectively, producing a mean larval development time of 76 days. Lasack and Pedigo (1986) indicated a developmental threshold of about 5.1°C. Cannibalism is common among stalk borer larvae, the larger individuals usually killing and consuming the smaller. Pupa. When larval development has been completed, larvae usually move to the soil and prepare a small cell just beneath the surface, though in some hosts such as corn pupation often occurs within the stem of the host. After a prepupal period of 1–6 days, pupation occurs. The pupa is typical in form—elongate, broadly rounded anteriorly, tapering posteriorly, and terminating in a pair of small curved spines. It is reddish brown or brown and measures 16–22 mm long and 5–7 mm wide. Duration of the pupal stage is normally 22–29 days. Adult. The adult is medium in size, with a wingspan of 25–40 mm. The general color is grayish brown, but close examination shows that the wings are dark brown and covered with scales bearing white tips. The basal two-thirds of the front wings tend to be darker, and separated from the distal portion by a thin white transverse line. The hind wings are paler, similar to the distal portion of the forewings. Some moths also bear spots on the forewings—three small whitish spots about one-third the distance to the wing tip and a bean-shaped yellowish or whitish spot just beyond the midpoint of the wing. Moths are nocturnal, and begin copulation and oviposition within three nights of emergence. The period of oviposition averages about 10  days (range 4–23  days), and is followed by death within a few days (mean 2.4 days, maximum 9 days). Females may produce 200–500 eggs daily, with average fecundity reported to be about 900 and maximum egg production just over 2000. Adults seem to be weak fliers, making only short flights. An excellent treatment of stalk borer biology was given by Decker (1931), and updates by Rice and Davis (2010). Temperature relations were given by Levine (1983). Egg diapause was described by Levine (1986b). A key for identification of larvae was provided by Crumb (1956),

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Capinera (1986), and Stehr (1987). A guide to common stalk boring caterpillars is also included in Appendix A.

Damage Historically, damage has been sporadic and limited to border rows of crops. In recent years, however, as reduced tillage practices have become more widespread in corn production areas, population densities have increased and damage has become more frequent. Young larvae enter a variety of hosts in the spring, but often choose grass plants, because these tend to predominate early in the season. Larvae usually enter the plant by burrowing into the stem, but they may also mine leaves. Leaf mining in young corn plants causes no significant loss, but when larvae burrow into the whorl, causing its death (called “dead heart”), significant damage occurs (Bailey and Pedigo, 1986). The youngest corn seedlings are most susceptible to injury, and little damage is observed once corn attains the six-leaf stage (Levine et al., 1984; Davis and Pedigo, 1990b, 1991). If a food source of a single stem is exhausted, larvae move to other stems or plants. The entrance to the plant may be anywhere along the stem and is usually made obvious by the large entrance hole. The stem is often completely hollowed out, causing the distal portions of the plant to perish. Stalk borer larvae sometimes feed on tissue of woody plants such as trees, but only the soft terminal tissue is damaged.

Management Sampling. Moths can be attracted with blacklight traps, though most males are captured (Bailey et  al., 1985). The distribution of eggs and young larvae is highly aggregated, but larval distribution is altered and it becomes more uniform as larvae disperse. Many samples are required to estimate density, especially in wild grasses. However, it is not always necessary to dissect grass stems to locate larvae, because infested stems wilt and discolor (Davis and Pedigo, 1989). Insecticides. Persistent insecticides, including systemic materials, can be applied to rows of crop plants at the margin of crop fields to reduce damage by invading larvae. However, if crop fields support grass, particularly in the autumn when eggs are deposited, the entire field may require treatment. Davis and Pedigo (1990a) demonstrated that treatment of weedy areas adjacent to crop fields, especially if timed to coincide with larval hatch, could provide good crop protection. Cultural Practices. The abundance of stalk borer is directly related to the availability of preferred weedy host plants in or near crop fields. Thus, field edges and small fields are more likely to experience damage. Stinner et  al. (1984) noted the preference by ovipositing moths for grasses within crop fields, especially fields that were grown under reduced tillage practices. However, the p­ resence of b­ road-leaf weeds has also been shown to be correlated with increased

a­ bundance of stalk borer (Pavuk and Stinner, 1991). Reduced tillage practices often result in higher weed densities within crop fields, and greater damage by stalk borer (Willson and Eisley, 1992; Levine, 1993). Thus, destruction of weeds and grasses at field margins is recommended to reduce the invasion potential by larvae dispersing from weeds, but weeds within fields must also be suppressed.

Striped Grass Looper

Mocis latipes (Guenée) (Lepidoptera: Noctuidae)

Natural History Distribution. Striped grass looper is native to the western hemisphere, where it occurs commonly from the southern United States south to Brazil, including the Caribbean. On occasion, it is reported from as far north as Labrador and as far south as Argentina, but its occurrence is always limited to east of the Rocky Mountains and Andes Mountains. In the United States it is damaging only in the Gulf Coast region, where warm weather favors the survival of this tropical insect. Other species of the genus occur in the same area but are not thought to be important pests. Host Plants. As implied by its common name, this species feeds only on grasses. Several crops support striped grass looper, including bahiagrass, Bermudagrass, corn, guineagrass, millet, oat, pangolagrass, paragrass, rice, ryegrass, St. Augustine grass, Sudan grass, sugarcane, sorghum, and wheat. Among vegetable crops, only sweet corn is injured. Wild grasses reported as hosts include barnyardgrass, Echinochloa crusgalli; bluestem, Andropogon spp.; crabgrass, Digitaria spp.; goosegrass, Elusine indica; Johnsongrass, Sorghum halepense; panicum, Panicum spp.; and others. Natural Enemies. Although there is extensive information on natural enemies affecting striped grass looper in Central and South America (e.g., Cave, 1992), data from North America are limited. Hall (1985) reported larval and pupal parasitism rates of 7.0%–44.6% with a mean value of 29% in southern Florida sugarcane fields. Sarcophagid and tachinid flies seem to be most important, and include Sarcodexia sternodontis Townsend and Sarcophaga sp. (both Diptera: Sarcophagidae); Archytas marmoratus (Townsend), Attacta brasiliensis Schiner, Belvosia bicincta Robineau-Desvoidy, Chetogena sp., Eucelatoria armigera (Coquillett), Euphorocera claripennis (Macquart), and Lespesia aletiae (Riley) (all Diptera: Tachinidae). Among the wasp parasitoids are Brachymeria ovata (Say), B. robusta (Cresson), Spilochalcis sanguineiventris (Cresson), Spilochalcis n. sp. (Hymenoptera: Chalcididae); Copidosoma truncatellum (Dalman) (Hymenoptera: Encyrtidae); Apanteles scitulus Riley, Meteorus autographe Muesebeck, Microplitis maturus Weed (Hymenoptera: Braconidae); Coccygomimus aequalis (Provancher), Enicospilus

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­purgatus (Say), Gambrus ultimus (Cresson) (Hymenoptera: Ichneumonidae); and Trichogramma sp. (Hymenoptera: Trichogrammatidae). The ectoparasitic nematodes Noctuidonema guyanense and N. diddbolia (Nematoda: Aphelenchoidea) are found on the bodies of moths (Rogers and Marti Jr., 1993; Marti and Rogers, 1995; Simmons and Rogers, 1996), and are thought to be debilitating but cause few detectable effects on the insects. Predation may be an important factor, if overlooked element in striped grass looper population dynamics. In addition to predation by such usual predators as lady beetles (Coleoptera: Coccinellidae), there are frequent reports of predation by Anolis spp. lizards and songbirds such as eastern meadowlark, Sturnella magna, and redwing blackbird, Agelaius phoeniceus. Pathogens associated with striped grass looper include the fungi Beauveria bassiana and Metarhizium (Nomuraea) rileyi and a virus, but the importance of these microorganisms has not been determined. Life History and Description. Striped grass looper is found throughout the year in southern Florida and Puerto Rico; elsewhere in the United States, it is abundant principally during late summer and autumn. There is no period of diapause in this species. Duration of the complete life cycle has been given by most authors as 40–60  days, though it varies with temperature. The developmental threshold for this insect is reported to be 13.7°C, and it tolerates temperature of 20–30°C readily. Thus, several generations occur annually in the south.

Larva. The larvae are slender and elongate throughout development. Initially, the larvae measure only about 3 mm long, eventually attaining a length of 55–70 mm. The body is yellowish or tan but marked with numerous, narrow, brown and black stripes. The stripes extend from the body onto the head. In many specimens, a dark brown or black band encircles the body between the first and second, and second and third abdominal segments. Viewed from the side, these bands appear to be dorsal spots. The larva bears three pairs of prolegs and moves with a looping motion. Duration of the larval stage is 20–40 days. Alvarez and Sanchez (1981) reported mean (range) development times of 2.0 (2–2), 2.5 (2–3), 2.7 (2–5), 3.5 (3–4), 1.8 (1–3), and 4.9 (4–5) days, respectively, for instars 1–6 when cultured at 30°C. They also reported a mean development time for larvae of 17.4 days but did not include the prepupal period in this calculation. There are 6–7 instars, with mean (range) head capsule widths of 0.38 (0.33–0.41), 0.56 (0.49–0.63), 0.90 (0.79–1.01), 1.40 (1.13–1.54), 1.70 (1.64–2.00), 2.3 (2.20–2.50), and 3.01 (2.95–3.26) mm for instars 1–7, respectively (Ogunwolu and Habeck, 1975). Larvae tend to be found singly on blades of grass and drop to the soil surface if disturbed. They are active principally at night. Pupa. Larvae fold the leaves and pupate within. The pupa is covered with a soft, flimsy cocoon. The pupa measures 16–21 mm long and bears a waxy bloom which imparts a whitish or bluish color. Duration of the pupal stage is 6–12 days. Alvarez and Sanchez (1981) reported a mean pupal period of 6.7 days at 30°C.

Egg. The eggs are deposited singly or in small groups of up to five or six eggs on either the upper or lower leaf surface, usually just before midnight. The egg is pale green initially, darkening with age. It is hemispherical, with the base flattened. The egg is marked by the presence of 29–33 ridges, which radiate from the center. It measures about 0.65 mm in diameter and 0.55 mm in height. Duration of the egg stage is 3–5 days at 25–28°C.

FIG. 10.101  Striped grass looper moth. (Photo by L. Buss.)

FIG. 10.100  Striped grass looper. (Photo by J. Castner.)

Adult. The moths are grayish tan or gray in general color, with dark lines and circular markings. Like other Mocis spp., striped grass looper males have a dark spot on the lower margin of the forewing about one-third the distance to the outer margin. However, most specimens of M. latipes bear a dark area or spot at about the midpoint along the

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outer margin of the forewing, a character that helps distinguish M. latipes from the other Mocis spp. Striped grass looper moths superficially resemble adults of velvetbean caterpillar, Anticarsia gemmatalis Hübner. However, the transverse stripe across the forewing of velvet bean caterpillar moths terminates at the apex, whereas in striped grass looper it runs parallel to the outer wing margin (Gregory et al., 1988). The wingspan of striped grass looper moths is 35–40 mm. Adults fly most actively during the early evening hours, though mating frequency is highest near midnight. Adults survive for 10–20  days when provided with food. They typically produce 200–300 eggs. The biology of striped grass looper was summarized by Genung and Allen Jr. (1974), Reinert (1975), and Dean (1985). Larval and pupal descriptions can be found in Ogunwolu and Habeck (1979). Striped grass looper is included in the key to looper pests of vegetables found in Appendix A.

Damage Over 40 species of grasses are fed upon, but other plants seem to be relatively immune to attack. The first and second instars feed on epidermal leaf tissue only, but later instars notch the leaf margins. When abundant, larvae completely defoliate grass plants, leaving only the midribs and stems. Larvae feed at night and remain curled on the soil or in clumps of grass during the day. Although not regularly abundant, during some seasons they can be devastating pests, causing high levels of defoliation.

Management Moths are attracted to light (Gregory et  al., 1988), sugarbased baits (Landolt, 1995), and phenylacetaldhyde-baited traps (Meagher and Mislevy, 2005) though none of these is selective. It is also possible to attract males to pheromone formulations, though velvet bean caterpillar may be attracted to the same chemicals (McLaughlin and Heath, 1989; Landolt and Heath, 1989). Larvae often are difficult to detect because they hide during the day, and their presence is observed only following damage. Grass weeds are often the site of initial infestation in crop fields, so grass weeds should be destroyed to discourage moths from ovipositing within crops. If larval numbers are high or damage is imminent, application of insecticides to crop foliage is recommended.

Sweetpotato Armyworm

Spodoptera dolichos (Fabricius)

Velvet Armyworm

Spodoptera latifascia (Walker) (Lepidoptera: Noctuidae)

Natural History Distribution. These native armyworms are found in ­eastern states, primarily along the Gulf Coast. They are also

found in the Caribbean and in Central and South America. Adults and sometimes larvae of sweetpotato armyworm, Spodoptera dolichos (Fabricius), may occur as far north as Kentucky and Maryland. Velvet armyworm, Spodoptera latifascia (Walker), rarely is numerous outside the Gulf Coast area and is more damaging than sweetpotato armyworm in Florida and Central America. Host Plants. These insects are general feeders, and only occasionally damage vegetables in the United States. Among vegetable crops damaged are asparagus, bean, corn, cowpea, pepper, potato, sweet potato, turnip, and probably others. Sweetpotato armyworm is known to damage cotton, and has also been called the “larger cotton cutworm.” Velvet armyworm also is a common pest of ornamental plants in Florida. In studies conducted in Honduras, velvet armyworm oviposited preferentially on Amaranthus spp. and Ixophorus unisetus weeds relative to corn and sorghum; however, only amaranths, Amaranthus spp., were good hosts for larvae (Portillo et al., 1996b). Other weeds known to sustain larvae are sea purslane, Thrianthema portulacastrum; sena, Cassia leiophila; morningglory, Ipomoea sp.; Melampodium divaricatum; and purslane, Portulaca oleracea (Portillo et al., 1991, 1996a). Natural Enemies. Sweetpotato armyworm is parasitized by Winthemia quadripustulata (Fabricius) (Diptera: Tachinidae). Velvet armyworm is parasitized by Archytas marmoratus (Townsend) and Winthemia sp. (Diptera: Tachinidae), Chelonus sp. (Hymenoptera: Braconidae), Euplectrus plathypenae Howard (Hymenoptera: Eulophidae), and Trichogramma sp. (Hymenoptera: Trichogrammatidae). Velvet armyworm is also known to be infected with a Vairimorpha sp. (Microsporidia: Nosematidae) and a NPV. Telenomus remus Nixon (Hymenoptera: Scelionidae) also can parasitize these armyworm spp. and is easily reared for release (Cave, 2000). An ectoparasitic nematode, Noctuidonema guyanese, parasitizes the adults of both species; it is thought to be debilitating to the moths, but not lethal (Simmons and Rogers, 1996). Life Cycle and Description. The biology of these insects is poorly known. Larvae have been collected from July to November, and “several” broods of S. dolichos are reported to occur annually in Tennessee (Crumb, 1956). Development from the egg to adult stage requires 35– 50 days. When reared at 25°C, mean durations of the egg, larval, prepupal, and pupal stages were about 5.0, 21.3, 3.1, and 20.8 days, respectively (Montezano et al., 2015b). Egg. The egg closely resembles that of yellow-striped armyworm, Spodoptera ornithogalli. It is a slightly flattened sphere, and measures about 0.46 mm in diameter and 0.36 mm in height. The egg bears about 50 narrow ridges that diverge from the center. They are laid in clusters of 200–500 eggs on the undersides of leaves, and bear scales from the

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female’s abdomen. Apparently, females produce 500–3000 eggs, although under laboratory conditions Montezano et al. (2015a) reported mean fecundity of S. dolichos to be over 4000 eggs and mean fertility (number of eggs hatching) to be about 3500 eggs. Duration of the eggs stage is 4–8 days.

FIG. 10.102  Sweetpotato armyworm caterpillar. (Photo by L. Buss.)

FIG. 10.103  Sweetpotato armyworm moth. (Photo by L. Buss.)

Larva. Larvae are gregarious during the early instars but disperse thereafter. There likely are 5–7 instars in both species. Crumb (1929) gave head capsule widths of 0.27, 0.41, 1.0–1.1, 1.5–1.6, 2.2–2.5, and 3.1–3.3 mm for instars 1–6 of S. dolichos. Mean duration of the larval stage when reared at 25°C was about 4.1, 3.0, 3.2, 3.4, 4.2, and 3.4 days for instars 1–6, respectively, followed by a prepupal period of about 3.1 days (Montezano et al., 2015b.). Santos et al. (1960) indicated head capsule widths of 0.3, 0.45, 0.75, 0.95, 1.4, 2.20, and 3.25 mm for instars 1–7 in S. latifascia. Duration of the larval stage is reported to be 23–30 days in S. dolichos and 15–27 days in S. latifascia. Instar-specific development time is reported to be about 3, 4, 4, 4, 3, 4, and 5  days for instars 1–7, respectively, in S. latifascia. Larvae of S. latifascia completed their larval period in about

15  days when fed cotton, and 18  days when soybean; no larvae completed development on lettuce. Larvae are variable, ranging from light gray or green to blackish, and wellmarked with spots and stripes. Larvae bear prominent dark triangular spots subdorsally along the abdomen, consistent with many other Spodoptera spp., and lateral yellowish lines are usually present both above and, to a lesser extent, below the spiracles. The subspiracular yellowish line is not interrupted by a spot on the first abdominal segment, as is usually the case with southern armyworm, S. eridania, but the supraspiracular line does bear a dark spot. Dark subdorsal markings found on the mesothorax are small and semicircular in S. latifascia but large and trapezoidal in S. dolichos. In contrast, the mesothoracic markings of yellowstriped armyworm, with which these species are easily confused, are triangular. Larvae attain body lengths of 43 and 48 mm in S. dolichos and S. latifascia, respectively, which makes them quite large for the genus. Pupa. Larvae pupate in the soil. The pupal stage is dark brown and measures 20–30 mm long. Duration of the pupal stage is 9–20 days.

FIG. 10.104  Velvet armyworm caterpillar. (Photo by L. Buss.)

FIG. 10.105  Velvet armyworm moth. (Photo by L. Buss.)

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Adult. The adults are grayish-brown moths with heavily mottled forewings. The forewing of sweetpotato armyworm usually bears an irregular orangish streak distally. Also, the thoracic region is marked with two broad, dark, subdorsal stripes running from the head to the abdomen; the moths of velvet armyworm and other common Spodoptera spp. lack these stripes. In both species, the hind wing is white, with a narrow dark band along the distal edge. The wingspan of these moths is 40–50 mm. The pre-, post-, and oviposition period in S. dolichos when reared at 25°C was reported to be 3.0, 0.4, and 10.4  days, and total adult longevity was 12.9  days in females and 12.4 days in males (Montezano et al., 2015a). Descriptions of sweetpotato armyworm are found in Crumb (1929, 1956). Velvet armyworm was described by Levy and Habeck (1976), developmental biology was given by Habib et  al., 1983, and the sex pheromone discussed by Monti et al. (1995). Larvae of both species were described briefly by Passoa (1991), and keys for many Spodoptera were presented by Levy and Habeck (1976), Passoa (1991), and Heppner (1998). These species also occur in a key to Louisiana noctuids (Oliver and Chapin, 1981), in Stehr (1987), and in a key to armyworms and cutworms in Appendix A. Keys to adults were presented by Todd and Poole (1980) and Heppner (1998). King and Saunders (1984) discussed biology in Central America.

Damage Larvae are defoliators, and because of their large size inflict considerable damage late in the larval stage. They can function as cutworms, severing young plants at the soil surface, and also may burrow into tomato and other soft fruits.

Management Larvae are most often controlled by the application of foliar insecticides. These insects are not frequent pests, however. In Central America, velvet armyworm sometimes feeds on corn and sorghum, but if weeds are present the larvae of this insect feed preferentially on them. Corn and sorghum, because they are not suitable for larval development, serve as sink habitat, whereas certain broadleaf weeds serve as a source habitat (Portillio et al., 1991).

Sweetpotato Leaf Folder

Lygropia tripunctata (Fabricius) (Lepidoptera: Crambidae)

Natural History Distribution. This species is known from Hawaii, Texas, and east to Florida and South Carolina. It is also recorded from the Caribbean and Central American regions, south to Brazil. Host Plants. This plant is found to feed mostly on the genus Ipomoea (Convolvulaceae), both wild and ­cultivated

varieties. In addition, it feeds on Turbina corymbosa (Linnaeus) and Merremia umbellata (Linnaeus) (both Convolvulaceae). Natural Enemies. Exorista pyste Walker (Diptera: Tachinidae) are known to attack the larvae, as are adults of Podisus maculventris Say (Hemiptera: Pentatomidae). Life Cycle and Description. This species apparently undergoes at least four, and sometimes five, generations per season. Each generation requires at least 25  days for completion, consisting of 4 days in the egg stage, 13 days in the larval stage, 2 days in the prepupal stage, and 6 days in the pupal stage. The eggs require about 3 days for development before they are laid. The biology is given by Jones (1917a, b). Egg. The egg is elliptical in shape. Initially, it is colorless, though later in development the embryo is evident. The eggs measure a mean length of 1.00 mm whereas the width is 0.79 mm. The eggs are deposited singly and in small groups of up to 5 per egg clutch. The eggs are typically found on the leaf tissue. Duration of the egg stage is about 4 days. Larva. Hatching larvae are about 1.5 mm long. At this stage, larvae are nearly colorless. After ingesting chlorophyll, they acquire a green color, and eventually may be brownish or bluish green. The head is pale yellow. Eventually, the larvae attain a length of about 27 mm. The larva typically constructs a shelter composed of the leaf material that shortens as the silk dries. Initially, the young larva creates small holes, but as the larva increases in size, the entire leaf may disappear. The larval stage persists for 13–14  days. Duration of the larval instars was about 3, 2, 2, 2, 2, and 6 days each for instars 1–6, respectively. Pupa. The pupa is dark brown in color. They measure about 15 mm in length, and 4 mm in width. Pupation requires about 6–9 days. Adult. The upper color of the moth is light yellow, with the wings iridescent. A dark-grayish-brown band occurs along the periphery of the wings, especially along the margin of the forewings. Two brown spots are located along the costal margin of each forewing, and a small black spot is found on the hind wings. A wavy dark line is found at about the midpoint of the fore- and hindwings. The measurements of the male and female are about 27 mm (range of 25–29 mm) and about 25–27 mm, respectively. Adults are active in the United States from March to October.

Damage Damage results from feeding by the larvae of sweetpotato leaf folder on the leaf tissue.

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Management The larvae can become quite numerous, and liquid insecticide is often applied to disrupt their development.

Tobacco Budworm

Chloridea virescens (Fabricius) (Lepidoptera: Noctuidae)

Natural History Distribution. Tobacco budworm is a native species and is found throughout the eastern and southwestern United States. This species has a recent name change, so most of the literature refers to it as Heliothis virescens. Like its close relative corn earworm, Helicoverpa zea (Boddie), it generally overwinters successfully only in southern states. However, it occasionally survives in cold climates in greenhouses and other sheltered locations. Tobacco budworm disperses northward annually and can be found in New England, New York, and southern Canada during the late summer. It also occurs widely in the Caribbean, and sporadically in Central and South America. Occasionally, tobacco budworm in found in the Pacific Northwest of United States. Host Plants. Tobacco budworm is principally a fieldcrop pest, attacking such crops as alfalfa, clover, cotton, flax, soybean, and tobacco. Tobacco budworm is also a common pest of geranium and other flower crops such as ageratum, bird of paradise, chrysanthemum, gardenia, geranium, petunia, mallow, marigold, petunia, snapdragon, strawflower, verbena, and zinnia. However, it sometimes attacks such vegetables as cabbage, cantaloupe, lettuce, pea, pepper, pigeon pea, squash, and tomato, especially when cotton or other favored crops are abundant. Weeds serving as a host for larvae include beardtongue, Penstemon laevigatus; beggarweed, Desmodium spp.; bicolor lespedeza, Lespedeza bicolor; black medic, Medicago lupulina; cranesbill, Geranium dissectum; deergrass, Rhexia spp.; dock, Rumex spp., groundcherry, Physalis spp.; Japanese honeysuckle, Lonicera japonica; lupine, Lupinus spp.; morning glory, Ipomoea spp.; a morning glory, Jacquemontia tamnifolia; passionflower, Passiflora sp.; prickly sida, Sida spinosa; sunflower, Helianthus spp.; toadflax, Linaria canadensis; and velvetleaf, Abutilon theophrasti (Brazzel et al., 1953; Neunzig, 1963; Graham and Robertson, 1970; Roach, 1975; Harding, 1976a; Stadelbacher, 1981; Pair, 1994; Sudbrink Jr. and Grant, 1995). In Georgia, Barber (1937) determined that tobacco budworm developed principally on toadflax during April-May for one to two generations, followed by one generation on deergrass during June-July and two to three generations on beggarweed during JulyOctober. In Mississippi, cranesbill was identified as the key early-season host plant (Stadelbacher, 1981). In southern Texas, cotton is the principal host, but such weeds as

wild tobacco, Nicotania repanda; vervain, Verbena neomexicana; ruellia, Ruellia runyonii; and mallow, Aubitilon trisulcatum, are important hosts early or late in the year (Graham et al., 1972). In cage tests and field studies conducted in Florida and which did not include cotton, tobacco was more highly preferred than other field crops and vegetables, but cabbage, collards, okra, and tomato were attacked (Martin et al., 1976a). Natural Enemies. Numerous general predators have been observed to feed upon tobacco budworm. Among the most common are Polistes spp. wasps (Hymenoptera: Vespidae); big-eyed bug, Geocoris punctipes (Say) (Hemiptera: Lygaeidae); damsel bugs, Nabis spp. (Hemiptera: Nabidae); minute pirate bugs, Orius spp. (Hemiptera: Anthocoridae), and spiders. Several parasitoids have also been observed, and high levels of parasitism have been reported (Lewis and Brazzel, 1968; Tingle et al., 1994). The egg parasitoid Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae) can be effective in vegetable crops. Other important parasitoids are Cardiochiles nigriceps Viereck in vegetables and Cotesia marginiventris (Cresson) in other crops (both Hymenoptera: Braconidae). Effectiveness of the parasitoids varies among crops (Martin et  al., 1981a, b). Other species known from tobacco budworm include Archytas marmoratus (Townsend) (Diptera: Tachinidae); Meteorus autographae Muesebeck (Hymenoptera: Braconidae); Campoletis flavicincta (Ashmead), C. perdistinctus (Viereck), C. sonorensis (Cameron), Netelia sayi (Cushman) and Pristomerus spinator (Fabricius) (all Hymenoptera: Ichneumonidae). Pathogens are also known to inflict mortality. Among the known pathogens are fungi such as Nosema spp. and Nomuraea rileyi, and NPVs. In a study conducted in South Carolina, Nomuraea was a more important mortality agent than the natural incidence of virus and was considered to be one of the most important natural mortality agents (Roach, 1975). Life Cycle and Description. Moths emerge during March-May in southern states, followed by four to five generations through the summer, with overwintering commencing in September-November. Four generations have been reported from northern Florida (Chamberlin and Tenhet, 1926) and North Carolina (Neunzig, 1969), and at least five from Louisiana (Brazzel et al., 1953). Moths have been collected in New York in July-September, but at such northern latitudes it is not considered to be a pest (Chapman and Lienk, 1981). This species overwinters in the pupal stage. Egg. The eggs are deposited on blossoms, fruit, and terminal growth. They are spherical, with a flattened base. They measure 0.51–0.60 mm wide and 0.50–0.61 mm long. They initially are whitish to yellowish white, turning gray as they age. Narrow ridges radiate from the apex of the egg, and number 18–25. Eggs of tobacco budworm are nearly

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indistinguishable from those of corn earworm. At high magnification, however, the primary ribs of tobacco budworm eggs can be observed to terminate before they reach the rosette of cells surrounding the micropyle; in corn earworm, at least some primary ribs extend to the rosette (Bernhardt and Phillips, 1985). Females normally produce from 300 to 500 eggs, but fecundity of 1000–1500 eggs per female have been reported from larvae cultured on artificial diet at cool temperature (Fye and McAda, 1972).

microspines on the first, second, and eighth abdominal segments that are about half the height of the tubercles. In corn earworm the microspines on the tubercles are absent or up to one-fourth the height of the tubercle. Larvae exhibit cannibalistic behavior starting with the third or fourth instar, but are not as aggressive as corn earworm. Pupa. Pupation occurs in the soil. Pupae are shiny reddish brown, become dark brown before the emergence of the adult. The pupa averages 18.2 mm long and 4.7 mm wide. Duration of the pupal stage is reported to be about 22 days at 20°C, 13.0 days at 25°C, and 11.2 days at 30°C. Diapause is initiated by either low temperature or short day length (Henneberry et al., 1993; Henneberry, 1994).

FIG. 10.106  Larva of tobacco budworm. (Photo by L. Buss.)

Larva. Tobacco budworm larvae have 5–7 instars, with five or six most common. Head capsule widths for larvae that develop through five instars measure 0.26–0.31, 0.46–0.54, 0.92–0.99, 1.55–1.72, and 2.38–2.87 mm for instars 1–5, respectively. Larval lengths are 1.1–4.0, 4.2–8.0, 8.7–14.7, 18.5–25.6, and 23.3–35.6 mm for these same instars. Head capsule widths for larvae that develop through six instars measure 0.26–0.31, 0.36–0.53, 0.72–0.85, 1.12–1.25, 1.60– 1.72, and 2.40–2.82 mm for instars 1–6, respectively. Larval lengths are 1.4–4.1, 3.0–7.0, 7.5–9.2, 12.0–15.8, 19.5–24.3, and 25.5–36.0 mm for these same instars. Development time was studied by Fye and McAda (1972) at various temperatures. When cultured at 20°C, development required about 4.6, 2.6, 3.1, 3.7, 10.1, and 9.8 days for instars 1–6, respectively. At 25°C, larval development times were 3.1, 2.0, 1.9, 2.1, 5.7, and 2.5 days, respectively. Young larvae are yellowish or yellowish green with a yellowish-brown head capsule. Later instars are greenish with dorsal and lateral whitish bands, and with a brown head capsule. Many of the bands may be narrow or incomplete, but a broad, lateral subspiracular band is usually pronounced. Body color is variable, and pale green or pinkish forms, or dark reddish or maroon forms are sometimes found. Larvae are very similar to corn earworm. As in corn earworm, its body bears numerous black thorn-like microspines. These spines give the body a rough feel when touched. Early instars are difficult to separate from corn earworm; Neunzig (1964) gave distinguishing characteristics. Starting with the third instar, close examination reveals tubercles with small thorn-like

FIG. 10.107  Tobacco budworm moth. (Photo by L. Buss.)

Adult. The moths are brownish, and lightly tinged with green. The front wings are crossed transversely by three dark bands, each of which is often accompanied by a whitish- or cream-colored border. Females tend to be darker. The hind wings are whitish, with the distal margin bearing a dark band. The moths measure 28–35 mm in wingspan. The preoviposition period for females is about 2-days long. The longevity of moths is reported to range from 25 days when held at 20°C, to 15 days at 30°C. A sex pheromone has been identified (Tumlinson et al., 1975). The biology of tobacco budworm was given by Neunzig (1969) and Brazzel et  al. (1953). The larva was included in keys by Okumura (1962), Stehr (1987), Sparks and Liu (2001), and Oliver and Chapin (1981); the latter publication also pictured the adult stage. Tobacco budworm is also included in a key to armyworms and cutworms in Appendix A. Larvae are readily cultured on bean-based rearing media or other diets (King and Hartley, 1985).

Damage Larvae bore into buds and blossoms (the basis for the common name of this insect), and sometimes the tender ­terminal foliar growth, leaf petioles, and stalks. In the absence of reproductive tissue, larvae feed readily on foliar tissue. Neunzig (1969)

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infested tobacco with both tobacco budworm and corn earworm and observed very similar patterns and levels of injury by these closely related species. In California, both budworm and earworm burrow into the heads of developing lettuce. The entry of larvae into fruit increases frequency of plant disease. Research in southern Arkansas tomato fields indicated that though tobacco budworm was present from May to July, they were not nearly as abundant or damaging as corn earworm (Roltsch and Mayse, 1984).

Management Sampling. Large cone-shaped wire traps baited with sex pheromone lures are commonly used to capture tobacco budworm moths (Hartstack et  al., 1979). Smaller bucket traps can be used to capture these moths, but they are not very efficient (Lopez Jr. et al., 1994). Insecticides. Foliar insecticides are commonly used in crops where tobacco budworm damage is likely to occur. However, the destruction of beneficial organisms often results, and this is thought to exacerbate budworm damage. Also, resistance to insecticides is widespread, particularly in crops where pyrethroid use is frequent (Kanga et al., 1995; Greenstone, 1995). Larvae also consume bait formulated from cornmeal and insecticide (Creighton et al., 1961). Cultural Techniques. Early season destruction of weeds with herbicide or mowing, or destruction of larvae on the weeds by treatment with insecticides, can reduce tobacco budworm population size later in the year (Bell and Hayes, 1994; Snodgrass and Stadelbacher, 1994). Biological Control. The microbial insecticide Bacillus thuringiensis is effective against budworm (Johnson, 1974; Stone and Sims, 1993). Heliothis NPV has been used effectively to suppress tobacco budworm on field crops (Andrews et  al., 1975) and on early season weed hosts (Hayes and Bell, 1994). Tobacco budworm is also susceptible to NPV from alfalfa looper, Autographa californica (Speyer) (Vail et al., 1978; Bell and Romine, 1980). Release of Trichogramma egg parasitoids has been shown to be beneficial in some vegetable crops (Martin et al., 1976b). Host-Plant Resistance. Although there is little evidence for natural resistance to tobacco budworm among many crops, cotton has been genetically engineered to express resistance (Benedict et al., 1996). Enhanced resistance to larval survival by cotton results in lower insect pressure on nearby vegetable crops.

Variegated Cutworm

Peridroma saucia (Hübner) (Lepidoptera: Noctuidae)

Natural History Distribution. Variegated cutworm is found in many areas of the world. It occurs throughout the western ­hemisphere, and in Hawaii, and also portions of Europe,

Asia, and North Africa. The origin of this insect is uncertain, but is thought to be Europe, where it was described in 1790. First observed in North America in 1841, it is now abundant in southern Canada and the northern United States, where it is often considered to be the most damaging cutworm pest of vegetables. Host Plants. This cutworm has an extremely wide host range. Unlike some cutworms that expand their dietary range only when confronted with overpopulation and starvation, variegated cutworm feeds readily on numerous plants. Among vegetables attacked are asparagus, bean, beet, Brussels sprouts, cabbage, cantaloupe, carrot, cauliflower, celery, Chinese cabbage, collards, corn, cowpea, cucumber, garbanzo, globe artichoke, kale, lettuce, lima bean, mustard, onion, pea, pepper, potato, sweet potato, Swiss chard, radish, rhubarb, rutabaga, spinach, squash, tomato, turnip, and watermelon (Rings et  al., 1976b). Based on the frequency of reports, variegated cutworm is most likely to be found damaging beet, cabbage, lettuce, potato, and tomato. Variegated cutworm is also known to damage fruit trees, including apple, apricot, avocado, cherry, currant, gooseberry, grape, lemon, mulberry, orange, plum, raspberry, and strawberry. Other crops injured include alfalfa, barley, clover, corn, cotton, flax, hops, mint, sunflower, sugarbeet, sweet clover, tobacco, wheat, and many flower crops. Weeds are occasionally consumed but do not seem to be preferred. Some of the weeds eaten are jimsonweed, Datura sp.; dock, Rumex sp.; dogfennel, Eupatorium capillifolium; plantain, Plantago sp.; ragweed, Ambrosia sp.; and shepherdspurse, Capsella bursa-pastoris. Natural Enemies. Numerous natural enemies are known for variegated cutworm. Walkden (1950) reported mortality of variegated cutworm over a 20-year period in the central Great Plains, and frequently observed 20%–75% mortality, with wasp and fly parasitoids accounting for most of the deaths among larvae. In a study of cutworms in Oklahoma, Soteres et  al. (1984) reported six species of Braconidae, three species of Ichneumonidae, and one species of Eulophidae, but among the Hymenoptera only Ophion sp. (Ichneumonidae) accounted for more than 5% mortality. In the same study, 12 species of Tachinidae were observed, but among the Diptera only Archytas apicifer (Walker) and Peleteria ­texensis Curran (both Tachinidae) accounted for more than 5% m ­ ortality. In Oregon, Coop and Berry (1986) reported eight species of Hymenoptera from variegated cutworm larvae. The parasitoid Meteorus communis (Cresson) (Braconidae), was recovered from about 35% of the intermediate-age larvae. Not only did parasitism affect abundance of cutworm in subsequent generations, but caused a 93% reduction in foliage consumption by parasitized larvae. A study of variegated cutworm in Hawaii demonstrated 33–80% parasitism due to Hyposoter exiguae (Viereck) (Ichneumonidae), Cotesia marginiventris (Cresson) and Meteorus laphygmae Viereck (both Braconidae) (all Hymenoptera) (Hara and Matayoshi, 1990).

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Other Hymenoptera known to parasitize variegated cutworm include Apanteles xylinus (Say), Chelonus ­insularis Cresson, C. militaris (Walsh), Meteorus autographae Muesebeck, M. leviventris (Wesmael), Microplitis feltiae Muesebeck, Rogas perplexus Gahan, and R. rufocoxalis Gahan (all Braconidae); Campoletis sonorensis (Cameron), Enicospilus merdarius (Gravenhorst), Nepiera fuscifemora Graf, and Ophion flavidus Brulle (all Ichneumonidae); and Dibrachys canus (Walker) (Pteromalidae). Other Diptera parasitizing this cutworm are Archytas aterrimus (Robineau-Desvoidy), A. cirphis Curran, Bonnetia comta (Fallén), Carcelia spp., Chaetogaedia monticola (Bigot), Clausicela opaca (Coquillett), Eucelatoria armigera (Coquillett), Euphorocera claripennis (Macquart), E. omissa (Reinhard), Gonia longipulvilli Tothill, G. porca Williston, G. sequax Williston, Lespesia archippivora (Riley), Madremyia saundersii (Williston), Peleteria texensis (Curran), Periscepsia helymus (Walker), P. laevigata (Wulp), Voria ruralis (Fallén), Winthemia leucanae (Kirkpatrick), W. quadripustulata (Fabricius), and W. rufopicta (Bigot) (all Tachinidae). The fungus Metarhizium anisopliae and viruses can inflict mortality, but the incidence is often low. Both a granulosis (Steinhaus and Dineen, 1960) and a NPV (Harper, 1971) are known to affect this cutworm. Life Cycle and Description. There are two to four generations annually, with two generations common in northern states and southern Canada, and 3–4 in southern areas. Flights of moths are protracted, and generations are difficult to discern based on adult populations. Overwintering may occur in the pupal stage, or perhaps the larval stage, but there is also evidence that moths migrate into northern areas from southern latitudes each spring, and return to the south in the autumn. In Iowa, 3%–9% of moths collected during the spring flight were considered to have emerged from overwintering pupae, with the balance migrating from southern latitudes (Buntin et  al., 1990). The AprilMay spring flight in Iowa tends to be followed by generations and adult flights in June-July, August-September, and October-November. The total duration of the life stages is usually 35–70  days. Overwintering in Canada is unlikely (Ayre, 1985) except in relatively warm areas such as coastal British Columbia. Egg. The egg of variegated cutworm is hemispherical; the egg is flattened at the point of attachment to a leaf or plant stem. The surface of the egg is marked with ridges, about 42 in number, radiating from the center. The egg measures about 0.55–0.58 mm in diameter and 0.40–0.45 mm in height. Initially, they are white but soon turn brownish. The developmental threshold for the egg stage is estimated at 3.0–6.0°C by various authors. Duration of the egg stage

is 4–6 days in warm weather (20–30°C), but 10 days when held at 15°C. Eggs are deposited in clusters, often numbering several hundred per egg mass. Females may deposit 1200–1400 eggs during their lifespan.

FIG. 10.108  Larva of variegated cutworm. (Photo by L. Buss.)

Larva. There are normally six instars. The developmental threshold for the larval stage was estimated at 2.6–6.7°C by various authors. Mean duration of the instars is reported to be 6.5, 4.6, 4.8, 4.7, 6.7, and 16.8 days for instars 1–6, respectively, at 15°C. When reared at 25°C, instar durations are reduced to 3.1, 1.9, 2.2, 2.2, 2.9, and 8.4 days, respectively (Shields, 1983). Head capsule widths are 0.30–0.35, 0.46–0.62, 0.80–1.00, 1.20–1.65, 1.9–2.6, and about 3.0–3.2 mm, respectively. Body lengths are estimated at 2.0–3.0, 3.6–6.5, 5.3–9.0, 12–16, 25–28, and 35–46 mm for instars 1–6, respectively. Body color is brownish gray to grayish black. The most distinctive character is a dorsal yellow or whitish spot, which is present on each of the first four abdominal segments, often on the first six segments, though this character may be absent in early instars. Less distinctive is the black “W”-shaped mark on the eighth abdominal segment of the last instar, and light brown or tan on the posterior end of the body. An inconspicuous black line is often present laterally above the spiracles. An orangish-brown line may connect the spiracles, and below the spiracles there is usually some irregular yellowish or orangish coloration. The head is orangish brown and marked with darker spots. Pupa. The larva forms a cell in the soil and pupates near the soil surface. The pupa is mahogany brown, and measures 15–23 mm long and 5–6 mm wide. The developmental threshold of the pupal stage was estimated at 4.3–8.5°C by various authors. Duration of the pupal stage is about 33 and 13 days at 15°C and 25°C, respectively.

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FIG. 10.109  Adult of variegated cutworm. (Photo by L. Buss.)

Adult. The adult is fairly large in size, measuring 43–50 mm wingspan. The front wings are normally grayish brown, tinged with reddish and shaded centrally and distally with darker brown. The background color varies, however, from dark brown to yellowish brown. A bean-shaped spot and a smaller round spot are usually evident centrally. The hind wings are iridescent or pearly white with brown veins and brown shading marginally. The head and thorax are dark brown, whereas the abdomen is lighter brown. Females produce a sex pheromone to attract males (Struble et al., 1976). Oviposition commences 7–14 days after emergence of the adults. Description and biology of variegated cutworm were given by Chittenden (1901), Crumb (1929), and Walkden (1950). Developmental biology was given by Simonet et al. (1981) and Shields (1983). Rearing methods were described by Finney (1964) and Harper (1970). A bibliography was published by Rings et al. (1976a). The larva was included in keys by Whelan (1935), Crumb (1956), Okumura (1962), Rings (1977b), Oliver and Chapin (1981), Capinera (1986), and Stehr (1987). It is also included in a key to armyworms and cutworms in Appendix A. The moth was included in pictorial keys by Rings (1977a) and Capinera and Schaefer (1983).

Damage The larvae cause considerable mortality to seedlings by cutting off the plant at the soil surface. Larvae are also defoliators, and though they commonly frequent low-growing herbage, they readily climb trees to feed on buds and foliage. Young larvae may remain on the foliage during the daylight hours, but feed principally during the evening hours. Large larvae often hide in the soil or other sheltered locations during the daytime hours, moving to exposed areas of foliage in the evening to feed. Larvae may burrow into tomato fruit and the heads of cabbage and cauliflower.

At high densities larvae may assume a gregarious, dispersive “armyworm” habit, but this is uncommon. It also invades greenhouses frequently. The larvae consume about 125 cm2 of sugarbeet or 160 cm2 of potato foliage during their larval development (Capinera, 1978c; Shields et  al., 1985). Potato and many other plants can tolerate some defoliation without significant yield decrease. Late in the season, after tuber formation is nearly complete, potato can withstand up to 75% leaf loss without yield decrease. At this time up to 40 variegated cutworms per plant can be tolerated. However, at an earlier period in the season such as at full bloom, plants may be able to tolerate only three cutworms per plant (Shields et al., 1985).

Management Sampling. The adult populations can be monitored with blacklight and pheromone traps. Captures by the two types of traps are correlated, but pheromone traps capture larger numbers from the spring generation, whereas blacklight traps capture larger numbers at other times of the year (Willson et  al., 1981). Pheromone traps, though not completely species-specific (Ayre et  al., 1983), provide considerable selectivity relative to blacklight traps, thereby reducing labor requirements associated with population monitoring. Larval populations are difficult to sample. Young larvae may be found clustered on foliage, but older larvae tend to hide in sheltered locations or burrow beneath the surface of the soil during the daylight hours. If plants are severed at the soil surface, or have disappeared, it is important to rake the soil surface and search for cutworm larvae. Insecticides. Insecticides are commonly recommended to protect young plants from cutting damage, and older plants from defoliation and fruit injury. Insecticide applications are directed at the foliage or soil, the latter because the larvae often seek shelter there. Insecticides differ greatly in their effectiveness, and larger larvae are considerably more difficult to kill (Harris et al., 1977). Insecticide-treated bran baits are effective against variegated cutworm. Bacillus thuringiensis is not recommended. Variegated cutworm larvae are sensitive to neem products, which act as feeding deterrents and disrupt larval growth and survival (Koul and Isman, 1991; Isman, 1993). Cultural Practices. Cutworm problems often develop in weedy fields or portions of fields infested with weeds. It is advisable to till, or otherwise destroy weeds, 10–14 days in advance of planting, as this should cause small larvae to starve. If seedlings are to be transplanted into a field or garden, larger plants are preferred, because they are less likely to be irreparably damaged by cutworms. Transplanted plants derive considerable protection if surrounded by a

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barrier such as a can or waxed paper container with the top and bottom removed. Aluminum foil wrapped around the base of the seedling also deters cutting by larvae. Biological Control. Variegated cutworm is susceptible to infection by entomopathogenic nematodes (Nematoda: Steinernematidae and Heterorhabditidae) (Morris and Converse, 1991), but the demonstration of practical use under field conditions is lacking.

Western Bean Cutworm

Striacosta albicosta (Smith) (Lepidoptera: Noctuidae)

late summer, and mature larvae overwinter. Pupation occurs in May or June of the following year. Egg. The eggs are deposited on the upper surface of leaves and within the whorl of corn plants. They are nearly spherical but flattened at the point of attachment to the foliage. They are white with a thin red line around the top initially and turn pinkish or purplish gray at maturity. The eggs are about 0.7 mm in diameter, and well-marked with ridges that radiate from the center. The mean number of eggs per cluster is reported to be 52 (range 21–195 eggs). The incubation period is 4–7 days.

Natural History Distribution. Historically, this native cutworm is found in the Rocky Mountain and western Great Plains regions of the United States and Canada. However, in recent years its geographic range has extended east, attaining Pennsylvania, New York, and Quebec in 2009 (Michel et  al., 2010). Its range expansion is possibly due to adoption by this insect of corn as a food plant in the 1950s. Western bean cutworm appears to be absent from the Pacific Coast States, but it also affects corn in Mexico. Host Plants. Originally known as a minor pest of bean, and perhaps of tomato, the host range of western bean cutworm apparently expanded during the 1950s to include corn. Acreage in the western states devoted to bean and corn also expanded during this period, so it is not clear whether host range expansion occurred, or whether damage simply became more noticeable as the availability of suitable hosts increased. It is now considered to be a locally important pest of grain (dent) corn and dry beans in western states, but increasingly injures sweet corn, snap beans, dry beans, and rarely tomato in the Midwest and southern Canada. Other legume crops may receive eggs and support partial larval development but are considered to be poor hosts. Weed hosts include the fruit of groundcherry, Physalis spp., and black nightshade, Solanum nigrum, though these are most suitable after larvae are partly grown (Blickenstaff and Jolley, 1982). Natural Enemies. Few natural enemies have been reported, though this is more likely due to lack of study than the absence of predation and parasitism. The wasp Apanteles laeviceps Ashmead (Hymenoptera: Braconidae) has been reared from western bean cutworm. Common predators such as lady beetles (Coleoptera: Coccinellidae), minute pirate bugs (Hemiptera: Anthocoridae), damsel bugs (Hemiptera: Nabidae); big-eyed bugs (Hemiptera: Lygaeidae), green lacewings (Neuroptera: Chrysopidae), and spiders consume western bean cutworm eggs and larvae under laboratory conditions (Blickenstaff, 1979). Life Cycle and Description. There is a single generation per year. Adults emerge in July and early August, with eggs found throughout this period. Larvae develop during

FIG. 10.110  Western bean cutworm larva. (Photo by J. Capinera.)

Larva. Upon hatching, larvae remain near the egg mass for several hours and then disperse, often to the tassel of corn where they feed on pollen. Larvae normally feed at night, but they can be observed feeding during the daylight hours if it is cloudy. On small plants, particularly on beans, larvae may move to the soil during the daylight hours to seek shelter. The larvae initially are brownish and marked with weak stripes. As they mature they lose the stripes and become pinkish brown dorsally, and grayish laterally. The thoracic plate bears broad dark brown or black stripes. The number of instars varies from 5 to 8, but six is normal. Larval head capsule width is 0.3–0.5, 0.5–0.8, 0.8–1.0, 1.1–1.7, 1.8–2.5, and 2.7–4.1 mm, for instars 1–6, respectively. Larval lengths are 1.9–3.3, 3.6–7.5, 8.2–11.5, 16.0–19.2, 19.0–27.2, and 28.0–37.1 mm for instars 1–6 respectively. Mean development time requires 31 days (range 21–45 days). The mature cutworm measures about 35 mm long. Pupa. The larvae burrow into the soil in the autumn, mostly to a depth of 7–15 cm, and create a small pupal cell from saliva and soil particles. However, they remain in the larval stage until late May or June, when pupation occurs. The pupa is dark brown and measures about 20 mm long.

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the seed pod at several locations. When not feeding, large larvae may seek shelter in the soil. The level of damage to dry beans historically has been low, but now that this insect has become established in the eastern United States and Canada, snap (green) beans are more available, and they are more easily damaged than dry beans. Goudis et al. (2016) report that insectical control is much more justified on snap beans than on dry beans.

Management FIG. 10.111  Adult of western bean cutworm. (Photo by J. Capinera.)

Adult. The moth is brownish, with a wingspan of 35–40 mm. The forewing is distinctively marked with a broad whitish or tan stripe along the anterior margin. The center of the forewing is generally well-marked with a small round spot basally and a bean-shaped spot distally. The spots are white or margined with white, and connected by a short black bar. The hind wing is white, but with delicate brown lines at the margin and along the veins. Adults live for a relatively brief period, averaging 7.2 days for males and 9.2 days for females. The preoviposition period is about 4 days, with females depositing about 400 eggs. The biology was described by Hoerner (1948), Hagen (1962, 1976), Blickenstaff (1979), and Michel et al. (2010). A sex pheromone has been described (Klun et  al., 1983). Western bean cutworm was included in the larval keys by Crumb (1956), Capinera (1986), and Stehr (1987), and is included in a key to armyworms and cutworms in Appendix A. The adult was included in the key by Capinera and Schaefer (1983).

Damage On corn, larvae feed on developing pollen and tassels. As the tassel matures, larvae disperse and feed on pollen that has collected on the foliage and on the leaves. If pollen is not available, larvae feed on corn silk. As ears develop, larvae feed beneath the leaf sheath on both silk and kernels. Unlike corn earworm, Helicoverpa zea (Boddie), which tends to attack the ear tip, larvae of western bean cutworm are likely to burrow randomly into the ear, attacking all areas. Also, western bean cutworm larvae, unlike corn earworm, are not cannibalistic, so several larvae may be found feeding on a single ear. Leaf and stalk feeding are trivial, but silk feeding can inhibit pollination. Damage to corn also allows entry of plant pathogens. Feeding by western bean cutworm significantly increases the development of Gibberella ear rot, Fusarium graninearum (Schwein.) (Parker et al., 2016). When feeding on bean, larvae initially remain near the top of the plant, feeding on buds and young leaves. Larger larvae feed on pods but usually do not burrow into the pod. Instead, they wander and feed on developing beans through

Sampling. Moths can be captured with both blacklight and pheromone traps. Blacklight traps capture moths earlier in the season than pheromone traps, but both trap types correlate well with damage potential. Although trap captures and overall damage potential are correlated, there is high among-field variation in damage (Mahrt et al., 1987). Initiation of moth capture in traps should trigger field scouting for eggs. Pretassel corn is preferred by moths, and infestation is patchy, so thorough scouting is advisable. Paula-Moraes et al. (2011) provided egg cluster sequential sampling plans that reduce the number of samples needed for decision making. The number of samples needed varies with the incidence of egg clusters, but generally, only 40–55 plants need to be sampled. Insecticides. Application of insecticides to foliage is effective, though it is important to time the application early in the larval period, so applications are usually timed shortly after peak moth flight. If moth populations are not being monitored, insecticide should be applied before all the corn tassels have emerged from the whorl. Applications of insecticide-treated bait and soil-applied systemic insecticides are not very effective (Hoerner, 1948; Blickenstaff and Peckenpaugh, 1981). Cultural Practices. Timing of planting affects the susceptibility of corn to western bean cutworm infestation. Moths oviposit preferentially on plants that are silking, avoiding early and late-planted corn. The emergence of moths from the soil is enhanced by high moisture conditions and high sand content; the rate of emergence from heavy and dry soil is low. Host-Plant Resistance. Transgenic Bt-based corn normally escapes damage by Lepidoptera, but western bean cutworm seems to be unaffected by Cry1A, Cry1Ab, and Cry2Ab corn hybrids, whereas European corn borer, Ostrinia nubilalis (Hübner) and corn earworm, Helicoverpa zea (Boddie), are adversely affected by Cry1Ab hybrids. This is a problem for the production of both sweet corn and dent (grain) corn. In some areas, western bean cutworm is moderately susceptible to Cry1F protein, but reports from Ontario, Canada indicate an insensitivity to Cry1F hybrids, so this source of resistance may fail as well (Smith et al., 2017). Insect behavior also affects susceptibility. If the caterpillars feed on pollen, tassels, husks, or leaves they

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n­ ormally ingest the insecticidal protein, but if they enter the ear via the silk channels they may avoid the toxin (Catangui and Berg, 2006; Eichenseer et al., 2008; Michel et al., 2010).

Winter Cutworm

Noctua pronuba (Linnaeus) (Lepidoptera: Noctuidae)

Natural History Distribution. Winter cutworm is also known as large yellow underwing, reflecting the appearance of the adult form instead of the larva. Yellow underwing is not a preferred name, because the hind wings are often orange, not yellow. It occurs widely around the world, including Europe, North Africa, the Middle East, and much of Asia. It has also attained North America, possibly in the 1970s, as it was documented first in Nova Scotia, Canada, in 1979. However, it soon spread in North America, and was detected in the New England states and New York, as well as Canada’s Atlantic Provinces, during the period of 1985– 1993, then North Carolina (1997), Pennsylvania (1998), and west to Colorado (1999), Wyoming (2000), California (2001), British Columbia (2002), and Alaska (2005). It is now found in all provinces of Canada, and much of the United States. It has long been known to be very dispersive in the adult stage, so it is not surprising that so much of North America quickly became home to this invasive insect. However, its spread may have been facilitated by human activity. Host Plants. Like many other noctuids, this species displays a wide host range, including such vegetables as beet, cole crops, carrot, lettuce, onion, potato, rhubarb, spinach, sugar beet, swiss chard, and tomato. In addition, it can damage fruit crops such as currant, grape, and strawberry, and ornamentals such as carnation, chrysanthemum, dahlia, freesia, gladiolus, hosta, iris, primrose, viola, and lawn grasses. Agronomic crops injured include alfalfa, canola, clover, forage grasses, and small grains such as oat, rye, and wheat. Among favored weed hosts are burdock, Arctium sp.; hawkweed, Hieracium sp.; dandelion, Taraxacum sp.; plantain, Plantago sp.; knotweed, Polygonum sp., and grasses, Poaceae. Natural Enemies. The natural enemies are not yet well known in North America, but tachinids are known to parasitize them in eastern North America and Trichogramma wasps have been reared from eggs in western North America. Life Cycle and Description. Overwintering occurs as partially grown to nearly fully grown larvae in protected places such as beneath plant residues. Apparently, larvae can enter diapause for part of the winter months, though many remain active at least during part of the winter. Larvae pupate in May or June, with moths observed during the

p­ eriod of June-October. Eggs can be produced soon after adult emergence or later, resulting in larvae of different ages entering the winter period. Some reports from Europe suggest two periods of egg laying and two complete generations, but this also could result from pupation at different times. In eastern North America, there seems to be an early period of flight in June-July and a later period of flight in August-September, with most of the moths in the latter period. Generally, it is believed that there is only one complete generation per year. Egg. Eggs are produced by females several weeks after fertilization. The eggs are deposited in clusters on leaves, stems, sticks, and elsewhere, apparently without regard to the availability of food. Females typically deposit three to five egg clusters of 180 to 522 eggs, totalling about 1400 eggs, but up to 2000 eggs can be produced by a single female. The eggs initially are cream-colored, but darken with age to brown, and eventually becoming dark gray or black before hatching. They are spherical and ribbed, and are arranged side-by-side in rows in a single layer, forming a sheet, on the underside of leaves. Eggs hatch in 2–6 weeks, depending on temperature. Madge (1962) reported that the egg incubation period was 43.1, 11.7, 7.5, 5, 5, and 2.5 days when reared at 10°C, 15°C, 20°C, 25°C, 27°C, and 28°C, though when reared at 28°C and above the survival rate was very low. He also noted that in England during early September almost all eggs hatched, however, as it became cooler (during October) egg hatch decreased dramatically, and during late October, when eggs were exposed to freezing temperatures, eggs failed to hatch.

FIG. 10.112  Larva of winter cutworm. (Photo by L. Buss.)

Larva. There are seven larval instars. The early developing larvae attain the mature larval state by autumn, but the later-developing larvae do not. It is the partly mature larvae that cause the most damage when they re-commence feeding in the spring (March to April).

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The larvae are quite variable in appearance. Most are either olive brown or green, with a line of black and cream ‘dashes’ subdorsally, on each side of the back, running the length of the body. The dashes tend to be black above and cream below. As with many cutworms bearing such markings, the dashes are heavier at the posterior end and faint toward the anterior end. The mean head capsule widths during instars 1–7 are 0.33, 0.50, 0.78, 1.15, 1.64, 2.45, and 3.65 mm, respectively. There are two dark markings on the head capsule. The prothoracic and anal shields are blackish brown. Mean body length (range) is 2.52 (1.92–2.95), 4.30 (3.46–4.93), 5.62 (5.25–6.80), 10.35 (9.02–11.57), 15.14 (14.90–16.50), 23.36 (20.80– 26.07), and 35.90 (31.30–42.00) mm during instars 1–7, respectively. The green larval phenotype varies from pale greenish to bright pea green. The mouthpart structures are used for diagnosis of this species (Neil and Specht, 1987). The optimum temperature for survival is about 15°C and 20°C. Larval activity patterns change over the course of development. During instar 1, larvae are diurnal and burrow into the soil at night. During instar 2, larvae are “intermediate,” with most larvae remaining on the foliage. During instars 3–7, however, larvae are repelled by light and thus are nocturnal. They emerge from the soil soon after dark and feed voraciously before reentering the soil as daybreak approaches (Madge, 1964). Pupa. The pupae are reddish brown or dark brown and about 2.5 cm in length. They resemble the pupae of many other noctuids. Pupation occurs on the soil surface beneath organic debris, or within the upper 1 or 2 cm of soil. There is no cocoon in this species.

nighttime temperatures, and negatively correlated with nighttime wind velocity. Males tend to fly low, with males outnumbering females by 10 to 1 at ground level. The adults have a wingspan of 50–60 mm. The orange or yellow hind wings, which bear a broad black border, are diagnostic. The forewings are more variable, normally blackish brown and mottled or with irregular dark spots in males, and orange to reddish with weak spots in females. Several phenotypes have been named. They are nocturnal, but if disturbed will take flight during the day. Summaries of N. pronuba biology in North America are provided by Difonzo and Russell (2010), Bechinski et  al. (2009), and Green et  al. (2016). All provide good photographs of the insects, and Green et al. (2016) also provide pictures of similar cutworms with which they might be confused. Madge (1962) provides important biological data from England.

Damage Generally, this species is considered to be a minor pest of vegetable gardens and flowers and not a commercial crop pest. There are exceptions, with damage to alfalfa and winter wheat crops observed in Idaho and Michigan. Damage by winter cutworm typically occurs during the autumn, winter, and early spring months, as this is when the larvae are large and plants usually are not growing rapidly. The larvae feed on foliage and stems, and like many other cutworms, can sever the stems of plants. The larvae may feed when temperatures are elevated above about 7°C. Thus, it has gained its name of winter cutworm based on its ability to forage under cool conditions, when most other cutworms are inactive. Larvae of this cold-tolerant species even have been observed crawling on snow. They also tend to be gregarious, which exacerbates their damage potential. Interestingly, incidental injury to pets can result from the consumption of numerous caterpillars by dogs and cats, which can cause vomiting and sickness. Poultry does not seem to be adversely affected (Difonzo and Russell, 2010).

Management

FIG. 10.113  Adult of winter cutworm. (Photo by L. Buss.)

Adult. Adults fly between June and October and deposit the eggs. Peak activity is, however, during August and September. Adult flight is positively correlated with mean

Noctua pronuba moths can be captured in traps baited with acetic acid plus 3-methyl-1-butanol, though this is a feeding attractant for many noctuid moths, so it is not selective (Landolt et al., 2015). Recommended practices for winter cutworm management include crop rotation, weed and crop residue management, tillage, and chemical control. Bechinski et al. (2009) provide a list of recommended insecticides. For additional information on cutworm management, see the section on variegated cutworm.

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Yellow-Striped Armyworm

Spodoptera ornithogalli (Guenée)

Western Yellow-Striped Armyworm Spodoptera praefica (Grote) (Lepidoptera: Noctuidae)

Natural History Distribution. Yellow-striped armyworm, Spodoptera ornithogalli (Guenée), is common in the eastern United States as far west as the Rocky Mountains and occurs in southern Canada. However, it is also reported from southwestern states, including California. The distribution of this native insect includes Mexico, Central and South America, and many Caribbean islands. As a pest, however, its occurrence is limited principally to the southeastern states. Western yellow-striped armyworm, Spodoptera praefica (Grote), is known only from the western states, principally California and Oregon. It is also a native insect. In California, S. praefica is much more important than S. ornithogalli. Host Plants. These species are general feeders, reportedly damaging many crops. Among vegetable crops injured are asparagus, bean, beet, cabbage, cantaloupe, carrot, corn, cucumber, lettuce, onion, pea, potato, rhubarb, rutabaga, salsify, sweet potato, tomato, turnip, and watermelon. Other crops damaged include alfalfa, blackberry, cotton, clover, grape, lentil, peach, rape, raspberry, sorghum, soybean, sugarbeet, sweetclover, sunflower, tobacco, wheat, and several flower crops. Some of the weed species known to be suitable hosts are castorbean, Ricinus communis; dock, Rumex sp.; gumweed, Grindelia sp.; horse nettle, Solanum carolinense; horseweed, Erigeron canadensis; jimsonweed, Datura sp.; lambsquarters, Chenopodium album; morning glory, Ipomoea sp.; plantain, Plantago lanceolata; prickly lettuce, Lactuca scariola; and redroot pigweed, Amaranthus retroflexus. In California, western yellow-striped armyworm develops for one to two generations on rangeland plants, preferring storksbill, Erodium sp.; and foxtail, Setaria sp. early in the year. However, as these plants senesce the armyworms move to irrigated areas, where they prefer alfalfa and morning glory. Alfalfa is a very suitable host, and such fields become heavily infested and serve as important sources of armyworms for other crops later in the season. A similar phenomenon was reported in Washington, where larvae developed on thistle, Cirsium spp.; wild lettuce, Lactuca spp.; wild mustard, Brassica spp.; and goosefoot, Chenopodium spp.; and then moved to crops as the weeds senesced (Halfhill, 1982). Natural Enemies. Several wasp parasitoids affect S. ornithogalli including Rogas laphygmae Viereck, R. terminalis (Cresson), Zele mellea (Cresson), Chelonus insularis

Cresson and Apanteles griffini Viereck (all Hymenoptera: Braconidae). Also, Euplectrus plathypenae Howard (Hymenoptera: Eulophidae) attacks larvae and causes a cessation of feeding within 2 days (Parkman and Shepard, 1981). Thus, this parasitoid is particularly valuable at minimizing damage. Numerous flies have been found to parasitize these armyworms including Archytas spp., Choeteprosopa hedemanni Brauer and Bergenstamm, Euphorocera omissa (Reinhard), E. tachinomoides Townsend, Lespesia aletiae (Riley), L. archippivora (Riley), Omotoma fumiferanae (Tothill), Winthemia quadripustulata (Fabricius), and W. rufopicta (Bigot) (all Diptera: Tachinidae). A NPV is highly pathogenic to larvae, and survivors that do not succumb exhibit reduced fecundity (Hostetter et  al., 1990; Young, 1990a, b). Undoubtedly, predators are important, but unlike the situation with western yellow-striped armyworm, their effect has not been quantified. Not surprisingly, western yellow-striped armyworm is also host to many parasitic insects. Among wasps associated with this caterpillar are Chelonus insularis Cresson, Cotesia marginiventris (Cresson) (both Hymenoptera: Braconidae), Hyposoter exiguae (Viereck), and Pristomeres spinator (Fabricius) (both Hymenoptera Ichneumonidae). Flies parasitizing western yellow-striped armyworm include Archytas californiae (Walker), Eucelatoria armigera (Coquillett), Euphorocera claripennis (Macquart), and Lespesia archippovora (Riley) (all Diptera: Tachinidae). Despite the numerous parasitoids, the first three species listed above, all wasps, account for 95% of recorded parasitism in alfalfa. The relative importance of the individual parasitoid species varies geographically (Miller, 1977). Of great interest, however, is the impact of predation. Bisabri-Ershadi and Ehler (1981) reported that over 96% of total mortality occurred in the egg and early larval stages, and most was attributed to predation. The most important predators were minute pirate bug, Orius tristicolor (White) (Hemiptera: Anthocoridae); big-eyed bugs, Geocoris spp. (Hemiptera: Lygaeidae); and damsel bugs, Nabis spp. (Hemiptera: Nabidae). The plant bug Lygus hesperus Knight (Hemiptera: Miridae) was a facultative predator, often feeding on armyworm eggs. A NPV is found in western yellow-striped armyworm, and when larvae occur at high densities it may be a significant mortality factor. Life Cycle and Description. There apparently are three to four generations annually, with broods of adults present in March-May, May-June, July-August, and AugustNovember. Some of the latter brood of yellow-striped armyworm and all members of the latter brood of western yellow-striped armyworm overwinter as pupae rather than emerging as adults. Although eggs, larvae, and adults of yellow-striped armyworm may be present in autumn or early winter they cannot withstand cold weather and perish. Development time, from egg to adult, is about 40 days.

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Egg. The eggs are greenish to pinkish brown and bear 45–58 small ridges. In shape, the egg is a slightly flattened sphere, measuring 0.46–0.52 mm in diameter and 0.38–0.40 mm in height. Females typically deposit clusters of 200–500 eggs, usually on the underside of leaves. Total fecundity was determined to be over 3000 eggs under laboratory conditions. They are covered with scales from the body of the adults. Duration of the egg stage is 3–5 days at warm temperature.

larval stage is 14–20 days, with the first three instars requiring about 2  days each and the last three instars requiring about 3 days each. Pupa. Larvae pupate in the soil within a cell containing a thin lining of silk. The reddish-brown pupa measures about 18 mm long. Duration of the pupal stage is 9–22 days, normally averaging 12–18 days.

FIG. 10.115  Yellow-striped armyworm moth. (Photo by L. Buss.) FIG. 10.114  Yellow-striped armyworm caterpillar. (Photo by L. Buss.)

Larva. The larvae initially are gregarious in behavior, but as they mature they disperse, sometimes spinning strands of silk upon which they are blown by the wind. There are usually six instars, though seven instars have been reported. Head capsule widths are about 0.28, 0.45, 0.8–1.0, 1.4–1.6, 2.0–2.2, and 2.8–3.0 mm, respectively, for instars 1–6. The larva grows from about 2.0 to 35 mm long over the course of its development. In California, mean development times for instars 1–7 of S. praefica were 2.5, 2.8, 3.0, 3.4, 4.5, 5.2, and 6.9  days, respectively, during the second generation. There was a total larval development time of 26.8 days during the second generation, but it was shortened to 21.3 days during the third. Coloration is variable, but mature larvae tend to bear a broad brownish band dorsally, with a faint white line at the center. More pronounced are black triangular markings along each side, with a distinct yellow or white line below. A dark line runs laterally through the area of the spiracles, and below this is a pink or orange band. Dark subdorsal spots are found on the mesothorax of y­ ellow-striped armyworm, and the triangular shape of these spots aids in distinguishing this insect from sweetpotato armyworm, Spodoptera dolichos, and velvet armyworm, S. latifascia, in eastern states. The head is brown but has extensive blackish markings. In western yellow-striped armyworm the blackish markings on the head form an arc on each side that extends from the mouth to the back of the head; in yellow-striped armyworm the band is less definite dorsally. Duration of the

Adult. The moths measure 34–41 mm in wingspan. The front wings are brownish gray with a complicated pattern of light and dark markings. Irregular whitish bands normally occur diagonally near the center of the wings, with additional white coloration distally near the margin. The hind wings of yellow-striped armyworm are opalescent white, with a narrow brown margin. In western yellowstriped armyworm the hind wings are similar but tend to be tinted with gray, and the underside bears a dark spot centrally. In S. praefica, both sexes are similar in appearance, whereas in S. ornithogalli the sexes are dimorphic. Under laboratory conditions average longevity of adults is 17 days, with most egg production completed by the tenth day (Adler et al., 1991). The most complete description of S. ornithogalli and its biology was given by Crumb (1929), with additional comments by Crumb (1956). Blanchard and Conger (1932) describes S. praefica. van den Bosch and Smith (1955) and Okumura (1962) described both species and their biology in California. Keys for identification are also found in these references. Keys for separation of Spodoptera adults can be found in Todd and Poole (1980) and Heppner (1998). Larvae can be distinguished using the keys of Passoa (1991), and Heppner (1998). Spodoptera ornithogalli can be distinguished from other common cutworms using Oliver and Chapin (1981), Stehr (1987), Sparks and Liu (2001), and a key to larvae included in Appendix A. Rearing on artificial diet was described by Adler and Adler (1988).

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Damage Larvae damage plants principally by consumption of foliage. The small, gregarious larvae tend to skeletonize foliage but as the larvae grow and disperse they consume irregular patches of foliage or entire leaves. However, they also feed on the fruits of tomato, cotton, and other plants. Larval consumption of soybean was estimated by King (1981) to total 115 cm2; this is an intermediate value relative to some other lepidopterous defoliators.

Management Insecticides. Insecticides are applied to foliage to prevent injury by larvae. The microbial insecticide Bacillus thuringiensis can be applied to kill armyworms but should be applied when the larvae are young, as they become difficult to control as they mature. Larvae consume bran bait containing insecticide. Cultural Methods. Proximity of crops to rangelandcontaining weed hosts, or to alfalfa, may be important factors predisposing vegetable crops to injury. At high densities, especially if alfalfa hay is mowed, larvae sometimes disperse simultaneously and invade nearby vegetable fields. Physical barriers such as trenches can be used to deter such dispersal. Biological Control. Liburd et  al. (2000) evaluated several biological insecticides on yellow-striped armyworm abundance and reported that although several Bacillus thuringiensis products and azadirachtin and a Steinernema carpocapsae product could reduce armyworm abundance significantly below untreated tomatoes, conventional chemical insecticides generally were more effective.

Zebra Caterpillar

Natural Enemies. A NPV of zebra caterpillar causes marked decreases in abundance, especially when larval densities are high (Adams et  al., 1968). Signs and symptoms of infection include loss of appetite, sluggish behavior, and reduced larval growth rate. Mortality occurs within 14 days of ingesting the polyhedral inclusion bodies, and dead larvae can be found hanging from foliage, suspended by their hind prolegs. Soon after death, the larval color darkens, and the cadavers turn black and rupture. Body contents, including virus polyhedra, contaminate the foliage, enhancing spread of the disease. There is some evidence that other armyworms and cutworms may be infected by M. picta NPV. Parasitoids known to attack zebra caterpillar include Limneria annulipes Harris (Hymenoptera: Ichneumonidae), Microplites mamestrae Weed (Hymenoptera: Braconidae), and Winthemia quadripustulata (Fabricius) (Diptera: Tachinidae) (Li et al., 1993). Life Cycle and Description. There are two generations per year over most of the range of this insect, generally occurring in June-July and August-October. A complete life cycle requires about 60 days during the summer. Zebra caterpillar overwinters in the pupal stage. Egg. The moths are highly fecund, often depositing 1200 white, ovoid, slightly flattened eggs in clusters of 100–200, usually on the underside of leaves. As might be expected from an insect with a northern distribution, eggs do not hatch at high temperature, 32°C and higher. Excellent survival occurs at about 27°C, and the hatching time is about 5 days. At 21°C, excellent hatching also occurs, but development is delayed to about 6 days. Hatching is inhibited at 15°C, and development time extended to about 12 days. At 10°C, eggs fail to develop.

Melanchra picta (Harris) (Lepidoptera: Noctuidae)

Natural History Distribution. This native insect is found in southern Canada and the northern United States from the Atlantic to the Pacific Coast. Nowhere it is considered to be a major pest, yet it can be a fairly regular nuisance, and a significant component of the defoliator pest complex of several crops. Host Plants. Zebra caterpillar feeds on several vegetable plants, and has been recorded as a pest of asparagus, bean, beet, broccoli, cabbage, celery, corn, lettuce, parsnip, pea, potato, rutabaga, spinach, tomato, and turnip; cabbage seems preferred. It also attacks flowers such as aster, hydrangea, and sweetpea, field crops such as alfalfa, clover, rape, sugarbeet, and tobacco, as well as berry crops such as raspberry, and trees such as apple, plum, and willow. As might be expected from an insect with such a broad host range, zebra caterpillar feeds on many weeds. Much of the economic entomology literature reports this insect as a pest of sugarbeet.

FIG. 10.116  Zebra caterpillar. (Photo by J. Capinera.)

Larva. The mature larva is boldly colored black, yellow, and white, and is immediately recognizable. The sides are yellow and white, with a vertical row of short-black stripes running the length of the body. There is a black band ­dorsally,

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separating the yellow and zebra-stripe lateral bands. The underside of the caterpillar is orange or red. Young larvae are very difficult to identify; they are principally black and green. However, they feed gregariously until they are about one-half grown, and this behavior helps to distinguish them. There are six instars. Head capsule widths are about 0.3, 0.5, 0.9, 1.5, 2.1, and 3.0 mm for the six instars, respectively. The average larval length is about 3.5, 6, 10, 17, 27, and 37 mm, respectively. Development time averages 3.5, 2.9, 3.3, 3.0, 3.4, and 7.2 days for larvae reared at 27°C. The last 3–4 days of the terminal instar is usually spent in the soil, where the larva prepares to transform into a pupa. Pupa. The pupa is dark brown and measures about 2 cm long. The males weigh about 400 mg and the females are slightly heavier, about 450 mg. Pupal development time is about 30 days. This is the normal overwintering stage.

before moving to new food. Foliage consumption and damage potential on sugarbeet were given by Capinera (1979a, b). Although individual zebra caterpillar larvae are voracious, they usually are not numerous enough to cause serious injury to crops.

Management These insects are not difficult to kill with foliar applications of chemical insecticides or Bacillus thuringiensis. They are also susceptible to the botanical insecticide neem, which functions as a feeding deterrent and growth regulator (Isman, 1993). Because of their seasonal biology, they sometimes develop to damaging levels late in the season, after the threat of insect damage is generally past. Thus, continued vigilance is suggested if the insects have been observed earlier in the season. Zebra caterpillar is susceptible to infection by Autographa californica NPV (Capinera and Kanost, 1979).

FAMILY PAPILIONIDAE— CELERYWORMS AND SWALLOWTAIL BUTTERFLIES Black Swallowtail

Papilio polyxenes Fabricius

Anise Swallowtail FIG. 10.117  Zebra caterpillar moth. (Photo by J. Capinera.)

Adult. The adult wingspan measures about 3.5–4.5 cm. The front wings are chocolate brown, with a weak gray spot centrally. The hind wind is white but bears a narrow brown band at the wing margin. Moths begin to emerge as early as 45 days after eggs are deposited, but usually, about 2  months is required for a complete generation to occur. The preoviposition period of moths is about 2 days. Moths may continue to deposit eggs for a period of up to 2 weeks, although most oviposition occurs within 1  week. Adults perish after 10–12  days. The biology of zebra caterpillar was given by Tamaki et  al. (1972) and Capinera (1979a). Keys to cutworm moths that include zebra caterpillar are provided by Rings (1977a, b), and Capinera and Schaefer (1983). Keys to cutworm larvae that include zebra caterpillar are provided by Capinera (1986) and Stehr (1987).

Damage The larvae are leaf-feeders, and initially, they feed gregariously. They may make small holes or skeletonize foliage early in their development, but they soon become voracious, eating large holes in foliage. Larvae often completely ­consume ­individual leaves, leaving only the stem or petiole

Papilio zelicaon Lucas (Lepidoptera: Papilionidae)

Natural History Distribution. Black swallowtail, also known as “American swallowtail,” is found in southern Canada and the eastern United States as far west as the Rocky Mountains. Its range also extends south into Mexico and northern South America. Anise swallowtail, also known as “western swallowtail,” inhabits southwestern Canada and the western United States from the Rocky Mountains to the Pacific coast. The ranges of these native species overlap in the Rocky Mountain region. Host Plants. Larvae of these species are commonly known as “parsleyworm” or “celeryworm” because they feed on plants in the family Umbelliferae. Vegetable crops fed upon by larvae include carrot, celery, fennel, parsley, and parsnip. The herbs anise, caraway, and dill also are eaten. Numerous umbelliferous weeds serve as suitable hosts, including angelica, Angelica spp.; cow parsnip, Heracleum spp.; lovage, Ligusticum spp.; water hemlock, Cicuta maculata; and wild carrot, Daucus carota. There are some chemical similarities between Umbelliferae and Rutaceae, the plant family containing citrus. Thus, occasionally these swallowtails are observed feeding on citrus, particularly orange trees, or other Rutaceae. This has

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o­ ccurred in California with greater frequency since fennel has been planted widely at lower elevations. Nowhere are these swallowtails considered to be a serious pest on citrus, however. Adults visit a variety of flowers to obtain nectar, they seem particularly fond of milkweed, Asclepias spp.; thistle, Cirsium spp.; and red clover, Trifolium pratense. Relative suitability of some plant hosts for black swallowtail larvae was provided by Finke and Scriber (1988). Natural Enemies. Avian predators are an important source of mortality for adults. Resting butterflies are especially susceptible to predation, and during inclement weather they spend more time roosting, thus incurring higher levels of predation (Lederhouse et al., 1987). Larvae are attacked by several predatory insects such as shield bugs (Hemiptera: Pentatomidae), damsel bugs (Hemiptera: Nabidae), and assassin bugs (Hemiptera: Reduviidae). A wasp parasitoid, Trogus pennator (Fabricius) (Hymenoptera: Ichneumonidae) attacks the larvae of both swallowtail species; black swallowtail, and likely anise swallowtail, are attacked by the fly parasitoids, Buguetia obscura (Coquillett), Compsilura concinnata (Meigen), and Lesesia frenchii (Williston) (all Diptera: Tachinidae). Life History and Description. These two species are very similar in life history, differing principally in the appearance of adults, as described below. There are two generations annually in northern regions of New York, but three generations in warmer climates. Overwintering occurs in the pupal stage, and in warm climates, some pupae from the second generation, as well as those from the third generation, enter diapause and emerge the following spring.

Larva. The color pattern among individuals and among the five instars is variable, but the general pattern follows. The first three instars are bird-dropping mimics, principally black with a median, dorsal spot resembling a saddle on the third and fourth abdominal segments. The third instar also bears some lateral orange and white markings. In the fourth instar, the caterpillar is principally black, but the anterior and posterior of each body segment is edged in green, and orange or yellow spots are located near the center of each body segment. The fifth instar is predominantly green, with black restricted to the center of each segment; this stage also has the orange spots found on the earlier instar. Larval development time is 10–30  days, depending on temperature. Larvae have an orange, eversible gland that resembles horns or antennae when extended. Located dorsally behind the head, the gland is exposed only when the caterpillar is disturbed. The gland, called an osmeterium, releases volatile chemicals that deter predation by some, but not all, insect predators (Berenbaum et al., 1992). Pupa. The pupa is attached with silk to a plant stem. The posterior end is attached closely to the plant, but the anterior end hangs away from the stem, at about a 30° angle, suspended by silk strands. The pupa is green or light brown, but also bears irregular black marks that help camouflage the pupa. The pupa measures 2.5–3.0 cm long. Except when the pupal stage is overwintering, its duration is 9–18 days.

Egg. The spherical eggs measure about 1 mm in diameter, and are pale green or cream initially, developing a reddish-brown cap with age. Eggs are deposited singly on leaves and flowers of host plants. Egg deposition begins about 2–4  days after adults emergence at 40–50 per day. The total number of eggs produced is estimated at about 200–400 per female, with oviposition occurring over about a 13-day period (Blau, 1981). Eggs hatch in 4–9 days.

FIG. 10.119  Adult butterfly of black swallowtail. (Photo by D. Hall.)

FIG. 10.118  Larva of black swallowtail butterfly. (Photo by J. Capinera.)

Adult. The adults of black swallowtail are principally black, but a row of yellow spots is found near the margin of the wings. In males, there is also a row of larger yellow spots parallel to the marginal spots but is located more centrally. In females, the interior row is greatly reduced. Both sexes bear, at the posterior margin of the hind wing, an elongation of the wing that forms a “tail,” a row of diffuse bluish spots, and a multicolored “eyespot.” The adults of anise swallowtail may greatly resemble black swallowtail, but more commonly they have the

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i­nterior row of yellow spots greatly expanded, filling the central region of the wings with yellow. The net result is that the western species, anise swallowtail, is predominantly yellow, whereas the eastern species, black swallowtail, is predominantly black. The adults of both species are quite large, the wingspan measuring over 7 cm. Adults are active during most of the day, and males are territorial, defending areas against other males. They often perch on elevated objects to maintain a good view of their territory, or they may patrol an open area, looking for females with which to mate. Males often frequent hilltops, and virgin females fly to hilltops to seek mates. The biology of these species was given by Chittenden (1912d), Fisher (1980), Scott (1986), and Opler and Krizek (1984). Culture techniques were described by Carter and Feeny (1985).

Damage Larvae are large and can consume considerable quantities of foliage toward the end of their development. This should be of concern only in the home garden because the butterflies are not sufficiently abundant ever to threaten commercial cultivation of an umbelliferous crop.

Management Owing to their infrequent occurrence, control of larvae should not be necessary. For at least the past 100  years, handpicking has been recommended in the home garden, and this recommendation remains valid. In instances where plot size is too big to make this practical, larvae can be killed easily with foliar insecticides, either chemical or Bacillus thuringiensis.

FAMILY PIERIDAE—CABBAGE WORMS, WHITE, AND SULFUR BUTTERFLIES Alfalfa Caterpillar

Colias eurytheme Boisduval (Lepidoptera: Pieridae)

Natural History Distribution. This native species can be found throughout the United States, and often disperses to southern Canada during the summer. Formerly limited to central and western North America east of the Appalachian Mountains, the eastern dispersal of alfalfa caterpillar in the early 1900s was helped by the clearing of forests and widespread culture of alfalfa. It remains most common and damaging, in the southwest. Host Plants. Alfalfa caterpillar feeds on several forage legumes, particularly alfalfa, sweet clover, white clover, and hairy vetch. Red clover is not suitable for development,

and adults rarely oviposit on this plant. Wild hosts include many species of milkvetch, Astragulus spp.; trefoil, Lotus spp.; clover, Trifolium; and vetch, Vicia spp. Among vegetable crops sometimes consumed are bean and pea, but not cowpea. The adults feed on a succession of nectar-producing flowers, though the sequence varies between localities. In northern Virginia, the preferred flowers are dandelion, Taraxacum spp., and winter cress, Barbarea spp., during the first generation; followed by dogbane, Apocynum sp., and clover, Trifolium spp., during the second generation; then by milkweeds, Asclepias spp., in the third generation; and goldenrods, Solidago spp., aster, Aster spp., and tickseed sunflower, Bidens coronata, during the fourth to fifth generations (Opler and Krizek, 1984). Natural Enemies. Natural enemies are often effective to keep this insect from becoming very abundant, and consequently, it generally is not a serious pest. An egg parasitoid, Trichogramma sp., occasionally destroys 50% of the eggs. As it develops rapidly, completes two generations in the very time it takes the host to complete a single generation, this egg parasitoid builds to high levels by the end of the season when caterpillar populations are high (Wildermuth, 1914). Young caterpillar larvae are attacked frequently by Apanteles flaviconchae Riley (Hymenoptera: Braconidae), with over 50% of them destroyed during some years. Michelbacher and Smith (1943) reported that when this parasitoid was abundant during the first generation of its host, the alfalfa caterpillar did not reach damaging levels. Other braconid parasitoids of lesser importance include Apanteles cassianus Riley, A. medicaginis Muesebeck, Meteorus autographae Muesebeck, M. laphygmae Viereck, and M. leviventris (Wesmael). Other wasps known from alfalfa caterpillar are Itoplectis vidulata (Gravenhorst), Thyrateles instabilis (Cresson), Nepiera benevola Gahan, Hyposoter exiguae (Vierick), and Pristomerus spinator (Fabricius) (all Hymenoptera: Ichneumonidae). A pupal parasite, Pteromalus puparum (Linnaeus) (Hymenoptera: Pteromalidae), can be very important, sometimes parasitizing up to 60% of the pupae (Wildermuth, 1920). Among other parasitoids known from alfalfa caterpillar are Euphorocera claripennis (Macquart), E. omissa (Reinhard), and Lespesia archippivora (Riley) (all Diptera: Tachinidae). The importance of predators has not been determined, but several are known. Ants (Hymenoptera: Formicidae) and lady beetles (Coleoptera: Coccinellidae) kill the caterpillar larvae, and robber flies (Diptera: Asilidae) capture adults. Pupae of alfalfa caterpillar are sedentary, and therefore easy prey, and are consumed by the softwinged flower beetle, Collops vittatus Say (Coleoptera: Melyridae), larvae of corn earworm, Helicoverpa zea Boddie (Lepidoptera: Noctuidae), and likely many others. A disease caused by a cytoplasmic polyhedrosis virus sometimes causes spectacular alfalfa caterpillar population

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collapse late in the season. However, as happens with many insect diseases, these become apparent only during seasons when other natural enemies are ineffective and caterpillar densities are high. Life Cycle and Description. The adults are observed from March to May until late November over most of the ranges of this species. The number of generations varies with latitude and elevation. In northern regions and in the Rocky Mountains, only two generations are reported annually. However, three to five generations occur over most of the United States, five to six are reported from southern California and Arizona, and seven generations occur in Louisiana. During mid-summer a complete life cycle can be completed in 30 days. In northern regions this species survives periods of cold weather in the pupal stage, though survival in northern latitudes is poor. In southern regions, alfalfa caterpillar tends to pass the cold periods in the larval stage. Egg. The egg of alfalfa caterpillar measures about 1.2–1.5 mm long and 0.4–0.6 mm wide. The egg is spindleshaped, tapering to a point at each end, and marked with 18–20 raised longitudinal ridges which are connected with smaller cross-ridges. They are deposited on end to the upper surface of foliage, and normally singly. They are greenish white initially, but turn reddish after the second day. Duration of the egg stage is 3–7 days, with mean development time of 2.5–3.0 days at 26–32°C. Mean fecundity in the field is about 350 eggs during the adult lifespan of about 2 weeks. However, under laboratory conditions and exceptionally in the field, the female can live for over a month and deposit 700 eggs.

FIG. 10.120  Alfalfa caterpillar larva. (Photo by J. Capinera.)

Larva. There are five instars. Body length is reported to be 1.5–3.0, 3.0–5.5, 4.5–8.0, 8.0–15, and 15–29.0 mm in instars 1–5, respectively (Michelbacher and Smith, 1943). Larvae are primarily uniform green. A white lateral line normally runs the length of the body; a thin yellow, pink, or red line often occurs within the white line. Dorsally, a dark dorsal line and pair of fine white subdorsal lines may also be found. In some specimens the lines are absent, and the

body is completely green. Mean larval development time during instars 1–5 is 2.6, 2.2, 1.8, 2.3, and 4.3 days when cultured at 27°C, and 2.1, 1.8, 1.7, 2.1, and 2.9 days when cultured at 32°C, respectively. Pupa. The pupa is angular, bearing keel-like projections on the thorax and tapering to a point at both the anterior and posterior ends. It is light green or pink, measures about 17.5 mm long, and bears a yellow line and three black dots on each side of the abdomen. The pupa is anchored loosely to a plant stem, anterior end pointed upward, by means of a thread and a posterior anchor. Duration of the pupal stage is often 7–10 days, but averages 4–5 days at 27–32°C.

FIG. 10.121  Alfalfa caterpillar moth. (Photo by J. Capinera.)

Adult. The adults usually are orange-yellow butterflies with a broad black band distally on the upper surface of each wing. A large black spot is also found near the midpoint of each forewing. In females, but not males, the black marginal band contains light spots. The undersurface of each wing, in both sexes, is uniformly pale yellow. The wingspan measures about 50–58 mm in males, and about 62–64 mm in females. The upper surface of the male’s wings reflects ultraviolet light. This light reflection is visible to butterflies and serves to separate alfalfa butterfly from a co-occurring species, Colias philodice Godart, and to minimize hybridization (Silberglied and Taylor Jr., 1978), though pheromones are also involved in mating (Sappington and Taylor, 1990). The adults of alfalfa butterfly are quite variable in appearance, and white variants are sometimes observed. Also, larvae exposed to cold temperature produce, after a lag of one generation, adults with wings that are mostly yellow, but containing some orange. Larvae exposed to warm temperature, in contrast, produce adults with orange wings (Tuskes and Atkins, 1973). Common sulfur butterfly, C. philodice, is most easily confused with alfalfa butterfly, but it is yellow rather than yellow orange. Hybrids between these two species are not uncommon.

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The biology of alfalfa caterpillar was described by Wildermuth (1914, 1920), though Floyd (1940) and Michelbacher and Smith (1943) added important observations. A useful synopsis was included in Opler and Krizek (1984). Methods of rearing were given by Taylor Jr. et al. (1981).

Damage Alfalfa caterpillar larvae are defoliators. Young larvae initially feed on the leaf surface or make small holes in leaves, but soon consume large quantities of leaf material. Although they are recorded from several types of bean and pea, damage to vegetable crops is usually minor. The pupal stage is an important contaminant of peas processed for freezing and canning.

Management Vegetables generally are not at risk unless the caterpillars are very abundant on other crops or weeds. Thus, it is useful to monitor nearby alfalfa crops for the presence of caterpillars. The larvae and pupae are difficult to detect visually on foliage, and a sweep net is often used for sampling. If larvae are abundant in alfalfa, especially early in the season, it is often beneficial to harvest the alfalfa early because such harvesting can result in caterpillar mortality, and subsequent crops may avoid infestation. Infested crops can also be treated with insecticide or Bacillus thuringiensis applied to the foliage.

Imported Cabbage Worm

Pieris rapae (Linnaeus) (Lepidoptera: Pieridae)

Natural History Distribution. Imported cabbage worm is found principally in temperate regions of Europe, Africa, Asia, and North America, but is now firmly established in Australia and New Zealand, southern Mexico, and Hawaii. Imported cabbage worm was first observed in North America in 1860 in Quebec City, Canada. It spread to Massachusetts and New Jersey by 1869, reached Ohio in 1875, and by 1886 it was found in the Gulf Coast and Rocky Mountain states. P. rapae attained British Columbia in 1898 and the Pacific Coast by 1901. It is now widespread in North America. It is a strong flier and is also spread long distances annually by strong winds. Few cabbage worms reportedly survive the winter in most of Canada, but much of the country is invaded annually by dispersants from the United States or from southern Canada (Beirne, 1971). Pieris rapae is easily confused with other common cabbage white butterflies: Pontia protodice, southern cabbage worm; Pieris napi, mustard white; band Ascia monuste (Linnaeus), southern white. Before the introduction of

i­mported cabbage worm, P. napi (Linnaeus) was the dominant cabbage butterfly in the north, and Pontia protodice (Boisduval and LeConte) was the principal cabbage-­feeding butterfly in the south. Both have been largely replaced by P. rapae, though they sometimes co-occur on cultivated crucifers or on weeds. A key for the differentiation of these species is included in Appendix A. Some taxonomists prefer to place this species in the genus Artogeia (Robbins and Henson, 1986). Host Plants. Larvae of this insect feed widely on plants in the family Cruciferae, but occasionally on a few other plant families that contain glucosinolate and related chemical compounds (mustard oils). Vegetable crops attacked include broccoli, Brussels sprouts, cabbage, cauliflower, collards, horseradish, kale, kohlrabi, mustard, radish, turnip, and watercress. Also attacked are flowers such as nasturtium and sweet alyssum, and weeds such as field pennycress, Thlaspi arvense; pepperweed, Lepidium spp.; wild mustard, Brassica kaber; horsehair mustard, Conringia orientalis; and yellow rocket, Barbarea vulgaris. Adults sip nectar from flowers, and are commonly seen at mustards, Brassica spp.; dandelion, Taraxacum officinale; aster, Aster spp.; purple heliotrope, Heliotropum spp.; thistle, Cirsium spp.; and both weedy and cultivated crucifers (Harcourt, 1963a). Sea kale is reported to be attractive for oviposition, but larvae fail to complete their development on this plant (Richards, 1940). Natural Enemies. Imported cabbage worm is subject to numerous predators, parasitoids, and diseases. General predators such as shield bugs (Hemiptera: Pentatomidae), ambush bugs (Hemiptera: Phymatidae), and vespid wasps (Hymenoptera: Vespidae) attack them, as do many insectivorous birds. Chittenden (1916a), for example, noted 90% predation of overwintering pupae by birds. Similarly, Schmaedick and Shelton (1999) documented high, but variable, levels of predation of eggs and young larvae in New York, averaging 53% but sometimes attaining up to 80% mortality. However, parasitoids are considered to be much more important mortality factors. Harcourt (1963a) identified three important species in Ontario. Cotesia glomeratus (L.) (Hymenoptera: Braconidae) attacks the early instars and emerges from the mature larva as it prepares to pupate. Phryxe vulgare (Fallon) (Diptera: Tachinidae) attacks mature larvae and emerges from the host pupa. Pteromalus puparum (L.) (Hymenoptera: Pteromalidae) attacks and kills cabbageworm pupae. Cotesia glomeratus has long been considered to be the most important parasitoid in Canada and in the northern United States. Cotesia glomeratus is readily observed in the field, searching diligently on foliage for larvae. Dead cabbageworm larvae are often found with clusters of 20–30, C. glomeratus cocoons attached. However, in Europe, Cotesia rubecula (Marsall) (Hymenoptera: Braconidae) is the most important parasitoid, and in recent years it has become established in North America where it is now assuming a dominant role (Godin and Boivin, 1998b, c).

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In contrast, tachinids were more important in California, particularly Madremyia saundersii (Williston) (Diptera: Tachinidae) (Oatman, 1966b). As observed in Canada, however, C. glomeratus was also a significant larval mortality factor in California. A low level of egg parasitism by Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae) occurred, but the failure of eggs to hatch, which was attributed to infertility caused by cool weather, resulted in more deaths than did egg parasitoids. The pupal parasitoid P. puparum was also effective in California, but this is not always the case. In some locations, relatively low levels of parasitism have been observed (Oatman and Platner, 1969; Ru and Workman, 1979; Lasota and Kok, 1986); high rates tend to occur late in the season. Virus and fungal diseases of imported cabbage worm have been reported, but the predominant natural disease in a granulosis virus (GV). Pieris rapae GV occurs most commonly under high-density conditions, and often among late instar larvae after they have consumed the exterior foliage of plants and are forced into close contact. Harcourt (1963a) observed over 90% mortality of larvae due to natural occurrence of this disease. In the early stages of infection, larvae are inactive and paler in color. As the disease progresses, the caterpillar body turns yellow, and tends to appear bloated. After death, the body blackens, the integument ruptures, and the liquefied body contents ooze on the plant foliage. Rainfall has a major roll in assisting the spread of the virus on the plant, and from the soil to the plant. Beirne (1971) suggested that outbreak of granulosis virus in the second generation of cabbage worm often prevented the third annual generation from causing extensive damage. Life Cycle and Description. The complete life cycle of this insect requires 3–6 weeks, depending on the weather. Godin and Boivin (1998a) reported that about 320 degreedays above a threshold of 10°C was required to complete a generation in Quebec. The number of generations reported annually is two-to-three in Canada, three in the New England states, three-to-five in California, and six-to-eight in the south. Imported cabbage worm can be found throughout the year in the south. Egg. The eggs are laid singly, usually on the outer leaves of plants. About 70%–85% of the eggs are deposited on the lower surface of the leaves, where the larvae also tend to feed. The egg measures 0.5 mm wide and 1.0 mm long, and initially it is pale white, but eventually turns yellowish. The egg is laid on end, with the point of attachment flattened and the distal end tapering to a blunt point. The shape is sometimes described to resemble a bullet. The egg is strongly ribbed, with 12 longitudinal ridges, and hatches in about 5 days (range 2–8 days) during August.

FIG. 10.122  Imported cabbageworm larva. (Photo by P. Choate.)

Larva. The larva is green, velvety in appearance, and bears five pairs of prolegs. There are five instars. Head capsule widths are about 0.4, 0.6, 0.97, 1.5, and 2.2 mm, respectively. Body lengths at maturity of each instar averages 3.2, 8.8, 14.0, 20.2, and 30.1 mm, respectively. The larva requires about 15 days (range 11–33 days) to complete its development during August. Average (and range) development time for each instar at 19°C was observed to be 4.5 (2.5–6), 3.0 (1.5–5), 3.3 (2–5), 4.1 (3– 6.5), and 7.8 (5–18) days, respectively. All larval stages except the first instar bear a narrow yellow line running along the center of the back; this stripe is sometimes incomplete on the early instars. A broken yellow line, or series of yellow spots, also occurs on each side. The mature lava typically wanders before pupation, spending an average of 1.8  days (range 1–4  days) without feeding, and finally spins a silk pad or platform, where pupation occurs. Pupa. Pupation often occurs on the food plant, but cabbage worm may leave the plant to pupate in nearby debris, especially when larval densities are high. The chrysalis is about 18–20 mm long, and varies in color, usually matching the background; yellow, gray, green, and speckled brown are common. A sharply angled, keel-like projection is evident dorsally on the thorax, and dorsolaterally on each side of the abdomen. At pupation, the chrysalis is anchored by the tip of the abdomen to the silk pad, and a strand of silk is loosely spun around the thorax. The silk line serves to anchor the anterior portion of the chrysalis, and also keeps the head from hanging down. Pupation during the summer generations lasts, on average, 11  days (range 8–20  days). The chrysalis is the overwintering stage, however, so its duration may be prolonged for months if the pupa diapauses. The proportion of pupae that diapause increases as autumn progresses so that at the time of the final generation all pupae are in diapause.

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Imported cabbageworm larvae are present, and potentially damaging, throughout most of the period that crucifer crops are under culture. In northern latitudes such as Wisconsin (Radcliffe and Chapman, 1966) and Ontario (Harcourt, 1963a) damage typically occurs during summer months, in southern California they are most abundant during the autumn (Oatman and Platner, 1969), and in southern latitudes such as South Carolina (Reid Jr. and Bare, 1952) and Louisiana (Smith and Brubaker, 1938), damage occurs during the autumn, early winter- and spring-month periods that coincide with peak crucifer production.

FIG. 10.123  Imported cabbageworm moth. (Photo by D. Hall.)

Adult. Upon emergence from the chrysalis, the butterfly has a wingspan of about 4.5–6.5 cm. It is white above with black at the tips of the forewings. The front wings are also marked with black dots—two in the central area of each forewing in the female, and one in case of males. When viewed from below, the wings generally are yellowish, and the black spots usually show through the wings. The hind wing of each sex also bears a black spot on the anterior edge. The black spots and the yellow coloration may be reduced or absent from both sexes, especially in the spring generation. The body of the butterfly is covered with dense hair, which is colored white in females, but darker in males. The adult typically lives about 3  weeks, and may be active very early in the spring. The female produces 300–400 eggs. The adult is very active during the daylight hours, often moving from the crop to flowering weeds to feed. This movement tends to result in a preponderance of the eggs being deposited on the edges of crucifer fields unless there are flowering weeds contained within the crop. The biology of imported cabbage worm was given by Chittenden (1916a), Wilson et al. (1919), Richards (1940), and Harcourt (1962, 1963a). Keys that include imported cabbageworm larvae include Capinera (1986) and Sparks and Liu (2001). The culture was described by Webb and Shelton (1988).

Damage The larvae defoliate crucifer crops, sometimes killing young plants. Severe damage to young plants often prevents head formation even when the caterpillars are later removed, so early season protection is important (Wilson et al., 1919). If left unchecked, cabbage worm often can reduce mature plants to stems and large veins. Although they prefer leafy foliage, larvae may burrow into the heads of broccoli and cabbage, especially as they mature. Larvae are often immobile and difficult to dislodge, and they may be overlooked when cleaning produce. Larvae produce copious quantities of fecal material which also contaminate and stain produce.

Management Sampling. Harcourt (1962) studied the distribution of imported cabbage worm on crops. He suggested that onehalf of each plant be examined visually for various stages. Recommended sample sizes were 20 plants for eggs, 30 for young larvae, 40 for mid-age larvae, 50 for large larvae, and 70 for pupae. Larvae often rest along the principal leaf vein and are very difficult to see because their body color closely matches the background. Damage and fecal material is often the most visible indication of infestation. The presence of highly visible butterflies suggests future problems. Shelton et  al. (1994) compared the benefits of sequential and variable-intensity sampling for cabbageworm management and recommended the latter as being more reliable and requiring fewer samples. Insecticides. Imported cabbage worm is readily killed by foliar application of insecticides, including the bacterial insecticide Bacillus thuringiensis and some botanical insecticides (Hamilton and Gemmell, 1934; Huckett, 1934, 1946; Dills and Odland, 1948). Biological Control. Several microbes have been investigated for control of imported cabbage worm, and have the potential to be developed as microbial insecticides. The imported cabbageworm granulosis virus (Pieris rapae GV) suppressed cabbageworm larvae in the laboratory (Payne et al., 1981) and in a field test, but required 4–10 days to inflict mortality, and was not superior to control provided by Bacillus thuringiensis (Jaques, 1973). The NPV from alfalfa looper (Autographa californica NPV), a granulosis virus from cabbage butterfly (Pieris brassicae GV), and the microsporidian Vairimorpha necatrix were shown by Jaques (1977) and Tompkins et al. (1986) to suppress imported cabbage worm, but the population reduction was not superior to that achieved using Bacillus thuringiensis. These pathogens have commercial potential because they are not very host-specific, and also suppress another important crucifer pest, cabbage looper, Trichoplusia ni (Hübner). Home gardeners sometimes collect dead or dying caterpillars, macerate them in water, and spray the resulting suspension onto foliage as a homemade biological insecticide.

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Flower flies (Diptera: Syrphidae) consume the eggs and small larvae of imported cabbage worm and numerous other insects. Populations of flower flies have been manipulated to increase predation of cabbage worm by interplanting cabbage with pollen-rich flowering plants. Although aphid populations on cabbage were suppressed, imported cabbageworm populations were not decreased significantly (White et al., 1995). Host-Plant Resistance. Crucifer crops differ in their susceptibility to attack by imported cabbage worm. Chinese cabbage, turnip, mustard, rutabaga, and kale are less preferred than cabbage, collards, Brussels sprouts, broccoli, and cauliflower (Harrison and Brubaker, 1943; Radcliffe and Chapman, 1966). Some cultivars of certain crops also have moderate levels of resistance to infestation by imported cabbage worm. One resistance character is due to, or correlated with, dark green, glossy leaves. This character imparts resistance to imported cabbage worm and other caterpillars but increases susceptibility to flea beetle injury (Dickson and Eckenrode, 1980). The red color found in many crucifer varieties also affects imported cabbage worm. Cabbage butterflies avoid ovipositing on red cabbage varieties (Radcliffe and Chapman, 1966). However, larval survival is favored by red cabbage. Thus, while important genetic material has been identified, in most cases existing varieties are not a practical solution to caterpillar problems. Research conducted in Virginia on the susceptibility of broccoli cultivars, and where pest abundance was relatively low, indicated that the principal factor in susceptibility to attack was the date of plant maturity; early maturing varieties were less infested (Vail et al., 1991). Cultural Practices. Herbs are sometimes recommended as companionate or repellent plants for vegetable cultivation. Herbs are hypothesized to give off odors that repel ovipositing vegetable pests, or prevent them from locating the vegetables. In most cases, this has not been investigated critically. However, for imported cabbage worm, there is ample evidence that herbs impart no benefit, and some herbs are associated with increased cabbageworm infestation (Latheef and Ortiz, 1983a, b). Paper caps early in the season, and row covers later, are effective in preventing oviposition by imported cabbageworm butterflies.

Mustard White

Pieris napi (Linnaeus) (Lepidoptera: Pieridae)

Natural History Distribution. This insect is found both in North America and Eurasia, where it has a decidedly northern

range. In the eastern United States, it occurs principally in the northernmost states, but in the West, its range extends as far south as New Mexico and California. It is widespread in Canada and Alaska, extending north to the edge of the Arctic tundra. Mustard white has declined in abundance in the northeastern United States, a phenomenon commonly attributed to competition with imported cabbage worm. However, adults frequent shaded areas, whereas imported cabbageworm butterflies prefer sunny fields, so competition may not completely explain the decline in mustard white abundance. It has also been suggested that change in habitat availability could account for this population decline. Host Plants. Larvae feed on wild and cultivated Cruciferae. Weeds such as Virginia pepperweed, Lepidium virginicum; toothworts, Dentaria spp.; and the Brassica and Descurainia mustards are suitable larval hosts. Females oviposit on yellow rocket, Barbarea vulgaris, and field pennycress, Thlaspi arvense, but larvae cannot complete their development on these plants. Vegetables attacked include Brussels sprouts, cabbage, radish, turnip, watercress, and probably others. Adults take nectar from flowers, preferring Geranium spp. if they are available. Life Cycle and Description. One to three generations are reported; two is most common. In areas with three generations, many members of the second generation enter diapause. Egg. The eggs are pale yellow or white, tinged with green, and bear 13 ridges. The eggs are elongate, tapering to a blunt point distally. They measure about 1.2 mm long and 0.45 mm wide. They are deposited singly, on end, to the underside of leaves and on stems of hosts. The female apparently produces about 300 eggs over the course of her life, which may be of several weeks’ duration. Larva. Larvae are covered with short, fine hairs, imparting a velvety appearance. They are green with small black spots, with a green stripe dorsally and yellow spots along each side. Mustard white larvae are easily confused with other pierid butterfly larvae; a key to separate these species is provided in Appendix A. Pupa. The chrysalis is green, gray, or tan flecked with black. It bears frontal and lateral projections similar to imported cabbage worm. Light-colored dorsal and lateral stripes may occur on the chrysalis. The mustard white undergoes diapause in the pupal stage.

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FIG. 10.124  Adult of mustard white. (Drawing by USDA.)

Adult. The wingspan is about 4–5 cm. This insect is white dorsally, and sometimes bears one or two black spots on the front wings. The lower side of the hind wing is usually yellow, as is the tip of the forewing. The most distinctive feature is the darkening along the wing veins; this is especially evident on the hind wings, and most pronounced on the underside of the hind wings. This insect has two color forms. The summer generation tends toward nearly white, while the spring generation has pronounced bands of gray along the veins. A key for separation of this species from other common cabbage white butterflies is included in Appendix A. The biology of mustard white was given by Opler and Krizek (1984), and Scott (1986).

Host Plants. Southern cabbage worm attacks a variety of cultivated crucifers but is much less abundant than imported cabbage worm, Pieris rapae (Linnaeus). Apparently southern cabbage worm, which formerly was quite abundant in some localities, has been largely displaced by Pieris rapae. Vegetable crops known to be attacked are cabbage, cauliflower, Chinese cabbage, radish, and turnip. The host range is probably equivalent to imported cabbage worm, but because southern cabbage worm is not a serious pest it has not been well studied. Wild crucifers that support the growth of southern cabbage worm include pepperweed, Lepidium spp.; yellow rocket, Barbarea vulgaris; shepherdspurse, Capsella bursa-pastoris; tansymustard, Descurainia pinnata; field pennycress, Thlaspi arvense; hoary cress, Cardaria draba; black mustard, Brassica nigra; and others. Life Cycle and Description. Three or four generations occur annually. Southern cabbage worm overwinters in the pupal stage. Egg. The eggs are pale yellow initially, but turn orange as they mature. They are barrel-shaped, but taper at the apex, and are bear deep ridges. Eggs are deposited singly on flowers or leaves of the larval host plants.

Damage Larvae feed on the foliage and produce small to large holes depending on larval size. They are rarely numerous enough to warrant concern.

Management Mustard white is rarely a serious pest and should respond to management practices developed for imported cabbage worm.

Southern Cabbage Worm

Pontia protodice (Boisduval and LeConte) (Lepidoptera: Pieridae)

Natural History Distribution. Southern cabbage worm is found throughout the United States and south into Mexico. It is native to North America. As its common name suggests, it is principally a southern insect and occurs in the North only sporadically. It is rare in the New England states and southern Canada. The adult is known to lepidopterists as the checkered white, and the species name is sometimes given as Pieris protodice.

FIG. 10.125  Larva of southern cabbage worm. (Photo by J. Capinera.)

Larva. Larvae are pale blue green or bluish gray, with numerous small black spots. They bear a pair of yellow stripes running the length of the back, and a stripe on each side. The head is yellow with reddish spots. Mature larvae attain a length of about 3 cm. larvae prefer to feed on flowers or fruits, but also eat foliage. A key to distinguish this caterpillar from similar pierid larvae feeding on crucifers is provided in Appendix A.

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Pupa. The chrysalis is light blue grey and speckled with black. White strips are sometimes found laterally. In general shape, the chrysalis resembles that of imported cabbage worm, bearing sharp keel-like structures on the back. This is the overwintering form.

d­ iapause, may be smaller and very light in color. Adults can be seen feeding on nectar from a wide variety of flowering plants, and are highly dispersive. When they mate, the male transfers a spermatophore to the female that represents 7%–8% its body weight. The female absorbs this, and it assists with her nutritional needs during ovposition. A key for differentiating this species from other cabbage white butterflies is included in Appendix A. The biology of southern cabbage worm was given by Opler and Krizek (1984), and Scott (1986).

Damage Larvae prefer to feed on flowers and flower buds, but they also eat leaves. On cabbage, they restrict their feeding to the outer leaves. This makes them much less damaging than imported cabbage worm, which bore into the head. FIG. 10.126  Southern cabbage worm, adult male. (Photo by D. Hall.)

Management Southern cabbageworm butterflies were observed to be highly attracted to fluorescent orange and to be captured with sticky traps, especially when the traps were positioned close to the soil surface (Capinera, 1980). The methods discussed for management of imported cabbage worm are also appropriate for southern cabbage worm.

Southern White

Ascia monuste (Linnaeus) (Lepidoptera: Pieridae)

Natural History

FIG. 10.127  Southern cabbage worm, adult female. (Photo by D. Hall.)

Adult. The adults have a wingspan of about 3.5–5 cm. They are similar in appearance to imported cabbage worm, but the southern cabbageworm butterflies are more heavily marked with scattered grayish brown or black spots, both dorsally and ventrally, on their white background. The female, in particular, may be dark, with up to 50% of the wings darkened. The males, in contrast, bear only a few dark spots, but are still more heavily marked than Pieris rapae. The first generation butterflies, which emerge in the spring following

Distribution. Southern white resides in Florida, the Gulf Coast areas of the southern states, southern Texas, Mexico, and most of Latin America. This butterfly exhibits strong migratory tendencies, moving northward in the summer and southward in the winter, but in North America, it rarely becomes numerous anywhere other than subtropical, coastal areas. They are more dispersive under high-density conditions. There are at least seven named subspecies of this insect. The subspecies found in North America is A. monuste monuste (L.), whereas in South America it is A. monuste orseis Godart. This is noteworthy because unlike most subspecies, the biology differs between these races (Liu, 2005). Host Plants. Larvae feed on plants in the families Cruciferae, Bataceae, Capparidaceae, and Tropaeolaceae. Principal hosts are saltwort, Batis maritima, in coastal

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r­egions and pepperweed, Lepidium virginicum, elsewhere. Vegetables fed upon are cabbage, cauliflower, collards, kale, radish, and turnip. Other food plants include beach cabbage, Cakile maritima; spider flower, Cleome spinosa; clammy weed, Polanisia sp. nasturtium, Tropaeolum majus; and others. Adults frequent a variety of flowering plants to obtain nectar. Life Cycle and Description. The number of generations is poorly documented, but at least three are known annually, and adults are active year-round in southern Florida and Texas. Reproductive diapause likely occurs during the winter months. Egg. The eggs are pale yellow, elongate, and bear 11 longitudinal ridges. Eggs average (± SE) about 0.15 ± 0.03 mm in length and 0.58 ± 0.02 mm in diameter. They may be laid singly, in small groups, or in clusters up to about 50, depending on the larval host. On average, eggs are laid in clusters of 16 on young leaves. The female may deposit 800–1000 eggs over the course of her life, which is typically 8–10 days in duration. Mean duration of the egg stage (± SE) at 25°C 4.7 ± 0.1 days.

At 25°C, development time (± SE) averaged 2.8 ± 0.1, 2.3 ± 0.1, 3.1 ± 0.1, 3.2 ± 0.2, and 6.0 ± 0.2 days for instars 1–5, respectively. Thus, the total larval period is about 17.4 ± 0.3  days. The mean body lengths (± SE) are 0.32 ± 0.02, 0.74 ± 0.03, 1.71 ± 0.04, 2.25 ± 0.26, and 4.43 ± 0.08 cm for instars 1–5, respectively. Mean head capsule widths (± SE) for instars 1–5 are 0.040 ± 0.002, 0.083 ± 0.003, 0.135 ± 0.002, 0.231 ± 0.007, and 0.344 ± 0.006, respectively (Liu, 2005). Pupa. The chrysalis is ivory white with numerous black markings and bears one orange-yellow band dorsally, and another on each side. The general shape is similar to that of imported cabbage butterfly, but the projections are greatly blunted in the southern white and appear to be nothing more than large bumps. The length is about 2.5 cm. Duration of the pupal stage at 25°C is about 9.6 ± 0.1 days.

FIG. 10.129  Southern white butterfly. (Drawing by USDA.)

FIG. 10.128  Larvae of southern white butterfly. (Photo by L. Buss.)

Larva. Young are often gregarious, and such larvae grow faster than solitary individuals. Larvae are mottled gray or brownish green, often with a purplish hue, and bear five longitudinal orange or yellow-green bands running the length of the body. Rows of black spots also occur laterally. The head is yellow and orange, and marked with black spots. The larvae feed gregariously. A key for differentiating this caterpillar from other crucifer-feeding pierids is provided in Appendix A.

Adult. The adult has a wingspan of about 6–8 cm. The male butterfly is almost pure white dorsally but has a black, deeply indented band along the distal edge of the front wing. On the underside of the male, the hind wings are tan, as are the tips of the front wings. The female has two rather distinct color forms. The long-lived winter form is principally white, but with wider black bands than the male, and a single black spot on the forewings. During the summer months, the female is much darker, brown to dark gray; such butterflies tend to have a broader dark band on the forewings, but less scalloping of the band. Caterpillars raised under short-day conditions produce the dark adult form. A key for differentiating this species from other cabbage white butterflies is included in Appendix A. The biology of southern white was given by Opler and Krizek (1984) and Scott (1986). A key that includes southern white is Sparks and Liu (2001).

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Damage Larvae feed on foliage and eat small and large holes in leaves. In coastal areas, they sometimes defoliate small plantings or areas of fields. In Texas, this species is reported to be increasing in importance (Liu, 2005).

Management Hayslip et al. (1953) considered southern white to be a minor pest in Florida, as it is in most locations where it occurs. Management techniques appropriate for imported cabbage worm should be suitable for this insect also.

FAMILY PLUTELLIDAE— DIAMONDBACK MOTHS Diamondback Moth

Plutella xylostella (Linnaeus) (Lepidoptera: Plutellidae)

Natural History Distribution. Diamondback moth is probably of European origin but has become rather cosmopolitan, and is now found throughout the Americas and in Europe, Southeast Asia, Australia, and New Zealand. It was first observed in North America in 1854, in Illinois, but spread quickly. It was found in Massachusetts and Maryland by 1870, had spread to Florida and the Rocky Mountains by 1883, and was reported from British Columbia by 1905. In North America, diamondback moth is now recorded everywhere that cabbage is grown, even as far north as Canada’s Northwest Territories. By virtue of its ability to feed on cruciferous weeds, diamondback moth is sometimes abundant even in some areas where cruciferous crops do not occur. However, it is highly dispersive and is often found in areas where it cannot successfully overwinter, including most of Canada (Beirne, 1971). Host Plants. Diamondback moth generally is reported to attack only plants in the family Brassicaceae (Cruciferae). Virtually all cruciferous vegetable crops are eaten, including broccoli, Brussels sprouts, cabbage, Chinese cabbage, cauliflower, collards, kale, kohlrabi, mustard, radish, turnip, and watercress. Not all are equally preferred, however, and collards usually are chosen by ovipositing moths relative to cabbage. Several weeds are important hosts, especially early in the season before cultivated crops are available. Yellow rocket, Barbarea vulgaris; shepherdspurse, Capsella bursapastoris; pepperweed, Lepidium spp.; and wild mustards, Brassica spp., are commonly cited as attractive. However, yellow rocket is reported to attract ovipositing moths, but to be unsuitable for larval development (Shelton and Nault, 2004). Even more interesting are the reports from Africa

of P. xylostella expanding its host range from cruciferous plants to include feeding on snap pea and snow pea, Pisum sativum (Fabaceae) (e.g., Rossbach et al., 2006). Natural Enemies. A comprehensive analysis of diamondback moth mortality factors was conducted in Ontario by Harcourt (1960, 1963b). Large larvae, prepupae, and pupae often were killed by the parasitoids Microplitis plutellae (Muesebeck) (Hymenoptera: Braconidae), Diadegma insulare (Cresson) (Hymenoptera: Ichneumonidae), and Diadromus subtilicornis (Gravenhorst) (Hymenoptera: Ichneumonidae). All are specific on P. xylostella. In Ontario, D. insulare was considered most important except during diamondback moth population outbreaks when the other species assumed greater importance. More recent studies conducted in Quebec are consistent with the Ontario studies (Godin and Boivin, 1998b). Diadegma insulare was also important in California (Oatman and Platner, 1969; Kennedy and Oatman, 1976). Nectar produced by wildflowers is important in determining parasitism rates by D. insulare (Idris and Grafius, 1995). Egg parasites are poorly known. However, a Trichogramma egg parasitoid is reported in Japan (Wakisaka et al., 1992) and also from Florida (Leibee, pers. comm.). Fungi, granulosis virus, and NPV sometimes occur in high-density diamondback moth larval populations. Weather. Harcourt (1960, 1963b) and Wakisaka et  al. (1992) found that a large proportion of young larvae were often killed by rainfall. However, the most important factor determining population trends was reported to be adult mortality. Adult survival was thought to be principally a function of weather, though this hypothesis has not been examined rigorously. Life Cycle and Description. Total development time from the egg to pupal stage averages 25–30 days, depending on the weather (range about 17–51  days). In Ontario, diamondback moth is present from May to October but is most abundant in July and September. There are four to six generations annually, but discrete broods are not apparent (Harcourt, 1957). In Quebec, Godin and Boivin (1998a) reported adults and eggs beginning in early June and estimated three to four generations annually. In Colorado, the number of annual generations is estimated to be seven, and overwintering survival is positively correlated with the abundance of snowfall (Marsh, 1917). There is continuous breeding in the southern states, so the number of generations is likely 12–15 per year. Egg. Diamondback moth eggs are oval and flattened, and measure 0.44 mm long and 0.26 mm wide. They are yellow or pale green, and are deposited singly or in small groups of 2–8 eggs in depressions on the surface of foliage, or sometimes on other plant parts. Females may deposit 250–300 eggs early in the year, but the number decreases

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in later generations by 90%; average total production is probably 150 eggs. Development time in temperate regions generally averages 5.6 days (range 4–8 days), but mean development time is 3.0 ± 0.09 (± SE), 2.4 ± 0.07, and 2.3 ± 0.06 days when reared at 24°C, 28°C, and 30°C, respectively (Liu et al., 2002).

FIG. 10.131  Pupa of diamondback moth. (Photo by L. Buss.)

FIG. 10.130  Larva of diamondback moth. (Photo by L. Buss.)

Larva. Diamondback moth has four instars. Average and range of development time generally is about 4.5 (3–7), 4 (2–7), 4 (2–8), and 5 (2–10) days, respectively. However, when instars 1–4 are cultured at 24, 28, and 30°C, development times are 2.0 ± 0.05 (± SE), 1.6 ± 0.17, and 1.5 ± 0.03  days for instar 1; 2.2 ± 0.08, 1.4 ± 0.10, and 1.3 ± 0.04 days for instar 2; 1.5 ± 0.04, 1.3 ± 0.11, and 1.1 ± 0.04 days for instar 3; and 2.0 ± 0.04, 1.7 ± 0.28, and 1.5 ± 0.04 days for instar 4, respectively (Liu et al., 2002). Throughout their development, larvae remain quite small and active. If disturbed, they often wriggle violently, move backward, and spin down from the plant on a strand of silk. The overall length of each instar rarely exceeds 1.7, 3.5, 7.0, and 11.2 mm, respectively, for instars 1–4. Mean head capsule widths for these instars are about 0.16, 0.25, 0.37, and 0.61 mm. Larval body form tapers at both ends, and a pair of prolegs protrudes from the posterior end, forming a distinctive “V.” The larvae are colorless in the first instar, but thereafter they become green. The body bears relatively few hairs, which are short in length, and most are marked by the presence of small white patches. There are five pairs of prolegs. Initially, the feeding habit of first instar larvae is leaf mining, though they are so small that the mines are difficult to notice. The larvae emerge from their mines at the conclusion of the first instar, molt beneath the leaf, and thereafter feed on the lower surface of the leaf. Their chewing results in irregular patches of damage and the upper leaf epidermis is often left intact.

Pupa. Pupation occurs in a loose silk cocoon, usually formed on the lower or outer leaves. In cauliflower and broccoli, pupation may occur in the florets. The yellowish pupa is 7–9 mm long. The duration of the cocoon generally averages about 8.5 days (range 5–15 days), but during the initial time the insect is in the cocoon, in the prepupal rather than the pupal stage. For example, when cultured at 24°C, the duration of the prepupa is 0.6.7 ± 0.16 days (± SE), but the duration of the female pupal stage is 3.8 ± 0.23  days (± SE); when cultured at 28°C, the duration of the prepupa is 0.5 ± 0.18 days (± SE), but the duration of the pupal stage is 3.1 ± 0.24 days (± SE); when cultured at 30°C, the duration of the female prepupa is 0.6 ± 0.02 days (± SE), but the duration of the pupal stage is 2.6 ± 0.13 days (± SE) (Liu et al., 2002).

FIG. 10.132  Adult diamondback moth. (Photo by L. Buss.)

Adult. The adult is a small, slender, grayish-brown moth with pronounced antennae. It is about 6 mm long and marked with a broad cream or light-brown band along the back. The band is sometimes constricted to form one or more light-colored diamonds on the back, which is the

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basis for the common name of this insect. When viewed from the side, the tips of the wings can be seen to turn upward slightly. Moths usually mate at dusk, immediately after emergence from the cocoon. Flight and oviposition take place from dusk to midnight, and moths can be found feeding at blossoms on nectar. Adult males and females live about 12 and 16  days, respectively, and females deposit eggs for about 10 days. The moths are weak fliers, usually flying within 2 m of the ground, and not flying long distances. However, they are readily carried by the wind. The adult is the overwintering stage in temperate areas, but moths do not survive cold winters, as is found in most of Canada. They routinely re-invade these areas each spring, evidently aided by southerly winds (Smith and Sears, 1982). Detailed biology of diamondback moth can be found in Marsh (1917), Harcourt (1955, 1957, 1963b), and in Philips et al. (2014). A survey of the world literature was published by Talekar et  al. (1985). Rearing techniques were provided by Biever and Boldt (1971) and Liu and Sun (1984). Chow et  al. (1974) identified two major components of the sex pheromone, and Chisholm et  al. (1983) evaluated a four-component mixture, but the exact blend remains undetermined.

Damage Damage is caused by larval feeding. Although the larvae are very small, they can be quite numerous, resulting in complete removal of foliar tissue except for the leaf veins. They are particularly damaging to seedlings and may disrupt head formation in cabbage, broccoli, and cauliflower. The presence of larvae in florets can result in complete rejection of produce, even if the level of plant tissue removal is insignificant. Diamondback moth was long considered a relatively insignificant pest. Its impact was overshadowed by such ­serious defoliators as imported cabbage worm, Pieris rapae (Linnaeus), and cabbage looper, Trichoplusia ni (Hübner). However, in the 1950s the general level of abundance began to increase, and by the 1970s it became troublesome to crucifers in some areas. Although this shift in abundance has been attributed to the increased availability of alternative weed hosts or destruction of parasitoids, insecticide resistance was long suspected to be a component of the problem. This was confirmed in the 1980s as pyrethroid insecticides began to fail, and soon thereafter virtually all insecticides were ineffective (Leibee and Capinera, 1995). Relaxation of insecticide use, which can be implemented by use of thresholds to trigger applications, rotation with Bacillus thuringiensis and particularly elimination of pyrethroid use, can return diamondback moth to minor pest status by favoring survival of parasitoids.

Management Sampling. Populations are usually monitored by making counts of larvae, or by the level of damage. In Texas, average population densities of up to 0.3 larvae per plant are considered to be below the treatment threshold (Kirby and Slosser, 1984; Cartwright et al., 1987). In Florida and Georgia, treatment is recommended only when damage equals or exceeds one hole per plant (Workman et al., 1980). When growers monitor fields and subscribe to these treatment thresholds rather than trying to prevent any insects or damage from occurring in their fields, considerably fewer insecticide applications are needed to produce a satisfactory crop. Harcourt (1961) studied the distribution of various life stages on cabbage and recommended a minimum plant sample size of 40–50 except for the egg stage, where 150 plants should be examined for accurate population estimates. Shelton et al. (1994) compared the benefits of sequential and variable-intensity sampling for diamondback moth management and recommended the latter as being more reliable and requiring fewer samples. Pheromone traps can be used to monitor adult populations and may predict larval populations 11–21 days later. However, because of variation among locations, each crop field requires independent evaluation (Baker et  al., 1982a, b). Reddy and Guerrero (2000) documented the benefits of using pheromone traps as a decision-making tool, thereby reducing the number of insecticide applications. In this case, however, natural enemies (Chrysoperla carnea [Neuroptera: Chrysopidae] and Cotesia plutellae [Hymenoptera: Braconidae]), Bacillus thuringiensis, neembased insecticide, and chemical insecticide were integrated into a cabbage production system. So this IPM system was much more complex than simply using pheromone traps, but it resulted in financial savings for the producers as compared to a conventional production system. Insecticides. Protection of crucifer crops from damage often requires the application of insecticide to plant foliage, sometimes as frequently as twice per week. However, resistance to insecticides is widespread and includes most classes of insecticides including some Bacillus thuringiensis products. Rotation of insecticide classes is recommended, and the use of B. thuringiensis is considered especially important because it favors survival of parasitoids. Even B. thuringiensis products should be rotated, and current recommendations generally suggest alternating the kurstaki and aizawa strains because resistance to these microbial insecticides occurs in some locations (Tabashnik et al., 1990; Tang et al., 1996). Mixtures of chemical insecticides, or chemicals and microbials are often recommended for diamondback moth control (Leibee and Savage, 1992). This is due partly to the widespread occurrence of resistance, but also because pest complexes often plague ­crucifer

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crops, and the insects vary in susceptibility to individual insecticides. Botanical insecticides, especially neem products (extracts if Azadirachta indica trees), are sometimes suggested for diamondback moth suppression, though the effectiveness is limited. Charleston et al. (2005) compared neem extract with extract from the syringa tree, Melia azedarach, and reported that both extracts deterred larval feeding, but not oviposition. Sex pheromone, though not usually considered to be an insecticide, can also be used as a chemical crop protectant. Continuous release of pheromone has been investigated as a technique to suppress diamondback moth mating activity. McLaughlin et  al. (1994) reported that the number of insecticide treatments could be reduced from 13 to 15 to only three when crops were grown in the continuous presence of diamondback moth pheromone. Pheromone also can be formulated in a viscous matrix gel that contains insecticide (Maxwell et al., 2006), resulting in an attract-and-kill product. This product was shown to reduce damage when moth populations were at low to moderate levels, but not at high densities. The pheromone can also be used as part of an insect pathogen delivery system. Males can be attracted to traps dosed with the fungus Zoophthora radicans. After contacting the fungus and habituating to the pheromone, the moths disperse again, thereby transporting the fungus to nearby plants (Pell et al., 2008). Biological Control. Although over 130 parasitoid species plus numerous predators and pathogens are known from diamondback moth worldwide (Sarfraz et al., 2005), only a few species can be manipulated for effective biological suppression. The larval parasitoids Diadegma insulare (Cresson) (Hymenoptera: Ichneumonidae) and Microplites plutellae (Muesebeck) (Hymenoptera: Braconidae) are quite important in North America (Philips et  al., 2014). In warmer climates such as the southeastern United States, Oomyzus sokolowski (Kurdjumov) (Hymenoptera: Eulophidae) assumes importance as a larval parasitoid. Generalist predator such as rove beetles (Coleoptera: Staphylinidae), ground beetles (Coleoptera: Carabidae), paper wasps (Hymenoptera: Vespidae), flower flies (Diptera: Syrphidae), green lacewings (Neuroptera:Chrysopidae), brown lacewings (Neuroptera: Hemerobiidae), minute pirate bugs (Hemiptera: Anthocoridae), and spiders (Arachnida) are also important, often causing up to 90% mortality of first instar diamondback moths (Philips et  al., 2014). The best strategy for taking advantage of these natural enemies (both parasitoids and predators) is to avoid disrupting their populations by applying broad-spectrum insecticides. Entomopathogens also affect diamondback moth, but effective use is limited largely to the bacterium Bacillus thuringiensis, and the fungi Zoophthora radicans and

Beauveria bassiana. However, the fungi Isaria fumosorosea and Metarhizium anisopliae are sometimes used (De Bortoli et al., 2013). Entomopathogenic nematodes in the genera Steinernema and Heterorhabditis can infect and kill the larvae but do not persist well under field conditions, so get little use. Cultural Practices. Rainfall has been identified as a major mortality factor for young larvae, so it is not surprising that crucifer crops with overhead sprinkle irrigation tend to have fewer diamondback moth larvae than drip or furrow-irrigated crops. Irrigation also tends to disrupt oviposition. Best results were obtained with daily evening applications (McHugh Jr. and Foster, 1995). Crop diversity can influence the abundance of diamondback moth. Larvae generally are fewer in number and more heavily parasitized, when crucifer crops are interplanted with another crop or when weeds are present. This does not necessarily lead to reduction in damage, however (Bach and Tabashnik, 1990). Surrounding cabbage crops with two or more rows of preferred hosts such as collards and mustard or attractive weeds such as yellow rocket (Barbarea vulgaris) can delay or prevent the dispersal of diamondback moth into cabbage crops (Srinivasan and Krishna Moorthy, 1992; Badenes-Perez et al., 2004). Barbarea vulgaris also serves as a dead-end host, because although this species is preferred for oviposition, the larvae do not survive (Shelton and Nault, 2004). Crucifer transplants are often shipped long distances before planting, and diamondback moth may be included with the transplants. In the United States, many transplants are produced in the southern states and then moved north as weather allows. Cryptic insects such as young diamondback moth larvae are sometimes transported and inoculated in this manner. The transport of insecticide-resistant populations may also occur (Wyman, 1992). Every effort should be made to assure that transplants are free of insects before planting. For small plots of crucifers, protection from diamondback moth can be attained when frames supporting netting are placed over the plants. This approach can be more effective (and economic) than some insecticide treatments, and the effectiveness can be improved by impregnating the netting with insecticide (Martin et al., 2006). Repellent insecticides, a property often associated with pyrethroids, is especially useful. Also, it allows the use of larger mesh size netting, alleviating some of the disadvantages of covering plants, such as excessive temperature and humidity (Martin et al., 2013). Host-Plant Resistance. Crucifer crops differ somewhat in their susceptibility to attack by diamondback moth. Mustard, turnip, and kohlrabi are among the most resistant crucifers, but resistance is not as pronounced as it is for

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i­mported cabbage worm and cabbage looper (Radcliffe and Chapman, 1966). Varieties also differ in susceptibility to damage by diamondback moth, and a major component of this resistance is the presence of leaf wax. Glossy varieties, lacking the normal waxy bloom and therefore green rather than grayish green, are somewhat resistant to larval feeding (Dickson et al., 1990; Stoner, 1990; Eigenbrode et al., 1991). Larvae apparently spend more time searching, and less time feeding, on glossy varieties. Glossy varieties also tend to have fewer imported cabbageworm larvae and cabbage aphids, but more cabbage flea beetles.

factors at this time, once larvae began to burrow they were much less affected by such mortality agents. Life Cycle and Description. There are three generations annually in California, where considerable overlapping of generations occurs, and all stages of development can be found throughout the year. The number of generation in New York was reported to be one, whereas two were reported from Minnesota. Development time is quite long, requiring about 110–140 days for a complete life cycle, so it seems improbable that there was more than one generation in most of the North.

FAMILY PTEROPHORIDAE—PLUME MOTHS

Egg. The oval eggs are glossy and yellow, and measure 0.52–0.66 mm long and 0.26–0.35 mm wide. They are deposited singly and externally on various plant tissues, especially the undersides of leaves, and they may be attached either on the side or on end. Females prefer to attach their eggs among leaf hairs and avoid smooth substrates. Mean production was estimated at 170 eggs (range 70–300 eggs) by Lange Jr. (1941) but at about 250 eggs by Bari and Lange (1980). Duration of the egg stage varies with temperature but averages 15  days (range 8–24 days) during the spring in California.

Artichoke Plume Moth

Platyptilia carduidactyla (Riley) (Lepidoptera: Pterophoridae)

Natural History Distribution. This native insect is found widely in North America. Artichoke plume moth damages throughout the west, from British Columbia to southern California, but is infrequent in the Rocky Mountains and arid Great Plains regions. It is also known from eastern North America, but in Canada it is known only as far east as Ontario, and in the United States it occurs as far east as New York and North Carolina but is not common in the southeastern states. Host Plants. Artichoke plume moth attacks thistles in the family Compositae. Several weedy Cirsium species are suitable hosts, and some are preferred over globe artichoke. Other weeds also known to be hosts are milk thistle, Silybum marianum; and at least on one occasion Napa thistle, Centaurea melitensis. Despite the reported preference for weed species, artichoke is regularly attacked in California, where most artichoke is cultivated. Artichoke can serve as a host during the entire year, but a sequence of weedy thistle species is usually required in nonagricultural habitats. Lange (1950) and Turner et  al. (1987) provided lists of known-host plants. Natural Enemies. Several natural enemies are known, including a variety of Hymenoptera (Braconidae, Ichneumonidae), Diptera (Tachinidae), lacewing (Neuroptera: Hemerobiidae), rove beetles (Coleoptera: Staphylinidae), the whirligig mite, Anystis agilis (Banks) (Acarina), as well as spiders and birds. In California, Diadegma acuta (Viereck) (Hymenoptera: Ichneumonidae) is the most widespread and effective parasitoid, attacking larvae in their burrows. Other parasitoids were documented by Lange Jr. (1941) and Bragg (1994). Life table analyses conducted by Goh and Lange (1980b) suggested that much of the natural mortality occurred soon after egg hatch. Predation and drowning were important

FIG. 10.133  Larva of artichoke plume moth. (Drawing by USDA.)

Larva. Upon hatching, the larva measures about 1 mm long and bears four pairs of prolegs. Larvae often feed on young foliage during the first two instars and then burrow into the stalks or flower buds. Lacking suitable young foliage, however, they immediately begin to burrow. There are four instars. Mean duration (range) of the instars is 8 (8), 5.2 (5–6), 7.1 (5–12), and 19.4 (16–26) days, respectively. Thus, larval development times of 40–60 days are common, depending on weather. The larva is yellowish for the first three instars, and whitish during the fourth instar. During all instars the head, true legs, cervical shield, and anal plate are black. The head capsule widths are 0.24, 0.43, 0.67, and 1.06 mm for instars 1–4, respectively. Body length increases to about 12 mm at maturity, and the larva is rather stout. Pupa. The pupa varies from yellow to brown, and measures about 10–12 mm long. The abdomen bears tooth-like spines that project posteriorly. Pupation does not always occur within a cocoon, but a thin silken cocoon tends to be present if the larva pupates in an exposed location such as among old senescing leaves and leaf litter. Often the naked pupa is found within the burrow of the larva. Duration of the pupa is about 24 days (range 22–28 days).

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FIG. 10.134  Adult of artichoke plum moth. (Drawing by USDA.)

Adult. The adult is a small yellowish-brown moth with a wingspan of 18–30 mm. The forewing is lobed, with a cleft extending inward about one-third the length of the wing. The forewing also bears a dark triangular mark adjacent to the cleft and extending to the leading edge of the forewing. The hind wing is even more divided, consisting of three large lobes; the color is uniform brown. The legs are light in color except that they bear dark bands at the joints. Adults normally are not active during the day and tend to rest on the underside of foliage, but they make short flights if disturbed. The temperature threshold for flight is about 8–10°C. Mating usually occurs within 3  days of emergence, and adults display a preovipositional period of about 3–8 days. Adults live for up to 30 days. Females produce a sex pheromone (Klun et al., 1981) with maximal pheromone release about 4 h after sunset (Haynes et al., 1983). The most complete accounts of artichoke plume moth biology were supplied by Lange Jr. (1941, 1950). Bari and Lange (1980) provided information on developmental biology.

Damage The larvae may feed on any portion of the plant, but they are usually found in the developing flower head, or bud, of the artichoke. Small larvae may burrow through the tissue of the outer bracts. As the larvae mature, they tunnel into the inner portions of the fruit. Larvae sometimes feed on the leaf tissue, especially the newly formed foliar tissue at the center of the plant, and sometimes burrow into the stalks and crown, including short distances below-ground. Artichoke plume moth is often considered to be a serious limiting factor in commercial artichoke production.

Management Sampling. Distribution of eggs and larvae was studied by Goh and Lange (1980a). During the vegetative or prebloom stage, most eggs and young larvae were found on the young leaf tissue at the center of the plant. As blossoms are produced, however, eggs are deposited on the leaves

just ­below the flower heads, and virtually all the larvae are found within the buds. Moths can be captured with light traps. Sex pheromone can be used to bait traps for population monitoring, but can also be used to permeate the atmosphere and disrupt mating (Haynes et al., 1981). Insecticides. Foliar insecticides, including Bacillus thuringiensis, are often applied to protect artichoke from artichoke plume moth. The preferred materials affect both the adult and larval stages, and this often limits the use of Bacillus thuringiensis to tank mixes with chemical insecticides. Insecticide use sometimes induces outbreaks of twospotted spider mite, Tetranychus urticae Koch. Cultural Practices. Sanitation is an important element in artichoke plume moth management. Infested artichokes should be shredded, deeply buried, fed to livestock, or otherwise destroyed and not discarded in the field, because moths emerge successfully from infested buds. Similarly, destruction of plants after harvest can be beneficial if they are infested. Weedy thistles should be eliminated if found growing near cultivated artichoke as these weeds can provide inoculum for crops. Biological Control. The bacterium Bacillus thuringiensis and the entomopathogenic nematode Steinernema carpocapsae have been shown to provide effective suppression of artichoke plume moth larvae. The dense vegetative growth and tunneling behavior of larvae provide suitable microhabitat for nematode survival during the prebloom period, but not during the fruiting period (Bari and Kaya, 1984). At planting time, artichoke cuttings can be soaked in a suspension of entomopathogenic nematodes to reduce the likelihood of planting insects into a field along with the new crop.

FAMILY PYRALIDAE—PYRALID MOTHS Lesser Cornstalk Borer

Elasmopalpus lignosellus (Zeller) (Lepidoptera: Pyralidae)

Natural History Distribution. This species occurs widely in the western hemisphere, but not elsewhere. It is known from much of the southern United States, though in the eastern regions it occurs much farther north than in western states. The northernmost limits seem to be Massachusetts to southern Iowa in the east, and Oklahoma to southern California in the west. It is found throughout Central and South America. Despite its wide range, the damage is limited principally to sandy soil, so it tends to cause injury in the coastal plain of the southeastern states from South Carolina to Texas. Host Plants. Lesser cornstalk borer damages several crops grown in the southeast, though it is mostly a pest of peanut, sorghum, and soybean. Among vegetable crops

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i­njured are bean, beet, cabbage, cantaloupe, corn, cowpea, lima bean, pea, pepper, sweet potato, tomato, and turnip. Legume and grass crops are most often damaged. Field crops injured are corn, chufa, millet, oat, rice, rye, sorghum, peanut, soybean, sudan grass, sugarcane, tomato, turnip, and wheat. It also infests crabgrass, Digitaria sanguinalis, wiregrass, Elusine indica; and Johnsongrass, Sorghum halepense. Natural Enemies. Several natural enemies of lesser cornstalk borer are known, though they are not thought to be major determinants of population trends. Smith Jr. and Johnson (1989) constructed life tables for populations in Texas and identified survival of large larvae as the key element in generation survival, but the causative factor remained unidentified. The predominant parasitoids are Orgilus elasmopalpi Muesebeck and Chelonus elasmopalpi McComb (both Hymenoptera: Braconidae), Pristomerus spinator (Fabricius) (Hymenoptera: Ichneumonidae), and Stomatomyia floridensis Townsend (Diptera: Tachinidae) through most of the range of lesser cornstalk borer. Other parasitoids sometimes are present include Bracon gelechiae Ashmead (Hymenoptera: Braconidae), Geron aridus Painter (Diptera: Bombyliidae), and Invreia spp. (Hymenoptera: Chalcididae) (Johnson and Smith Jr., 1981; Funderburk et  al., 1984b; Smith Jr. and Johnson, 1989). Parasitoids rarely cause more than 10% mortality. Among the predators thought to be important mortality factors are a ground beetle, Plilophuga viridicolis LeConte (Coleoptera: Carabidae); Geocoris spp. bugs (Hemiptera: Lygaeidae); and larval stiletto flies (Diptera: Therevidae). Pathogens are commonly present in lesser cornstalk borer populations. The most important pathogen appears to be a granulosis virus, but a Beauveria sp. fungus, microsporidia, and mermithid nematodes have also been found (Funderburk et al., 1984b). Weather. Lesser cornstalk borer seems to be adapted for hot, xeric conditions, and therefore tends to be more abundant and damaging following unusually warm, dry weather. Mack et  al. (1993) used data from Alabama and Georgia to develop a predictive equation that forecasts the potential for crop injury and the need to monitor crops. It is based on the concept of “borer-days.” Borer-days is calculated as the sum of days during the growing season in which the temperature equals or exceeds 35°C and the precipitation is less than 2.5 mm, less the number of days in which the temperature is less than 35°C and the precipitation equals or exceeds 2.5 mm. Thus, it is the sum of the number of hot, dry days less the number of cooler, wetter days. If the number of borer-days equals or exceeds 10, the damage is likely. If borer-days equals 5–9, the damage is possible and fields should be scouted. The relationship between borerdays and larval abundance is nonlinear, and small increases in borer-days beyond 10 results in large increases in larval

abundance. Reduction in larval feeding at high soil moisture levels was documented by Viana and da Costa (1995). Life Cycle and Distribution. There are three complete generations annually in Georgia, and some members go on to form the fourth generation. Other southeastern states also experience three to four generations, but in the southwest there are only three generations annually. Activity extends from June to November, with the generations overlapping considerably and little evidence of breaks found between generations. Overwintering apparently occurs in the larval and pupal stage; diapause is not present. A complete life cycle usually requires 30–60 days. Egg. The egg is oval, measuring about 0.6 mm long and 0.4 mm wide. When first deposited, the egg is greenish, soon turns pinkish, and eventually reddish. The female deposits nearly all her eggs below the soil surface, adjacent to plants. A few, however, are placed on the surface or on leaves and stems (Smith Jr. et al., 1981). Duration of the egg stage is 2–3 days.

FIG. 10.135  A soil tube in which a lesser cornstalk borer dwells. (Photo by J. Castner.)

FIG. 10.136  A grass stem split to reveal a lesser cornstalk borer larva. (Photo by L. Buss.)

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Larva. Larvae live in the soil, constructing tunnels from soil and excrement tightly woven together with silk. They leave the tunnel to feed in the basal stalk area or just beneath the soil surface, returning and constructing new tunnels as they mature. Thus, tunnels often radiate out from the stem of the food source, just below the soil surface, to a depth of about 2.5 cm. Normally there are six instars, though 5–7 have been observed. During the early instars, larvae are yellowish green, with reddish pigmentation dorsally, tending to form transverse bands. As the larva matures, whitish longitudinal stripes develop, so that by the fifth instar they are pronounced. The mature larva is bluish green, but tends toward reddish brown, with fairly distinct yellowish-white stripes dorsally. The head capsule is dark, and measures about 0.23, 0.30, 0.44, 0.63, 0.89, and 1.2 mm wide, respectively, for instars 1–6. Larval lengths are about 1.7, 2.7, 5.7, 6.9, 8.8, and 16.2 mm, respectively. Mean development time is estimated at 2.8, 2.9, 3.1, 3.4, 401, and 8.9  days, respectively, for instars 1–6 when reared at 24°C. Total larval development time varies widely but normally averages about 20–30 days (Sandu et al., 2010). Pupa. At maturity, the larva constructs a pupal cell of sand and silk at the end of one of the tunnels. The cocoon measures about 16 mm long and 6 mm wide. The pupa is yellowish initially, turning brown and then almost black just before the adult emerges. It measures about 8 mm long and 2 mm wide. The tip of the abdomen is marked by a row of six hooked spines. Pupal development time averages about 9–10 days (range 7–13 days). Adult. Moths are fairly small, measuring 17–22 mm in wingspan. Sexual dimorphism is pronounced. The forewing of the male moth is yellowish centrally, bordered by a broad dark band bearing purplish scales. In females, however, the entire forewing is dark, sometimes almost black, but also bears reddish or purplish scales. The thorax is light in males but dark in females. The hind wings of both sexes are transparent with a silvery tint. Adults are most active at night when the temperature exceeds 27°C, relative humidity is high, and there is little air movement. Such conditions are optimal for mating and oviposition. Indeed, these activities cease if the temperature falls below 18–20°C. Mack and Backman (1984) studied adult longevity and fecundity in relation to temperature. Longevity varied from 20  days if held at 17°C, to 8 days if held at 32–35°C. Mean fecundity varied from only about 20 eggs per female when reared at 17°C, to a maximum of 110 eggs at 30°C, then decreased at higher temperatures. Mean fecundity per day was estimated at about 12 eggs. However, these values included an unknown proportion of individuals that did not reproduce. Other reports indicate that fecundity is about 100–450 eggs among females that successfully reproduce, with an average of about 200 eggs per female. Adult longevity under field conditions is estimated at about 10 days.

The biology was described by Luginbill and Ainslie (1917) and a review was published by Tippins (1982). Developmental data were given by Dupree (1965) and Leuck (1966). Rearing was described by Chalfant (1975). A sex pheromone blend was identified by Lynch et  al. (1984b). A key to stalk borers associated with corn in southern states was presented by Dekle (1976); this publication also includes pictures of the adults. A guide to common stalk boring caterpillars is also found in Appendix A. Sparks and Liu (2001) included lesser cornstalk borer in their key to caterpillar larvae affecting vegetables.

Damage Damage is caused by the larval stage which feeds upon, and tunnels within, the stems of plants. Normally the tunneling is restricted to the basal region of stalks, including the below-ground portion, and girdling may occur. In affected plants wilting is one of the first signs of attack, but buds may wither, and stunting and plant deformities are common. Plant death is not uncommon, and infested areas of fields often have a very thin stand.

Management Sampling. The egg stage is difficult to sample because they are small and resemble sand grains. Eggs can be separated by flotation, however (Smith Jr. et al., 1981). Larval populations are aggregated and can be separated from soil by sieving or flotation (Mack et  al., 1991; Funderburk et al., 1986). Adults are attracted to light traps but are difficult to monitor with this technique because the moths of lesser cornstalk borer are difficult to distinguish from many other species. This is especially true of the females, which are less distinctive than the males. Pheromone traps have been used successfully to monitor adult populations, and adults can be flushed from fields by beating the vegetation. Adult pheromone trap catches and flush counts are correlated (Funderburk et al., 1985). Adult and larval counts are often highly correlated, indicating that flush counts can be used to predict the abundance of larvae in subsequent weeks (Mack et al., 1991). Loera and Lynch (1987) successfully used pheromone trapping to monitor moth populations in bean, and reported that trap heights of about 0.5 m were most effective. Insecticides. Insecticides applied for suppression of lesser cornstalk borer are usually applied in a granular formulation in the seed furrow or in a band over the seedbed. Liquid formulations can also be applied, but it is important that they are directed to the root zone. Cultural Practices. Lesser cornstalk borer damage is largely restricted to sites with sandy soil. Modified planting practices have long been used to minimize crop loss in such locations. Populations tend to increase over the course of a season, so some damage can be avoided by early planting.

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Tillage and destruction of weeds are recommended before planting because this helps to destroy larvae that may be present in the soil and damage seedlings, the stage most susceptible to destruction. However, crop culture that uses conservation tillage (i.e., retention of crop residue at the soil surface) experiences little injury from lesser cornstalk borer feeding because the larvae feed freely on crop residue and other organic matter, sparing the young crop plants (All and Gallaher, 1977; All et al., 1979).

Limabean Pod Borer

Etiella zinckenella (Treitschke) (Lepidoptera: Pyralidae)

Natural History Distribution. Limabean pod borer is found throughout the world but is most common in the tropics and subtropics. It occurs in the warmer regions of the temperate zone but is absent from cold climates such as northern Europe. In North America, limabean pod borer is widespread in the western United States, from Washington to southern California and east to Colorado and Texas. Limabean pod borer is known from certain eastern states, such as Florida, North and South Carolina, and Maryland, but is not considered to be a serious pest in the East. It is a problem, however, in the Caribbean, including Puerto Rico. In Canada, though limabean pod borer has been detected, it is not considered to be a field pest. It was first observed in the United States in California, in 1885, but its origin is unknown. Host Plants. This insect limits its attack to plants in the family Leguminosae. Vegetables damaged by limabean pod borer include cowpea, faba bean, snap bean, lima bean, pea, and pigeon pea. Other legumes such as lupine, Lupinus spp.; rattlebox, Crotalaria sagittalis and C. incana; locoweed, Astragalus antiselli; and milkvetch, Astragalus trichopodus, also serve as hosts. In California, severity of limabean pod borer damage is inversely related to the distance between crops and wild hosts, particularly lupine (Stone, 1965). In Puerto Rico, Crotalaria spp. are important alternate hosts (Segarra-Carmona and Barbosa, 1988). Natural Enemies. Limabean pod borer is not heavily parasitized in North America. Several native egg and larval parasitoids (Hymenoptera: various families) have been detected in California and Washington, but none are effective. Numerous parasitoid species are known from Europe (Parker, 1951; Stone, 1965), but attempts to introduce parasitoids into California and Puerto Rico during the 1930s were unsuccessful (Clausen, 1978). Life Cycle and Description. The number of generations varies according to the weather. The complete life cycle requires at least 60 days, and often considerably longer during cool weather. In California, one generation usually occurs

during March-June on lupines or other wild host plants, and then another two to four generations occur during JuneDecember on crop plants or perennial lupines. Starting in about July, some of the mature larvae diapause in their cocoons. The proportion of larvae entering diapause in July is small, only about 8%, but the proportion increases to 53% in August, 89% in September, and 100% in October and later. Pupation of overwintering larvae occurs from January to March. The emergence of adults from overwintering larvae begins in March, but it is protracted. Egg. The egg is oval and measures about 0.6–0.7 mm long and 0.3–0.45 mm wide. The egg is white when first deposited, but turns pink and then gray as the embryo develops. Duration of the egg stage is about 15.4 days but varies from 5 to 33 days depending on temperature. The eggs are deposited singly or in small groups of up to 12. They are deposited on the flower, stems, or pod petiole. Estimates of egg production vary widely. Laboratory-reared moths often produce only 50–90 eggs, whereas field-collected moths may produce about 140–260 eggs. The latter values are probably much better estimates of fecundity.

FIG. 10.137  Larva of limabean pod borer. (Drawing by USDA.)

Larva. The larva is white or cream at hatching, and measures only 1 mm long. Young larvae immediately bore into a bean pod and develop internally. The entrance hole into the pod usually heals, leaving no indication that a larva was feeding within. There are five instars, with mean head capsule widths of about 0.15, 0.35, 0.65, 1.05, and 1.60 mm, respectively. Duration of the instars is about 3, 3, 3, 3, and 4 days, respectively. The mature larva measures 12–17 mm long and is pinkish or tan. The head and pronotum are yellow with black markings. The larva has five pairs of prolegs

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in addition to three pairs of thoracic legs. Duration of the larval stage is usually about 35 days, but it may vary from 13 to 65 days. At maturity, the larva eats through the wall of the pod, exits, and drops to the soil. Pupa. The mature larva burrows into the soil to a depth of 1–5 cm and spins a cocoon. The soil particles adhere to the cocoon, so if dug from the soil the cocoon is an elongate cylinder of soil measuring 15–20 mm long and 6–8 mm wide. The duration of the prepupal period is variable, but during the summer months, it is typically 8–24 days. During the winter, of course, it is greatly prolonged, because this is the overwintering stage. The pupa, which eventually develops in the cocoon, is 8–10 mm long and 2.5–3 mm wide. Usually, it is amber or light brown. Duration of the pupal period is about 36 days (range 16–101 days).

FIG. 10.138  Moth of limabean pod borer. (Drawing by USDA.)

Adult. The adult is a small brownish-gray moth, with a wingspan of 24–27 mm. The most distinctive features are the forward-protruding mouthparts, a characteristic feature of pyralid moths, and a broad white band along the leading edge of the forewings. There is also a transverse yellowish band slightly anterior of the midpoint of the wing. Copulation commences about 24 h after emergence. The preoviposition period is usually 4–6 days, and adult longevity is about 10–12 days. The biology of limabean pod borer was described by Hyslop (1912b), Abdul-Nasr and Awadalla (1957), and Stone (1965). Rearing procedures were outlined by Hattori and Sato (1983).

Damage The larvae feed on buds and blossoms, and burrow into the pods of legumes to feed on the developing seeds. They typically feed on only a portion of seed and then move on to attack adjacent seeds. Silk and fecal material accumulate in the pods. In the case of small pods, larval feeding usually causes the pod to drop from the plant, but large pods remain attached. Larvae sometimes leave one pod and move to another to continue feeding, especially following pod drop. Market standards often cause a loss in excess of the direct feeding injury by larvae, because even low levels of

damaged pods or beans are considered to be undesirable. Formerly, this insect was considered to be an important pest in the western United States, but with the introduction of modern insecticides it has assumed minor status. In tropical climates, however, it remains a serious pest of beans and soybeans.

Management Sampling. Light traps can be used to sample populations of moths. Also, sex pheromone has been identified and used successfully to trap moths in the field (Toth et al., 1989). Insecticides. Residual insecticides provide good control of borers. As the larvae feed internally, it is essential that the insecticides are on the vegetation at the time moths are ovipositing and eggs are hatching. Protracted emergence of adults and multiple overlapping generations often necessitate numerous applications of insecticides. Bacillus thuringiensis is not usually recommended for this insect. Cultural Practices. Because larvae overwinter in the soil beneath legumes, tillage can reduce emergence in the spring. Autumn plowing to a depth of at least 20 cm is recommended. Early season planting is also helpful because the crop can reach maturity before pod borers attain high densities. Host-Plant Resistance. Host-plant resistance is a viable option for some crops. Several pigeon pea and soybean cultivars have been shown to exhibit considerable resistance to attack (Cruz, 1975; Armstrong, 1991; Talekar and ­Lin, 1994).

FAMILY SESIIDAE—VINE BORERS AND CLEARWING MOTHS Squash Vine Borer

Melittia cucurbitae (Harris)

Southwestern Squash Vine Borer

Melittia calabaza Duckworth and Eichlin (Lepidoptera: Sesiidae)

Natural History Distribution. Squash vine borer, M. cucurbitae, is found throughout the United States east of the Rocky Mountains and is also known from Central America. In Canada, it occurs in southern Ontario. In the southwest (Texas, New Mexico, and Arizona), M. cucurbitae is replaced by M. calabaza (Eichlin and Duckworth, 1988), which also occurs in portions of Mexico. This latter species is not well studied, but seems to have biology similar to M. cucurbitae; the only known exception is host range, as noted below. Further south in Mexico and in Central America, Melittia calabaza is replaced by M. putchripes Walker (which often has been referred to as M. satyriniformis Hübner).

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Host Plants. Squash vine borer feeds on wild and cultivated species of Cucurbita. Summer squash, Cucurbita pepo, and some winter squash, C. maxima, are most preferred for oviposition by adults, and most suitable for larval development. Some pumpkin, C. mixta, are intermediate in suitability. Winter squash derived from C. moschata does not support complete larval development. These plant species vary in susceptibility, however, with some cultivars quite resistant and others susceptible, so the local determination of susceptibility of specific cultivars is important information. Among weeds, C. texana and C. andreana, are very suitable hosts, while C. okeechobeensis is intermediate and some cucurbit weed species (C. digitata, C. foetidissima) are unsuitable (Howe and Rhodes, 1973). Other cucurbit crops such as cucumber and melon may be attacked, but this occurs mostly when the more favored hosts are not present (Friend, 1931). Southwestern squash vine borer displays basically the same host preferences (Sondak, 1981). However, M. ­calabaza, though generally having a host range similar to M. cucurbitae, was able to complete larval development in C. moschata and was not found in C. mixta (Eichlin and Duckworth, 1988). Natural Enemies. The natural enemies of squash vine borer are not well studied. General predators such as robber flies (Diptera: Asilidae) are reported to prey upon adults, and a wasp egg parasitoid, Telenomus sp. (Hymenoptera: Scelionidae) is known. Larvae seem to be relatively free of natural enemies (Friend, 1931). Life Cycle and Description. The squash vine borer completes its life cycle in about 60 days. In the southern states, it has two generations per year. In New York, New Jersey, and other cold climates, only a single generation occurs regularly. In Ohio, most borers diapause after the first generation, but some go on to complete a second generation (Smith, 1893). In the spring, emergence of adults is protracted. There has been considerable confusion concerning the number of generations in northern states because in the field it is difficult to discern a single protracted generation from two overlapping generations. In North Carolina, the first generation is reported to occur from April to June and the second from July to September (Smith, 1910a, b). In Connecticut, squash vine borer may not be observed until June or July and persist until September (Friend, 1931). Thus, each generation may remain active for 2 or 3 months, and overlapping generations are normal. Egg. Oval eggs, reddish brown in color, are glued to the plant tissue and are slightly flattened at the point of attachment. They measure about 1.1 mm long and 0.85 mm

wide. Each female is capable of producing 150–200 eggs. They are distributed singly on all parts of the plant except the upper surface of the leaves; most appear to be attached to the basal region of the vine, probably because this tissue is oldest and therefore exposed longer. The basal part of leaf petioles receives a moderate number of eggs. Some eggs are deposited in cracks in the soil near the base of the plant. Eggs hatch in 10–15 days and larvae soon bore into the plant. Those that hatch on the vine usually remain in this location to feed, whereas those hatching on leaf petioles eventually work their way to the base and enter the vine.

FIG. 10.139  Larva of squash vine borer. (Photo by J. Capinera.)

Larva. Larvae are white with a dark-brown or black head. Initially, the larval body is markedly tapered posteriorly and equipped with numerous large hairs. As the larva matures it loses its tapered shape and hairy appearance and acquires a dark thoracic shield. There are four instars, with head capsule widths of about 0.4, 0.7, 1.3, and 2.0 mm, respectively. The larva requires 4–6 weeks to complete its ­development and eventually attains a length of about 2.5 cm. The mature larva exits the plant tissue and burrows into the soil to a depth of 4–5 cm where pupation and formation of a black cocoon occurs. Pupa. The pupa is mahogany brown and the anterior end is equipped with a short but sharply pointed cocoon breaker, a structure used to cut a hole in the cocoon when the moth is ready to emerge. There is some disagreement whether squash vine borer always pupates soon after producing the cocoon. Friend (1931), working in Connecticut, reported that the fully grown larva overwintered in the cocoon, and pupated in the spring before producing the single northern generation. In contrast, Smith (1910a, b) maintained that among two-generation populations in North Carolina the pupa overwintered. In any event, a distinctive, heavy-­ bodied moth eventually emerges from the cocoon.

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occur. Commercial cucurbit production rarely suffers significant damage by squash vine borer. These borers seem to plague small plots especially severely; almost all home gardeners have had some unpleasant experience with squash vine borers.

Management

FIG. 10.140  Moth of squash vine borer. (Photo by J. Capinera.)

Adult. The wingspan of the moth is about 3 cm. The front wings are blackish and tinged with olive green while the hind wings are colorless. The abdomen is usually dull orange and marked with black dots dorsally, though on some specimens the abdomen is entirely black. The hind legs are prominently tufted and are orange and black. Adults are capable of oviposition about 3 days after emergence. Moths are active during the daylight hours, especially in the early morning hours. Although the moths of these two species are very similar in appearance, the second abdominal segment is orange dorsally in M. calabaza, whereas it is olive green in M. cucurbitae. Friend (1931) gave a detailed summary of the life history of squash vine borer. Sondak (1981) and Middleton (2018) provide summaries of the biology of southwestern squash vine borer and squash vine borer, respectively. Sparks and Liu (2001) included this species in their key to vegetableinfesting caterpillars. Good rearing techniques for these insects are unknown.

Damage The damage caused by these insects is accurately described by the common name—squash vine borer. The larva spends almost its entire life feeding within the plant stem. Because larvae feed within plant tissues they are hidden from view and easily overlooked. However, upon close examination frass can be found accumulating beneath small entrance holes. The presence of one, or even several, larvae is not always deadly to the plant. However, up to 142 larvae have been removed from a single plant, and obviously, such large numbers can disrupt the physiology of the plant beyond its ability to compensate successfully for the feeding injury. Often the first sign of infestation is wilting of the plant in the heat of the day while other plants remain turgid. If the vine is thin or heavily infested the portion beyond the feeding site of the larva(e) may be killed. Infestation of fruit can

Sampling. Shallow pans painted yellow and filled with water can attract and capture squash vine borer moths. Although water pan traps are recommended by investigators in Ontario for population monitoring, it is not known whether such traps are attractive enough to be used as a method of population reduction (McLeod and Gualtieri, 1992). A sex pheromone produced by females has been identified (Klun et  al., 1990). Male moths of M. cucurbitae may be attracted to traps baited with the insect’s sex pheromone, but thus far pheromone technology has not been demonstrated to improve management practices. In fact, one study reported the increased injury of squash plantings when pheromone was released; the biological basis for this is uncertain (Pearson, 1995). Melittia calabaza is also attracted to some components of the M. cucurbitae pheromone. Insecticides. Squash vine borer can be difficult to control with chemical insecticides because it is protected from contact with most insecticides by its burrowing activity. Therefore, it is imperative that insecticide residues are in place during oviposition so that when the larva hatches from the egg it has an opportunity to make contact with a lethal dose. Pollinators, particularly honeybees, are very important in cucurbit production, and insecticide application can interfere with pollination by killing honeybees. If insecticides are to be applied when blossoms are present, it is advisable to apply insecticides late in the day, when honeybee activity is minimal. Cultural Practices. It has long been known that in the north, early planted crops experience minimal injury, so early planting is recommended. Also, the moths preferentially attack certain species such as summer squash; this can be used as a trap crop if the plants are then destroyed or sprayed, thereby protecting less preferred cucurbit species or preferred crops planted later. In the southwestern areas, however, it may be advisable to wait until the spring oviposition period has occurred before planting susceptible squash, as in these warm-weather areas the summers are long enough to plant squash well into the summer. Deep plowing can disrupt the overwintering borer population. Overwintering insects, whether larvae or pupae are often damaged by plowing, and when they are brought to the soil surface they are susceptible to predation by vertebrates.

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Cucurbit plants are very resilient, recovering well from injury. If accumulated frass can be located adjacent to vine tissue, indicating the presence of active feeding, larvae can be killed or removed with a knife without severely affecting the growth of the remaining tissue. Vines also develop additional roots if covered with soil, which can compensate for damage by borers. Covering the vines near the base of the plant discourages oviposition near critically important tissue. Dying vines should be removed and destroyed (Britton, 1919). Sondak (1981) suggested also cutting back the new growth of young squash plants to destroy apical dominance in the plants, as this induces production of squash vines. The vines can be lost to feeding injury without the death of the plants, so the production of additional vines plus mounding of soil around the base of the plant to deter oviposition in a damage-­ susceptible area reduces the negative impacts of the insects. Biological Control. Using a medicine dropper or similar device, the bacterium Bacillus thuringiensis, and the entomopathogenic nematode Steinernema carpocapsae, can be injected into squash vines to kill existing larvae or to prevent their establishment. The hollow, moist vines are especially conducive to the spread and survival of nematodes.

Diptera: Tachinidae) (Arnaud Jr., 1978). The aforementioned tachinids attack the larval stage and have a wide host range; they have also been reared from many other species of Lepidoptera. Life Cycle and Description. The biology of this insect is not well documented. The number of generations is reported to be at least two in North Carolina, and probably more in Gulf Coast states. Egg. The eggs are deposited singly on the underside of the foliage. They are nearly spherical in shape, whitish or greenish, and measure about 1.35 mm in diameter. The female is reported to produce about 30–40 eggs. Duration of the egg stage is 6–10 days.

FAMILY SPHINGIDAE—HORNWORMS AND SPHINX MOTHS Sweetpotato Hornworm

Agrius cingulatus (Fabricius) (Lepidoptera: Sphingidae)

FIG. 10.141  Young larva of sweetpotato hornworm. (Photo by L. Buss.)

Natural History Distribution. Sweetpotato hornworm is native to the western hemisphere, where it is found commonly throughout the tropical and subtropical areas. In the United States, it occurs widely in the southern states and is found through most of the year in Florida and southern Texas. Each summer the moths are found as far north as Arkansas, and occasionally a stray is found in such northern locations as Michigan and Nova Scotia. Sweetpotato hornworm also occurs in Hawaii and Puerto Rico. This species has recently appeared in Africa, and it is believed that sub-Saharan Africa will prove to be a suitable environment for this insect (Ballesteros-Mejia et al., 2011). Host Plants. Larvae feed on various species of Ipomoea in the plant family Convolvulaceae. Among vegetable crops, only sweet potato is injured. The adults feed on nectar from various deep-throated flowers. Natural Enemies. The eggs of this species are heavily parasitized by Trichogramma semifumatum (Perkins) (Hymenoptera: Trichogrammatidae). Several fly parasitoids are known, including Agryophylax spp., Belvosia bifasciata (Fabricius), Chaetogaedia monticola (Bigot), Drino inca (Townsend), D. incompta (Wulp), and D. rhoeo (Walker) (all

FIG. 10.142  Older larva of sweetpotato hornworm. (Photo by L. Buss.)

Larva. Young larvae measure only about 3 mm long when they hatch and are greenish white with white granulations. The head, however, is greenish yellow. There are five instars and the larvae attain a length of about 115 mm at maturity. Head capsule widths are about 0.5, 1.0, 1.8, 3.2, and 6.0 mm, ­respectively. Duration of the larval stage is about 30–60 days, depending on temperature. The appearance of the larva is variable, the body color ranging from light green or brown to ­almost black. Typically, in the early instars the body is green

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and marked by oblique black stripes along each side on a whitish or light background, or with a narrow white band along the lower edge of the oblique stripes. The oblique lines may also form “V”-shaped lateral markings. The spiracles and anal appendage, or “horn” are blackish. In the final instar, the larva may remain green or becomes mostly brown. The green form has dark bands on the head capsule, and the oblique bands are found in the earlier instars; the posterior oblique band terminates in the horn. In the brown form, the body is gray with brown spots. The lateral oblique lines are weak, lighter beneath, and less likely to be “V”-shaped markings. The black spiracles are surrounded by a dark ring and give the appearance of very large spiracles. The prolegs are dark brown or purplish. As in the green form, the head is marked with dark bands and the posterior oblique lines terminate in the dark horn.

sometimes they have been so abundant as to defoliate entire fields and to acquire the “armyworm”-like habit of dispersing in groups (Watson, 1944).

Pupa. At maturity, the larva pupates in the soil. The pupa is dark brown or mahogany red and measures 55–65 mm long. Duration of this stage is 20–30 days.

Natural History

FIG. 10.143  Moth of sweetpotato hornworm. (Photo by L. Buss.)

Adult. The moth is sometimes known as the “pinkspotted hawk moth.” The adult is fairly typical in form for sphingids, heavy bodied with long pointed front wings, and short hindwings. The front wings are dark gray mottled with brown and black. The hind wings are pinkish basally with 2–3 dark bands that run parallel to the wing border. The abdomen bears a dark band dorsally and transverse pink or rose transverse bars on each side. The moth measures about 45 mm long and 90–100 mm in wingspan. Moths have often been observed feeding from flowers at dusk. The biology of the sweetpotato hornworm was given by Fullaway (1911), with additional notes provided by Hodges (1971). Larvae were described by Dyar (1895).

Damage The larvae of this species feed on foliage. They are rarely abundant enough to be considered a serious pest, though

Management Larvae are easily suppressed with foliar insecticides, including the microbial insecticide Bacillus thuringiensis.

Tobacco Hornworm

Manduca sexta (Linnaeus)

Tomato Hornworm

Manduca quinquemaculata (Haworth) (Lepidoptera: Sphingidae)

Distribution. The tobacco and tomato hornworms are very similar in appearance, biology, and distribution. However, the tobacco hornworm, Manduca sexta (Linnaeus), is more common in the southern United States, especially the Gulf Coast states. Its range extends northward to New York, and even sometimes into southern Ontario. The tomato hornworm, Manduca quinquemaculata (Haworth), in contrast, is uncommon along the Gulf Coast, but relative to tobacco hornworm it is more likely to be encountered in northern states and southern Canada. These are native species, occurring from the Atlantic to the Pacific Oceans, though for tobacco hornworm the range extends south into Central and South America, and the Caribbean. Host Plants. These insects feed almost entirely on solanaceous plants, particularly tomato and tobacco. They are recorded from other vegetables such as eggplant, pepper, and potato, but such feeding is unusual. Several solanaceous weeds are reported to serve as hosts, including groundcherry, Physalis spp.; horsenettle, Solanum carolinense; jimsonweed, Datura stramonium; and nightshade, Solanum spp.; but wild hosts are unimportant larval food sources relative to crops. Adults imbibe nectar from flowers of several plants such as catalpa, Catalpa speciosa; daylily, Hemerocallis sp.; hollyhock, Althaea rosea; jimsonweed; 4 o’clock, Mirabilis jalapa; mallow, Hibiscus lasiocarpus; mimosa, Albissia julibrissin; and tobacco. Despite the long history of these species being associated exclusively with Solanaceae, in Arizona, M. sexta is reported to ovipoit, feed, and develop successfully on Proboscidea spp. (Martyniaceae), which are taxonomically distant from Solanaceae (Mechaber and Hildebrand, 2000). Natural Enemies. Many natural enemies are known from tobacco and tomato hornworm. Lawson (1959) provided a good summary of their importance in North Carolina. Natural enemies of the egg stage include Trichogramma sp. (Hymenoptera: Trichogrammatidae) and Telenomus sp. (Hymenoptera: Scelionidae), but these are thought to be

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of little importance. Of greater importance are the stilt bug Jalysus spinosus (Say) (Hemiptera: Berytidae) and other general predators such as big-eyed bugs (Hemiptera: Lygaeidae) and lacewings (Neuroptera: Chrysopidae). The larvae often are parasitized by Cotesia congregatus (Say) (Hymenoptera: Braconidae), and they sometimes are seen bearing clusters of white pupal cases formed by this wasp, attached to their bodies. In North Carolina, 30%–40% of hornworms may be parasitized by this wasp. In contrast to this, one species of wasp accounting for most of the parasitism by Hymenoptera, numerous insects in the order Diptera, particularly the Tachinidae, attack tobacco and t­omato hornworms. Records of tachinid parasitism from t­ omato hornworm include Compsilura concinnata (Meigen), Drino incompta (Wulp), D. rhoeo (Walker), Lespesia frenchii (Williston), Winthemia leucanae (Kirkpatrick), and W.  quadripustulata (Fabricius). Records of tachinids from tobacco hornworm include Carcelia spp., Drino incompta, D.  rhoeo, Lespesia spp., Metavoria orientalis Townsend, and Winthemia quadripustulata. In North Carolina, the proportion of hornworm larvae parasitized by tachinids increases during the season, sometimes attaining 100%. The number of tachinid eggs per hornworm larva also increases seasonally, indicating a greater abundance of flies. Also of great significance as a mortality agent of larvae is Polistes spp. (Hymenoptera: Vespidae). These wasps kill and consume, or sometimes carry away to provision their nest, a very high proportion of larvae. Apparently, Polistes spp. is the most important larval mortality factor in North Carolina. Pathogens sometimes affect hornworms. A bacterial disease which causes the larva to blacken and to droop is sometimes reported to kill larvae, but this does not occur regularly and is observed mostly at very high larval densities. Larvae are also susceptible to infection by the fungus Entomophaga aulicae, but this disease also does not occur widely (Morrow et al., 1987). Vertebrates may prey on the pupae. Skunks and moles are particularly important. Tachinids emerge from the prepupal and pupal stages but attack only the larvae. Life Cycle and Description. Although these insects are common, they are not serious pests except in home gardens. Thus, there is surprisingly little information about these insects, and particularly about tomato hornworm. The following information applies specifically to tobacco hornworm except where noted but apparently is equally applicable to tomato hornworm. The number of annual generations ranges from 1 to 2 in Canada to 3–4 in northern Florida, but two generations per year is common over most of the ranges of these species. The proportion of insect that enters diapause increases from about 5% in June to 95% in mid-August, as day length decreases. In northern Florida the insects are active from April to November, but they are abundant only for the first two generations because many pupae enter diapause. In North Carolina they occur from mid-May to October, and

e­vidence of a third generation is poor. Throughout their range, hornworms overwinter in the pupal stage, and in the northernmost parts of their range they may not overwinter successfully, occurring only after dispersal northward during the summer. The life cycle can be completed in 30– 50 days, but often is considerably protracted. Egg. The eggs are spherical to oval and measure 1.25– 1.50 mm in diameter. They are smooth and vary in color from light green or yellow when they are early in development, to white at maturity. They are deposited principally on the lower surface of foliage, but also on the upper surface. Mean fecundity was reported by Madden and Chamberlin (1945) to be about 250–350 eggs with seasonal fluctuations and maximum egg production at mid-season. However, Yamamoto (1968) showed that with adequate adult nutrition, the fecundity of nearly 1400 eggs per female could be attained. Duration of the egg stage is 2–8 days but averages 5 days.

FIG. 10.144  Larva of tobacco hornworm. (Photo by J. Capinera.)

FIG. 10.145  Larva of tomato hornworm. (Photo by P. Choate.)

Larva. The larva is cylindrical and bears five pairs of prolegs in addition to three pairs of thoracic legs. The most striking feature of the larva is a thick-pointed structure or “horn” located

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dorsally on the terminal abdominal segment. The “horn” is as long as the body in newly hatched larvae but becomes relatively smaller as the larva matures.Young larvae are yellowish white but during the second instar become green with white lines laterally. The tobacco hornworm develops seven straight oblique whitish lines laterally. The white lines are edged with black on the upper borders, and the “horn” is usually red. The tomato hornworm is superficially similar, but instead of the seven oblique lateral bands it bears eight whitish or yellowish “V”-shaped marks laterally, and pointing anteriorly. The “V”-shaped marks are not edged in black. Also, in tomato hornworm the “horn” tends to be black. The body of both species is usually light green, but occasionally dark brown or blackish forms occur. There are normally five instars, but occasionally six are observed. The mean head capsule width is 0.8, 1.2, 2.0, 3.0, and 5.0 mm for instars 1–5, respectively. Corresponding mean larval body lengths are 6.7, 11.2, 23.4, 49.0, and 81.3 mm, respectively. Larval development time averages about 20 days, but ranges from 13 to 44 days depending on temperature. Mean development time for larvae reared under insectary conditions in northern Florida is 3.4, 2.9, 3.0, 3.9, and 6.6 days for instars 1–5, respectively. Pupa. Mature larvae drop to the soil at maturity and burrow to a depth of 10–15 cm. There they form a pupal cell measuring about 7 cm long and 4 cm wide, and pupate within. The interval between entering the soil and pupation is usually 4–8 days. The pupa is large and elongate-oval but pointed at the posterior end. It measures 45–60 mm long and 13–14 mm wide. The pupa bears a pronounced maxillary loop, a structure which encases the mouthparts. The maxillary loop in tobacco hornworm extends back about one-fourth the length of the body, whereas in tomato hornworm it is longer, usually extending for about one-third the length of the body. The color of the pupa is brown or reddish brown. Duration of the pupal stage is protracted and variable. Pupal duration often exceeds 100  days, even in the summer generations, but may be as short as 15  days, and 21 days is about average. Overwintering pupae do not emerge synchronously, with emergence spread from May to early August; a few even diapause through two winters.

FIG. 10.146  Moth of tomato hornworm. (Photo by J. Capinera.)

FIG. 10.147  Moth of tobacco hornworm. (Photo by J. Capinera.)

Adult. The adults of both species are large moths with stout, narrow wings, and a wingspan of 80–130 mm. The front wings are much longer than the hind wings. The females are larger than the males and can be differentiated by the narrower antennae. Both species are dull grayish or grayish brown, though the sides of the abdomen are usually marked with six orange-yellow spots in tobacco hornworm and five spots in tomato hornworm. The hind wings of both species bear alternating light and dark bands. Adults become active at sunset when they can be observed feeding at flowers. Although some adults remain active throughout the night, most activity occurs early in the evening and just before dawn. Both hornworm species likely produce sex pheromone, but only that of the tobacco hornworm has been identified. The preoviposition period of moths is about 2 days, and eggs are deposited for about 4–8 days. A detailed description of tobacco hornworm biology was given by Madden and Chamberlin (1945), and a more abbreviated but useful description of both species was established by Gilmore (1938). Diapause development was reported by Rabb (1966). Rearing was described by several authors, including Yamamoto (1968, 1969), Stewart and Baker Jr (1970), and Bell and Joachim (1976). Keys to the adults were included in Hodges (1971).

Damage Larvae are defoliators, and usually, attack the upper portion of plants initially. Rather than chewing holes in the leaf, they usually consume the entire leaf. Sometimes they attack green fruit. As the larvae of hornworms attain such a large size, they are capable of high levels of defoliation. Madden and Chamberlin (1945) calculated the consumption of tobacco foliage by larvae and reported that 5.2, 9.7, 33.5, 175.4, and 1941.4 cm2 of foliage were eaten by instars 1–5, respectively, for a total of 2165 cm2. Notice that about 90% of the foliage consumption occurred during the final instar, and though the exact value of foliage consumption would differ on tomato, the pattern would be the same. Larvae blend in with the foliage and are not easy to detect. Thus, it

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is not surprising that they often were not observed until they caused considerable damage at the end of the larval period.

Management Sampling. Moths are attracted to light and can be captured in light traps. Light traps were also used to attempt suppression of hornworm populations, and though some reduction was noted, this approach did not prove practical (Gentry et al., 1967). Isoamyl salicylate is attractive to both species of hornworms (Scott and Milam, 1943). Visual examination of foliage usually is recommended for monitoring of larval populations. Young larvae of tobacco hornworm tend to be found in the upper regions of the plant, whereas larvae of tomato hornworm tend to be lower; differential flight and oviposition behavior between the species is implicated (Yamamoto, 1972). Insecticides. Chemical insecticides or Bacillus thuringiensis are applied to the foliage for larval suppression (Creighton et al., 1961). Cultural Practices. The pupae are large and are not buried very deeply in the soil, so greater than 90% mortality is caused by normal soil tillage practices. Hand picking and destruction of larvae is often practical in the home garden. To take advantage of the preference of Polistes wasps for hornworm larvae, wasp shelters or nesting boxes were placed in tobacco fields to encourage the wasps, and wasp colonies were relocated into tobacco (Lawson et al., 1961). Although wasp predation was inadequate to prevent damage to tobacco, this approach might be satisfactory for tomato. Tobacco and tomato hornworms thrive on tobacco plants that are allowed to revegetate after harvest of the leaves, leading to high populations during the next year. Destruction of tobacco stalks, or inhibition of sprouting by application of plant growth regulators, greatly reduces hornworm populations in subsequent seasons (Rabb, 1969). Although not documented, timely destruction of tomato crop residue likely would have similar beneficial effects.

Portulaca spp. (Portulacaceae). If preferred host are eliminated, of course, larvae attempt to feed on nearly any nearby plant. Among vegetables reportedly injured are beet, cantaloupe, lettuce, tomato, turnip, and watermelon. Fruits such as apple, currant, gooseberry, grape, pear, and plum are also listed among hosts. Other plants consumed include bitter dock, Rumex obtusifolius; evening primrose, Oenothera spp.; fuchsia, Fuchsia spp.; 4 o’clock, Mirablis spp.; and willow herb, Epilobium spp. The adults have a long proboscis and take nectar from several flowering plants. Natural Enemies. Several flies are known to parasitize the larvae, including Compsilura concinnata (Meigen), Drino incompta (Wulp), Winthemia deilephilae (Osten Sacken), and W. quadripustulata (Fabricius) (all Diptera: Tachinidae). Although there is no quantitative data on incidence of parasitism, it is evident from the literature that larvae frequently are parasitized by tachinids. Many mature larvae, when collected, are observed to have tachinid eggs adhering to their bodies, usually located dorsally behind the head. Also attacking whitelined sphinx is a pupal parasitoid, Brachymeria robusta (Cresson) (Hymenoptera: Chalcididae). Life Cycle and Description. There are two generations annually, with the pupa being the overwintering stage. Adults are usually observed in June and September. Egg. The eggs are oval, and yellow-green initially, but become bluish later in development. They are deposited on host foliage. Eggs normally hatch in 6–7 days.

Whitelined Sphinx

Hyles lineata (Fabricius) (Lepidoptera: Sphingidae)

Natural History Distribution. This native insect is probably the most widely distributed sphingid moth in North America. It is found throughout the United States and southern Canada, and its range extends south into Central America and the Caribbean. Host Plants. This insect reportedly has a wide host range, but feeds principally on weeds. Its presence among garden plants often results in the assumption that it is developing at the expense of crops when it is actually grazing on an understory of competing plants, especially purslane,

FIG. 10.148  Larva of whitelined sphinx. (Photo by J. Capinera.)

Larva. There are six instars. At hatching the young larva measures about 4 mm long, and measures 6–7, 12–13, 22– 23, 37–40, and 55–60 mm long at the start of the subsequent instars, respectively. At maturity, the larva attains a length of 75–90 mm. Development time for the instars is about 8, 4, 5, 5, 4, and 11  days, respectively. During the initial five instars, larvae are green or black, with an yellowish to

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orange-brown head. The posterior end bears a “horn” that is usually black or yellow and black. A horizontal yellowish subdorsal line is present along each side. Although larvae are fairly variable in color during the early instars, variation is exceptionally marked in the terminal instar. In the sixth instar the green form has a yellowish-green body with a subdorsal row of pale spots bordered above and below with a black line, and brightly colored spots around the spiracles. The corresponding black form has a blackish body with three narrow yellow lines dorsally. Pupa. Pupation often occurs at the soil surface in a loosely constructed cocoon of brown color, but some larvae apparently pupate without forming a cocoon, and some enter the soil to pupate. The pupa is light brown, and measures about 44–48 mm long. Duration of the pupal stage is 30–40 days.

reports of this species completing its larval development on vegetable crops.

Management Moths are attracted to light, and can be captured in light traps. The presence of adults does not necessarily indicate future problems with crops, however, unless portulaca or another favored weed is also present. Larval infestation of vegetable crops can be avoided by preventing portulaca from growing amongst crop plants. Should purslane become extensively established, it should not be killed or removed if infested with larvae, as this action will force the larvae to feed on the crop.

FAMILY TORTRICIDAE—LEAFROLLER MOTHS Pea Moth

Cydia nigricana (Fabricius) (Lepidoptera: Tortricidae)

Natural History

FIG. 10.149  Whitelined sphinx moth. (Photo by J. Capinera.)

Adult. The adults are more active at dusk, but can also be observed feeding during the day, hovering at flowers while sipping nectar. The moth has a wingspan of 60–90 mm. The body is dull brown with white lines, running the length of the head and thorax. The abdomen bears white and dark brown spots dorsally. The olive front wings are marked with white-lined veins and a whitish stripe, extending from the base to the tip of the wing. The hind wings are dark brown, with a rosy band across the middle. Life history information on white-lined sphinx is quite limited. Soule (1896), Cooley (1905), and Eliot and Soule (1921) provided notes on this species.

Damage The mature larvae of white-lined sphinx are quite robust, and can consume large quantity of foliage. However, they normally limit their feeding to portulaca, moving onto crop plants only when their favored food is completely consumed, and they are faced with starvation. Much of the larva's reputation for damaging crops seems to stem from the habit of resting or nibbling on crop plants. There are no

Distribution. Pea moth is native to Europe, and was first observed in North America in 1893 at Toronto, Canada. The insect likely had been in residence for a number of years before attracting attention. It quickly spread across the northern United States, causing great damage in the Great Lakes region in the early 1920s, and reached Washington in 1926. It inflicted considerable damage in both Washington and British Columbia in 1933. Now found throughout the northern United States and southern Canada, it is rare in the Prairie Provinces, and infrequently is a serious pest anywhere in North America. It remains a serious problem in Europe, however, and is known from Japan. Host Plants. This insect feeds only on plants in the family Leguminosae. The only crop injured is pea, but several other legumes are suitable hosts, including Canada pea, Vicia cracca; common vetch, Vicia angustifolia; hairy vetch, Vicia villosa; lupines, Lupinus spp.; Scotch broom, Cytisus scoparius; spring vetch, Vicia sativa; sweet pea, Lathyrus odoratus; yellow vetchling, Lathyrus pratensis; and possibly other members of these plant genera. Hanson and Webster (1936) reported that the most suitable host was pea, followed by sweet pea, and then the vetches. Larvae feeding on lupines and Scotch broom experience high mortality. Natural Enemies. Several natural enemies have been imported from Europe and introduced into Canada and the United States. Although some species failed to establish, the parasitoids Ascogaster quadridentata Wesmael, Phanerotoma fasciata Provancher (Hymenoptera: Braconidae), and Glypta haesitator Gravenhorst (Hymenoptera: Ichneumonidae) became established. Ascogaster quadridentata proved to

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be very important, causing over 70% parasitism in British Columbia. Although the parasitoids undoubtedly are important in checking the abundance of pea moth, changes in cropping practices also are important. Specifically, the dramatic decline in the production of dried peas is of major significance, because the major host plant is now much less abundant. Also, harvesting of peas while they are green results in the destruction of larvae before most are able to mature. Clausen (1978) summarized the parasitoid introduction program. Pea moth seems to be free of important diseases. However, it has been shown to be susceptible to infection by a granulosis virus isolated from a related insect, codling moth, Cydia pomonella (Linnaeus) (Payne, 1981). Life Cycle and Description. Generally there is a single generation per year, though in Washington a few insects apparently go on to form a second generation. In England, a small proportion of the insects have been observed to have a 2-year life cycle, larvae spending two winters and a summer in the soil; this likely occurs in North America as well. Oviposition occurs in June and July, followed by development of larvae until September. Larvae overwinter, with pupation occurring in the spring. Egg. Moths become active in June, and after a preoviposition period of 8–9 days (range 5–13 days), eggs are deposited. Oviposition occurs principally on the upper half of the plant, and eggs are scattered on or near the flowers and pods. When deposited on the leaf surface, the lower surface is preferred. The egg is flattened, oval, and measures about 0.7 mm long and 0.5 mm wide. The lower theshold for development is about 8.5–9.5°C, with the upper threshold about 30°C. Initially, the eggs are whitish and somewhat translucent, but two pink streaks develop after 2–3  days. Duration of the egg stage is about 9, 6, and 3 days at 15°C, 20°C, and 25°C, respectively.

FIG. 10.150  Pea moth larva. (Drawing by USDA.)

Larva. The larva is whitish with a black head and thoracic plate during the first two instars. Afterwards it assumes a yellowish color, but retains the dark markings. The body bears short and sparse hairs. It grows from a length of about 1.5 to 12–15 mm. Duration of the larval stage, which includes five instars, is 18–35 days. The first instar is very active, searching extensively on the plant for suitable food. When the larva locates a suitable pod it burrows in, the entry hole developing into a small brown blister. All stages of pods are attacked,

except for the very old, dry pods. Usually 2–4 seeds are destroyed in each pod, though occasionally more than one larvae enter a pod, resulting in greater damage. Mature larvae drop from the plant and enter the soil, usually to a depth of 5–7 cm. They spin a thick silken cocoon measuring about 10 mm long and 4.5 mm wide, and enter diapause. The larva remains in the cocoon until spring, when they escape the old cocoon, and move closer to the soil surface. They then spin a second, much thinner cocoon, and pupate. Pupa. The pupa is dark brown and measures 7–8 mm long and about 1.7 mm wide. The dorsal surface of the abdominal segments is armed with rows of spines, structures which aid the escape of the pupa from the cocoon when the moth is about to emerge. Duration of the pupal stage is 12–14 days.

FIG. 10.151  Pea moth. (Drawing by USDA.)

Adult. The adult is a small brownish-gray moth with a wingspan of about 12 mm. The forewing is also marked with black, and spots of white, particularly along the leading edge of the wing. The hind wing is nearly uniform brownish gray, but is bordered with a light-colored fringe with a thin, dark inner line. The moths tend to be active only in the afternoon, and a threshold of about 18°C must be exceeded before flight occur. Flight also is reported to be favored by high humidity. Female moths emerge with nearly mature ovaries, mate upon emergence, and commence egg laying almost immediately. Females disperse freely in search of suitable oviposition sites, but males are less dispersive. Males are attracted to a female-produced sex pheromone that has been identified, and synthesized, for use in population monitoring. Wright and Geering (1948) provided detailed biology of pea moth, though the observations were made in England. The work of Hanson and Webster (1936) in Washington, though less complete, better reflects American and Canadian conditions. Lewis and Sturgeon (1978) gave important infomation on egg ecology.

Damage The larvae damage the pea crop by tunneling into the pod and feeding on one or more seeds in each pod. Often the larva does not consume the entire seed before attacking

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a­ nother. Silken webbing and fecal material also are found in the pea pod. In addition to yield loss caused by destruction of pea seed, and quality loss caused by contamination of undamaged seed with insect-damaged seed, additional loss can occur when the larvae contaminate peas harvested for processing (freezing or canning). The latter problem is minimal with this insect because it is not hidden within the pea seed. Damage potential varies with intended use of the peas. Pea moth damage is of little consequence for peas grown as livestock food because the quality standards are low. For processing peas, the damage potential is significant, but commercial processors have established procedures to eliminate damaged peas and insects. Pea grown for seed is another instance where damage potential is great, because even a small level of damage, 1.0%–1.5%, is sufficient to inflict loss to farmers due to downgrading of the seed quality.

Management Sampling. A sex pheromone is known for this insect (Greenway, 1984). In England, great effort has gone into refinement of a pheromone-based monitoring system to farmers of pea moth problems and to time insecticide applications carefully. At least two pheromone traps are placed in each field, and when 10 or more moths are collected in either trap for two consecutive nights the farmer can be assured that moths are present in sufficient number to warrant control measures (Gratwick, 1992). After consulting weather data and estimating the time of egg hatch, producers can apply insecticide to kill first instar larvae at hatch. Usually a delay of 10–15 days is recommended between initiation of sustained moth catches and application of insecticide to allow for oviposition and egg hatch. This system has found good acceptance in England (Wall et al., 1987; Wall, 1988). Release of large quantities of the sex pheromone can also be used to disrupt chemical communication in pea moth and disrupt mating (Bengtsson et al., 1994). However, this mating disruption system is difficult to implement because the moths are quite dispersive and gravid females can invade fields treated with pheromone (Saucke et al., 2014).

Insecticides. Insecticides are applied to foliage to kill first instar larvae before they enter the pea pod, so timing is critical. This is accomplished most easily with determinant pea varieties, because the synchrony of fruiting allows growers to protect pods with fewer applications of insecticides. With indeterminant varieties, or when planting of determinant varieties is staggered, more applications are required to protect the crop from damage. If crops remain in the field in June, they may require two insecticide treatments at about 10 day intervals to protect them adequately, though early crops may escape injury. Application of granular formulations of systemic insecticides is reported to be less satisfactory than foliar application (Thompson and Sanderson, 1977). Cultural Practices. Several cultural practices help reduce damage from pea moth (Wright and Geering, 1948). Tillage can destroy the larvae overwintering in the soil. Disking the soil twice is commonly recommended, especially if the field, or a nearby field, is to be planted to peas during the following growing season. Destruction of wild vetches is desirable to decrease the potential of pea moth to develop on these alternate hosts. Thick vegetation, either in the form of leafy pea varieties or weedy vegetation, exacerbates pea moth problems, probably by providing shelter for the moths. Thus, weeds should be destroyed and pea varieties with minimal foliage should be favored. Abundance of pea moths and associated damage are linearly related to land area devoted to pea production (Huusela-Veistola and Jauhiainen, 2006). Peas that mature early may also escape attack, so early planting or selection of early maturing varieties is desirable. Crops that are harvested before mid-June may escape attack. It is also beneficial to either pick all peas or to quickly destroy remaining pods after harvest, as this can minimize successful development of pea moth in the field. Peas that are grown for livestock food seldom receive insecticide applications, so they may serve as a source of infestation. Cultivation of fresh market and processing peas at a considerable distance away from livestock peas is suggested.

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Chapter 11

Order Orthoptera—Grasshoppers, Crickets, and Katydids

FAMILY ACRIDIDAE—GRASSHOPPERS American Grasshopper

Schistocerca americana (Drury) (Orthoptera: Acrididae)

Natural History Distribution. This grasshopper is found widely in the eastern United States, west to Iowa and Texas. Occasionally, it is reported from southern Canada and the New England states, but is likely a migrant there. Even in the midwestern states, where it is common, the resident population receives a regular infusion of dispersants from southern locations. In the southeast, it is quite common, and is one of the few grasshopper species to reach epidemic densities. The American grasshopper seems to be native to North America, but its distribution also extends south through Mexico to northern South America. Host Plants. The adult American grasshoppers tend to be arboreal in habit, and considerable feeding by adults occurs on forest, shade, and fruit trees. The nymphs, however, feed on numerous grasses and broadleaf plants, both wild and cultivated, in open fields. During periods of abundance, almost no plants are immune to attack, and vegetables, fruit, grain crops, and ornamental plants are injured by both the nymphal and adult stages. The feeding preferences of the nymphs and adults are highly correlated. The American grasshopper consumes bean, corn, okra, and yellow squash over some other vegetables when provided with choices (Capinera, 1993b). Among the vegetables less preferred are eggplant, pepper, and cucumber. Other food crops not preferred include peanut, sugarcane, tangerine, orange, and

Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00011-X © 2020 Elsevier Inc. All rights reserved.

loquat. As is normally the case with polyphagous insects, however, preferences can shift somewhat based on the availability of alternate hosts. Thus, during periods of great abundance in Florida brought on by the unusual availability of annual weeds, these grasshoppers caused considerable damage to orange, a relatively unpreferred but readily available crop plant. Natural Enemies. The natural enemies of S. americana are not well known. Birds such as mockingbirds, Mimus polyglottos polyglottos (Linnaeus), and crows, Corvus brachyrhynchos brachyrhynchos Brehm, have been observed to feed on these grasshoppers. Fly larvae, Sarcophaga sp. (Diptera: Sarcophagidae) are sometimes parasitic on overwintering adults. Life Cycle and Description. The American grasshopper has two generations per year and overwinters in the adult stage. In Florida, eggs produced by overwintered adults begin to hatch in April–May, producing spring generation adults by May–June. This spring generation produces eggs that hatch in August–September. The adults from this autumn generation survive the winter. Egg. The eggs of S. americana initially are light orange, turning tan with maturity. They are ­elongate-spherical, widest near the middle, and measure about 7.5 mm long and 2.0 mm wide. The eggs are clustered together in a whorled arrangement, and number 75–100 eggs per pod, averaging 85. They are inserted into the soil to a depth of about 4 cm and the upper portion of the oviposition hole is filled by the female with a frothy plug. The duration of the egg stage is about 14 days. The nymphs, upon hatching, dig through the froth to attain the soil surface.

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FIG. 11.2  Adult American grasshopper. (Photo by J. Capinera.)

FIG. 11.1  American grasshopper nymphs, showing solitary or low density (green) and gregarious or high-density (orange) nymphs. (Photo by J. Capinera.)

Nymph. Normally, there are six instars in this grasshopper (Kuitert and Connin, 1952; Capinera, 1993a), though Howard (1894) reported only five instars. The young grasshoppers are light green. They are extremely gregarious during the early instars. At low densities the nymphs remain green throughout their development, but at higher densities they gain increasing amounts of black, yellow, and orange coloration commencing with the third instar. Instars can be distinguished by their antennal, pronotal, and wing development. The first and second instars display little wing development but have 13 and 17 antennal segments, respectively. In the third instar, the number of antennal segments increases to 20–22, the wings begin to display weak evidence of veins, and the dorsal length of the ventral lobe of the pronotum is about 1.5 times the length of the ventral surface. Instar four is quite similar to instar three, with 22–25 antennal segments, though the ratio of the length of the dorsal to ventral surfaces of the pronotal lateral lobe is 2:1. In instar five, there are 24–25 antennal segments, and the wing tips assume a dorsal rather than ventral orientation, but the wing tip does not exceed the first abdominal segment. In the sixth instar, there are 24–26 antennal segments and the wing tips extend beyond the second abdominal segment. The overall body length is about 6–7, 12–13, 16–18, 22–25, 27– 30, and 35–45 mm for instars 1–6, respectively. The development time is about 4–6, 4–6, 4–6, 4–8, 6–8, and 9–13 days for the corresponding instars when reared at about 32°C.

Adult. The adult is rather large but slender bodied, and measures 39–52 and 48–68 mm long in the male and female, respectively. A creamy white stripe normally occurs dorsally from the front of the head to the tips of the forewings. The front wings bear dark-brown spots, the pronotum dark stripes. The hind wings are nearly colorless. The hind tibiae normally are reddish. Overall, the body color is yellowish brown or brownish with irregular lighter and darker areas, though for a week or so after attaining the adult stage a pinkish or reddish tint is evident. Adults are active, flying freely, and sometimes in swarms. They normally are found in sunny areas, but during the warmest periods of the day move to the shade. Adults are long-lived, persisting for months in the laboratory and apparently in the field as well. This can lead to early season situations where overwintered adults, all instars of nymphs, and new adults are present simultaneously. Mild winters favor survival of overwintering adults and apparently lead to population increase if preferred food supplies are also plentiful. The biology of the American grasshopper was described by Howard (1894) and Kuitert and Connin (1952). It is also included in a field guide to grasshoppers (Capinera et al., 2004).

Damage The American grasshopper is a defoliator, and eats irregular holes in leaf tissue. Under high-density conditions, it can strip vegetation of leaves, but more commonly leaves plants with a ragged appearance. Adults display a tendency to swarm, and high densities can cause severe defoliation. As the American grasshopper is a strong flier, it also sometimes becomes a contaminant of crops. When the lateseason crop of collards in the southeast is harvested mechanically, the American grasshopper may be incorporated into the processed vegetables. In the winter and early spring, a similar problem can occur with specialty lettuce if it is harvested mechanically. Although most grasshoppers can be kept from

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dispersing into crops near harvest by treating the periphery of the crop field, it is much more difficult to prevent invasion by the American grasshopper because, owing to its being a strong flier, it can fly over any such barrier treatments.

Management Sampling. Sampling methods are not well defined. Populations normally originate in weedy areas such as fence rows and abandoned fields. Thus, margins of fields are first affected and where monitoring should be concentrated. It is highly advisable to survey weedy areas in addition to crop margins if grasshoppers are found, as this gives an estimate of the potential impact if the grasshoppers disperse into the crop. Also, it is important to recognize that this species is highly dispersive in the adult stage, and can fly hundreds of meters or more to feed. Grasshoppers can be flushed from a predetermined unit of ground, such as a square meter, and counted as they hop or fly, but this only works with low vegetation such as pastures. In taller vegetation such as crops, the grasshoppers are usually sampled using a sweep net. The American grasshoppers take flight readily, so they can be difficult to sample even when using a net. Insecticides. Foliar application of insecticides can suppress grasshoppers, but they are difficult to kill, particularly as they mature. Toxic bait formulations, which often are recommended for grasshopper suppression, are not usually recommended for this species because these grasshoppers spend little time on the soil surface (where the bait is scattered), instead preferring to climb high in vegetation. Cultural Practices. Land management is an important element of S. americana population regulation. Grasshopper densities tend to increase in large patches of weedy vegetation that follow the cessation of agriculture or the initiation of pine tree plantations. In both cases, the mixture of annual and perennial forbs and grasses growing in fields that are untilled seems to favor grasshopper survival, with the grasshoppers then dispersing to adjacent fields as the most suitable plants are depleted. However, as abandoned fields convert to dense woods or the canopy of pine plantations shades the ground and suppresses weeds, the suitability of the habitat declines for these grasshoppers. The disturbance or maturation of crops may cause the American grasshopper to disperse, sometimes over long distances, into crop fields. Therefore, care should be taken not to cut vegetation or till the soil of fields harboring grasshoppers if a susceptible crop is nearby. Planting crops in large blocks decreases the relative amount of crop edge, and the probability that a crop plant within the field will be attacked. Biological Control. Products for grasshopper control that display minimal toxicity to mammals and other nontarget organisms are always desirable. Insecticidal products derived from plants and microorganisms tend to offer the best

potential in this regard (Foster et al., 1996; Sieglaff et al., 1997, 1998; Amarasekare and Edelson, 2004; Demirel and Cranshaw, 2006d; Capinera and Froeba, 2007; SandovalMojica and Capinera, 2011). The American grasshoppers display a reluctance to feed on plants treated with neem (azadirachtin, neem oil) products, which are extracted from the neem tree (Azadirachta indica), because neem can also be insecticidal. However, neem does not persist when exposed to sunlight, so the benefits may be transient. The fungus Beauveria bassiana strain GHA has been shown to have insecticidal effects on grasshoppers, as has spinosad, a naturally derived insecticide produced by fermentation of the actinomycete bacterium Saccharopolyspora spinosa. Beauveria bassiana can be effective by contact and ingestion, whereas spinosad works better by ingestion. Like many fungi, B. bassiana is inactivated by high temperatures. These products do not cause rapid mortality, but after 2–4 days, mortality may be discernable. The fungus Metarhizium flavoviride is used in other areas of the world for grasshopper and locust suppression. By mixing this fungus with oil, the fungus survives even in hot, dry climates. Experimentally, it has been demonstrated that the American grasshopper is very susceptible to M. flavoviride (Sieglaff et al., 1997, 1998), but products containing this fungus are not available in the United States.

Differential Grasshopper

Melanoplus differentialis (Thomas) (Orthoptera: Acrididae)

Natural History Distribution. This native grasshopper occurs widely in the central and western regions of the United States, and in northern Mexico; in Canada, it occurs in southern Saskatchewan and British Columbia. Within the United States, it is absent from the Atlantic and Gulf Coast region, except that it occurs in the Pennsylvania, New Jersey, and Maryland areas. It is infrequent in the Pacific Northwest area. Also, within the large geographic area generally inhabited by the differential grasshopper, it is rare in arid environments. Host Plants. The host plants preferred by the differential grasshopper are tall broadleaf plants such as those typically associated with fence rows, irrigation ditches, and fallow fields. It prefers plants in the family Compositae such as ragweed, Ambrosia spp.; sow thistle, Sonchus asper; sunflower, Helianthus annuus; and prickly lettuce, Lactuca scariola; though it will feed on other broadleaf plants such as kochia, Kochia scoparia; and smartweed, Polygonum sp.; and on such grasses as bermudagrass, Cynodon dactylon; slender oat, Avena barbata; barley, Hordeum sp.; and Johnsongrass, Sorghum ­halepense. In North Dakota alfalfa fields, the differential grasshopper reportedly ate kochia, Kochia scoparia; quack grass,

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Agropyron repens; squirreltail grass, Hordeum jubatum; bristly foxtail, Setaria spp.; and field bindweed, Convolvulus repens, in addition to alfalfa (Mulkern et  al., 1962). On the prairie, they ate mostly stickseed, Lappula echinata; wavyleaf thistle, Cirsium undulatum; quack grass, Agropyron repens; and pepperweed, Lepidium densiflorum (Mulkern et al., 1964). Crops sometimes injured include alfalfa, clover, corn, cotton, soybean, sugar beet, timothy, and small grains such as barley and wheat. The differential grasshopper is not normally an important vegetable pest. It occurs among vegetables if weeds are present within, or adjacent to, crops. However, during periods of great abundance all vegetables are at risk, because under such conditions virtually all green vegetation may be consumed. Natural Enemies. The natural enemies of the cropfeeding Melanoplus spp. are quite similar. For information on natural enemies of the differential grasshopper, see the section on natural enemies of the migratory grasshopper, Melanoplus sanguinipes (Fabricius). Weather. Weather affects the distribution of the differential grasshopper, but in a manner somewhat different from some other grasshopper species. The differential grasshopper is associated with dense vegetation, so it follows that it would thrive in areas with adequate moisture to support the lush growth of plants. It is a common and damaging species on the eastern edge of the Great Plains, where rainfall is plentiful, and is relatively infrequent along the drier western edge of this region. Wakeland (1961) documented the expansion of the differential grasshopper populations into areas of the northern Great Plains dominated by the migratory grasshopper, Melanoplus sanguinipes (Fabricius), a species that is better adapted to dry conditions. The migratory grasshopper was supplanted by the differential grasshopper as longterm mean precipitation levels increased and temperatures decreased in this area. When the weather returned to normal, however, the migratory grasshopper resumed its status as the dominant species. The irrigated cropland is also favored by the differential grasshopper, allowing this species to become abundant in dry areas due to “artificial rainfall.” High levels of precipitation are not entirely advantageous for the differential grasshopper. Precipitation during the warm months leads to the outbreak of disease in the differential grasshopper populations. This is a short-term response, and disease outbreaks occur only when grasshoppers are abundant. The differential grasshopper seems to be more susceptible to disease than some other species, including the migratory grasshopper (Wakeland, 1961), or perhaps it is an artifact of the more moist environments favored by the differential grasshopper. Precipitation accompanied by cool weather during the hatching period is also detrimental to the differential grasshopper, as with all grasshoppers, largely because it disrupts feeding during the critical early life of the grasshopper. The late onset of winter can favor grasshopper population increase, because it allows adults additional time to produce eggs.

Life Cycle and Description. A single generation occurs annually, with the egg stage overwintering. This is a late-season species, with eggs hatching about 3 weeks after those of the twostriped grasshopper, Melanoplus bivittatus (Say), and 2 weeks after those of Melanoplus sanguinipes (Fabricius). In Colorado, eggs hatch in June, usually within a 2-week period. Nymphs complete their development in July–August; adults are present from August to October. Egg. The eggs are creamy white, yellowish, or light brown. They are elongate-cylindrical, and taper to a blunt point at each end. The eggs measure about 4–5 mm long and 0.85 mm in diameter. They are clustered within an elongate, cylindrical pod consisting of about 40–200 eggs arranged in four columns and held together by frothy material. They are deposited in the soil, and the upper portion of the pod is plugged with additional froth. They are normally deposited among the roots of grasses and weeds, especially along edges of fields, and in moist soil. Embryonic development occurs in the autumn after eggs are deposited, but the embryo enters diapause at about the point of 50% development, and must endure a period of cold before commencing growth. Nymph. The nymphs normally develop during the warmest period of the summer, and complete their development in about 30 days, though development times are extended by cool weather. There are six instars, but the nymphs are not distinctive in appearance. The first instars are greenish, yellowish, or brownish with an indication of a black stripe on the outer face of the hind femora. Instars 2–6 are similar but possess a curved dark stripe extending from the back of the eye across the pronotum, and are bordered below by a narrower white stripe. The black stripes on the femora are pronounced. The hind tibiae are light green or gray. The nymphs increase in body length from 5.3 to 6 mm in the first instar, to 5.2–6.8, 9.4–12.6, 12–14, 18–21.5, and 22–32 mm in instars 2–6, respectively. The number of antennal segments is 12, 14–17, 19–20, 21–22, 25–26, and 26 in instars 1–6, respectively.

FIG. 11.3  Yellow adult of the differential grasshopper. (Photo by J. Capinera.)

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increased by the abundance of weeds associated with crops, and the irrigation practices of western farms. It also readily exploits disturbed sites in cities and towns. Unlike some of the arid environment-loving species, its numbers and damage may increase following long-term increases in precipitation. The damage caused by the differential grasshopper principally takes the form of leaf removal. Plants may be completely defoliated, or left ragged. As this grasshopper tends to roost in elevated locations at night, where they may nibble while resting, trees and shrubs outside the normal dietary range are sometimes severely injured.

Management FIG.  11.4  Blackish adult of the differential grasshopper. (Photo by J. Capinera.)

Adult. The adults are large grasshoppers, the males ­ easuring 28–34 mm long and females 32–44 mm. They m display more color variability than most grasshoppers. They may be yellowish, principally brownish-green, or almost entirely black, though the yellow form is most abundant. The most distinctive feature of these grasshoppers is the row of black marks, arranged in herringbone fashion on the outer face of the hind femora. These markings are not so evident in the infrequent black form, which instead bears four white blotches on its otherwise black hind femora. The front wings in all except the black form are uniform grayish or brownish; the hind wings in all forms are colorless. The males bear large, boot-shaped cerci. Like many other grasshoppers, the differential grasshopper tends to roost on elevated locations at night. This allows them to bask in the morning sun, and to assume activity early in the day. Bushes and other tall vegetation are favorite perches. Aspects of the biology of the differential grasshopper were treated by many authors, including Parker and Shotwell (1932), Parker (1939), and Shotwell (1941). Kaufmann (1968) gave some interesting biological information, but due to the low rearing temperature the relevance of this study to field biology is questionable. An excellent synopsis was presented by Pfadt (1994e), who also pictured all stages of development. Melanoplus differentialis is included in many grasshopper keys, including those by Blatchley (1920), Dakin and Hays (1970), Helfer (1972), Capinera and Sechrist (1982), and Richman et  al. (1993), and also in a field guide to grasshoppers (Capinera et  al., 2004). This species was also included in a key to grasshopper eggs by Onsager and Mulkern (1963). Rearing of Melanoplus spp. was described by Henry (1985).

Damage This species seemingly has benefited from agricultural practices more than most grasshoppers, with grasshopper s­ urvival

Management of the various Melanoplus spp. grasshoppers is substantially the same. For information on differential grasshopper management, see the section on management under the migratory grasshopper, Melanoplus sanguinipes (Fabricius).

Eastern Lubber Grasshopper

Romalea microptera (Beauvois) (Orthoptera: Acrididae)

Natural History Distribution. This native grasshopper is common in the southeastern states from North Carolina to eastern Texas, including the entire peninsula of Florida. It also sometimes is known as R. guttata (Houttuyn). Host Plants. The eastern lubber grasshopper has a broad host range. Jones et al. (1987) indicated that at least 26 species from 15 plant families containing shrubs, herbs, broadleaf weeds, and grasses are eaten. Watson (1941) indicated its preference for pokeweed, Phytolaca americana; treadsoftly, Cnidoscolus stimmulosus; pickerelweed, Pontederia cordata; lizard’s tail, Saururus sp.; sedge, Cyperus; and arrowhead, Sagittaria spp. Although their preferred habitat seems to be low, wet areas in pastures and woods and along ditches, lubbers disperse long distances during the nymphal period. Lubbers are gregarious and flightless; their migrations sometimes bring large numbers into contact with crops where they damage vegetables, fruit trees, and ornamental plants. Although lubbers will consume a large number of vegetable plants, in choice tests they favor plants in the families Liliaceae, Cucurbitaceae, Asteraceae, and Brassicaceae (Cruciferae). Watson and Bratley (1940) indicated their preference for corn silk, cowpea, and peanut under field conditions. Also, they seek out and defoliate amaryllis, Amazon lily, crinum, narcissus, and related plants in flower gardens. In Florida, lubbers sometimes damage citrus trees. Although they cannot fly, they readily climb trees. Capinera (2014) assessed the relative preference of lubber grasshoppers for 104 plants by comparing the consumption of each to romaine lettuce (a favored host). Only a few

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plants (3%) were more preferred than romaine, but 20% were consumed as readily as romaine, confirming a broad host range (polyphagy) in this species. The most preferred species were oleander, Nerium oleander; painted leaf, Poinsettia cyathophora; and wild taro, Colocasia esculenta. Interestingly, all of these plants are considered to be toxic to vertebrates. This research also included a study of choice and no-choice (starvation) tests, and found a statistically significant correlation between the feeding behaviors in the two types of tests, so each should have a good degree of predictability. Natural Enemies. The natural enemies of lubber grasshoppers are not well documented. Vertebrate predators such as birds and lizards learn to avoid these insects due to the production of toxic secretions. Naive vertebrates gag, regurgitate, and sometimes die following the consumption of lubbers. However, loggerhead shrikes, Lanius ludovicianus Linnaeus, capture and cache lubbers by impaling them on thorns and the barbs of barbed wire fences. After 1–2 days, the toxins degrade and the dead lubbers become edible to the shrikes (Yosef and Whitman, 1992). The parasitic flies Anisia serotina (Reinhard) (Diptera: Tachinidae) (Lamb et al., 1999), Blaesoxipha opifera (Coquillett) and B. hunteri (Hough) (Diptera: Sarcophagidae) (Capinera, unpublished) and nematodes have been reported from lubbers, and it is possible to infect lubbers experimentally with the ­grasshopper-infecting nematode Mermis nigrescens. Pathogens known from eastern lubber include the gregarine Boliviana floridensis (Johny and Whitman, 2005) and the fungus (microsporidian) Encephalitozoon romalea (Lange et al., 2009). Life Cycle and Description. There is a single generation annually, with the egg stage overwintering. There is not an obligatory diapause in this species. These grasshoppers are long-lived, and either nymphs or adults are present throughout most of the year in the southern portions of Florida. In northern Florida and along the Gulf Coast, they may be found from February–March to about October–November. Egg. The eggs of lubber grasshoppers are yellowish or brown. They are elongate-elliptical and measure about 9.5 mm long and 2.5 mm wide. They are laid in neatly arranged clusters, or pods, which consist of rows of eggs positioned parallel to one another and held together by a secretion. Normally, there are 30–80 eggs in each pod, and one to five egg pods per female are reported. Watson (1941) reported that ovipositing females preferred mixed broadleaf tree-pine habitats with intermediate soil moisture levels. He indicated that they avoided both lowland moist, compact soil and upland dry, sandy soil. The female deposits the pod in the soil at a depth of 3–5 cm and closes the oviposition hole with a frothy secretion or plug. The plug allows the young grasshoppers easy access to the soil surface when they hatch. The duration of the egg stage is 6–8 months, with eggs typically hatching in the morning.

FIG. 11.5  Nymph of the eastern lubber grasshopper; at this stage they are mostly black in color. (Photo by J. Capinera.)

Nymph. Young nymphs are highly gregarious, and remain gregarious through most of the nymphal period, though the intensity dissipates with time. Normally, there are five instars, though occasionally six instars occur. The nymphs are mostly black with a narrow median yellow stripe along the pronotum and abdomen, the edges of the pronotum, and on the lower side of the abdomen. The legs are well-marked with red. Their color pattern is distinctly different from the adult stage, and so the nymphs commonly are mistaken for a different species than the adult form. The early instars can be distinguished by a combination of body size, the number of antennal segments, and the form of the developing wings. The nymphs measure about 10–12, 16–20, 22–25, 30–40, and 35–45 mm long during instars 1–5, respectively. Antennal segments, which can be difficult to distinguish even with magnification, number 12, 14–16, 16–18, 20, and 20 per antenna during instars 1–5, respectively. The shape of the wing pads immediately behind the pronotum changes slightly with each molt. During the first instar, the ventral surface is broadly rounded; during the second instar, the ventral edges begin to narrow and point slightly posteriorly, and also acquire a slight indication of wing veins; during the third instar, the ventral edges of the wing pads are markedly elongate, point strongly posteriorly, and the veins are pronounced. At the molt to the fourth instar, the orientation of the small, developing wings shifts from pointing downward to upward and posteriorly. In instar four, the small front and hind wings are discrete and do not overlap, though the front wings may be completely or partly hidden beneath the pronotum. In instar five, the slightly larger wings overlap, appearing as a single pair of wings. Nymphs can complete instars 1–4 in about 7 days each, with the terminal instar requiring 10 days. However, under cool conditions, 60 days are required for nymphal development.

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be an antipredator defense, consisting of chemicals both synthesized and sequestered from the diet. Vertebrate, but not invertebrate, predators are affected (Whitman et  al., 1991, 1992). The life history and ecology of the eastern lubber grasshopper are not well documented. Rehn and Grant (1961) provided important descriptive notes on field biology and taxonomic information, and Hunter-Jones (1967) provided the basic life history information under laboratory conditions. Capinera and Scherer (2016) provide a recent summary of its biology. This species is included in a field guide to grasshoppers (Capinera et al., 2004).

Damage FIG.  11.6  A light-colored version of the eastern lubber grasshopper. (Photo by J. Capinera.)

Lubber grasshoppers are defoliators, consuming the leaf tissue of numerous plants. They climb readily, and as they are gregarious they can completely strip foliage from plants. More commonly, however, they eat irregular holes in vegetation and then move on to another leaf or plant.

Management

FIG. 11.7  A dark-colored version of the eastern lubber grasshopper; intermediate color patterns also occur. (Photo by J. Capinera.)

Adult. Adults are colorful, but their color pattern varies. In strong contrast to the nymph, the adult eastern lubber normally is mostly yellow or tawny, with black on the distal portion of the antennae, on the pronotum, and on the abdominal segments. The front wings extend two-thirds to three-fourths the length of the abdomen. The hind wings are short and are incapable of providing lift for flight. The front wings tend to be pink or rose centrally, whereas the hind wings are entirely rose. Darker forms of this species also exist, wherein the yellow color becomes the minor rather than the major color component, and in northern Florida a predominantly black form is sometimes found. Adults attain a large size, males measuring 43–55 mm long and females often measuring 50–70 mm, sometimes 90 mm. Both sexes stridulate by rubbing the forewing against the hind wing. When alarmed, lubbers spread their wings, hiss, and secrete foul-smelling froth from their spiracles. They can expel a fine spray of toxic chemicals to a distance of 15 cm. The chemical discharge from the tracheal system is b­ elieved to

Sampling. Quantitative estimates of grasshopper abundance in pastures or low vegetation are normally determined visually. This can be accomplished by estimating the boundaries of a unit of land (typically 1 m2 or yard), then flushing the grasshoppers from the sample unit and counting them as they leap or fly. A variation on this procedure is to preposition frames that outline the unit, allow the grasshoppers to redistribute, and then flush them. The latter approach is more accurate, though more time consuming. A third manner of sampling, which is more applicable to taller vegetation, is to sweep the vegetation with a sweep net. This will provide relative estimates of abundance, though the number captured is dependent on the nature of the vegetation as well as the abundance of the insects. Insecticides. Management practices for the lubber grasshopper are not well developed. Insecticides applied directly to the grasshopper can be lethal. However, due to their large size they often prove difficult to kill, and sometimes they recover after being temporarily paralyzed by insecticide application, suggesting an effective detoxification mechanism. Insecticide treatment is more effective for young grasshoppers. Because they are dispersive and may continue to invade an area even after it is treated with insecticide, it is difficult to provide protection to plants. A good practice is to identify the source of grasshoppers before they invade crop fields, and to treat these potential sources before the invasion occurs, while they are in a more juvenile (and more susceptible) stage of development. Pyrethroid insecticides have become popular for grasshopper control, but unless lubbers are sprayed directly the results often are disappointing.

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Toxic bait formulations are commonly used for grasshopper suppression, and Barbara and Capinera (2003) assessed the efficacy of several bait components for control of lubbers. They formulated several grain-based baits with several vegetable oils and established that wheat bran plus corn oil was optimal among the materials tested. With the addition of carbaryl insecticide, even consumption of a single flake of bran could be lethal. The bait was readily consumed in the presence of less preferred host plants, including some vegetables (tomato, pepper, eggplant, leek, parsley, and fennel) and ornamentals (daylily, lily of the Nile, canna lily). However, in the presence of preferred plants, including some vegetables (butter crunch lettuce, carrot foliage, yellow squash, cauliflower, collards, green onion, chive, cucumber, cabbage, cantaloupe, endive, and red leaf lettuce) and ornamentals (society garlic, caladium, and amaryllis), the bait was less readily consumed. On average, plants in the families Solanaceae and Apiaceae (less preferred plants) induced high levels of bait consumption, whereas plants in the families Liliaceae, Cucurbitaceae, Asteraceae, and Brassicaceae (preferred plants) interfered with bait consumption. These authors suggested that in ­lubber-infested areas, the garden plan be modified to have the perimeter dedicated to less preferred plants, with toxic bait applied to these areas, where the toxicant would more likely be consumed. The center of the garden, of course, would contain more susceptible plants, but hopefully the insects would not gain access because they would be poisoned following the consumption of bait at the perimeter. Similarly, Capinera (2014) evaluated a carbaryl-­ containing commercial bait formulation designed for mole cricket control, evaluating lubber mortality in cages on bare soil or including either preferred (butterfly weed, Asclepias tuberosa; Mexican petunia, Ruellia simplex) or unpreferred (bush daisy, Euryops pectinatus; penta, Pentas sp.) plants. As had been reported by Barbara and Capinera (2003), the preferred plants interfered with bait consumption relative to the unpreferred plant species (and bare soil). Biological Control. Products for grasshopper control that display minimal toxicity to mammals and other nontarget organisms are always desirable. Insecticidal products derived from plants and microorganisms tend to offer the best potential in this regard (Foster et al., 1996; Sieglaff et al., 1997, 1998; Amarasekare and Edelson, 2004; Demirel and Cranshaw, 2006d; Capinera and Froeba, 2007; Sandoval-Mojica and Capinera, 2011). Grasshoppers display a reluctance to feed on plants treated with neem (azadirachtin, neem oil) products, which are botanically derived, because neem can also be insecticidal. However, neem does not persist in sunlight, so the benefits can be transient. The fungus Beauveria bassiana strain GHA has been shown to have insecticidal effects on grasshoppers, as has spinosad, a naturally derived insecticide produced by fermentation of the actinomycete bacterium Saccharopolyspora spinosa. Beauveria bassiana can be

e­ ffective by ­contact and ingestion, whereas spinosad works better by ingestion. Like many fungi, B. bassiana is inactivated by high temperatures. These products do not cause rapid mortality, but after 2–4 days, mortality may be discernable.

Migratory Grasshopper

Melanoplus sanguinipes (Fabricius) (Orthoptera: Acrididae)

Natural History Distribution. This native grasshopper is extremely adaptable. It is found in every state in the continental United States, and in every province in Canada. It is absent from only the northernmost, coldest regions of Canada, and from southern Florida and Texas. Host Plants. This species feeds on a wide range of food plants and occurs in numerous habitats. Relative to the other common crop-feeding Melanoplus spp., the migratory grasshopper is more tolerant of arid, shortgrass environments. It tends to prefer annual broadleaf plants, but also eats grasses. Dry plant material is an important element of the diet in addition to succulent leaf tissue (McKinlay, 1981). Many authors have noted that the population abundance of the migratory grasshopper is correlated with the availability of annual broadleaf plants. Among the preferred plants are dandelion, Taraxacum officinale; stinkweed, Thlaspi arvense; Johnsongrass, Sorghum halepense; Kentucky bluegrass, Poa pratensis; shepherd’s purse, Capsella bursapastoris; pepperweed, Lepidium spp.; tansy mustard, Descurainia sophia; western wheatgrass, Agropyron smithii; winter mustard, Sisymbrium irio; young Russian thistle, Salsola kali; and young rabbitbrush, Chrysothamnus spp. (Pfadt, 1949; Scharff, 1954). Among the preferred weeds eaten in North Dakota alfalfa fields were awnless bromegrass, Bromus inermis; kochia, Kochia scoparia; field sow thistle, Sonchus arvensis; field bindweed, Convolvulus arvensis; and Russian thistle, Salsola kali (Mulkern et  al., 1962). On the prairie, however, the preferred host plants were Kentucky bluegrass, Poa pratensis; leadplant, Amorpha canescens; white sage, Artemisia ludoviciana; and western ragweed, Ambrosia psilostachya (Mulkern et al., 1964). The migratory grasshopper does not normally infest vegetable crops, but prefers to inhabit weedy areas along fences, irrigation ditches, roadsides, and in pastures. However, as favored food plants become overmature, desiccated, or depleted, grasshoppers move into vegetable crops. This is especially likely during periods when grasshoppers are extremely abundant. Among the vegetable crops reported to be injured by the migratory grasshopper are asparagus, bean, cabbage, carrot, cauliflower, celery, cucumber, lettuce, melon, onion, pea, radish, squash, tomato, and watermelon. Field crops are more often injured, particularly

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alfalfa, barley, corn, oat, and wheat. However, as happens with vegetables, when grasshoppers are numerous, buckwheat, clover, flax, millet, rye, young sorghum, soybean, sugar beet, timothy, and tobacco may be damaged. Even fruits such as apple, cherry, currant, grape, peach, plum, and strawberry, as well as numerous flowers and shrubs are attacked during periods of abundance. Pfadt (1949) studied host preferences and nymphal survival on many rangeland grasses, weeds, and some field crops. Not surprisingly, there was a strong positive relationship between preference and survival. Among the plants most suitable for nymphal survival were wheat, sunflower, alfalfa, corn, and barley, accounting for the reputation of this species as a severe pest in central and western North America. (Alfalfa is an unusual host, however, because though it is quite suitable for large nymphs, it is but a poor source of food for the youngest of the species.) Several broadleaf weeds including dandelion; downy chess, Bromus tectorum; tumble mustard, Sisymbrium altissimum; slimleaf scurf pea, Psoralea tenuiflora; and prickly lettuce, Lactuca scariola were quite satisfactory for survival, though not as suitable as the crop plants. Among the least suitable plants were common prairie grasses. Diet also affected fecundity, with favored food such as dandelion resulting in the production of a mean value of 3.5 egg pods per female during a 3-week period, whereas grasshoppers fed a mixture of prairie grasses produced only 0.3 pods per female. Natural Enemies. Many insects parasitize or prey on Melanoplus grasshoppers. The most comprehensive listing of arthropod natural enemies was published by Rees (1973); this publication also contains keys to many of the important species. The most important parasitoids are nymph- and adultattacking flies (Diptera) in the families Anthomyiidae, Nemestrinidae, Sarcophagidae, and Tachinidae, though egg parasitoids (Hymenoptera: Scelionidae) also cause mortality in grasshopper populations. In Oregon, over 70% of migratory grasshoppers were parasitized by the nemestrinid, Neorhynchocephalus sackenii (Williston), resulting in decreased longevity and reproduction. Other Melanoplus spp. are also affected by this fly, though grasshopper populations on rangeland, not cropland, are usually affected (Prescott, 1960). In a study of migratory grasshopper parasitism conducted in Ontario, the incidence of parasitism reached about 7% by the end of September, with Blaesoxipha hunteri (Hough) and B. atlantis (Aldrich) (both Diptera: Sarcophagidae) the most effective parasitoids (Smith, 1965). Sanchez and Onsager (1994), using more accurate methods to estimate parasitism of the migratory grasshopper, reported generation parasitism levels of 15%–41% in Montana, with anthomyiids and sarcophagids accounting for 50% and 35% of the parasitism, respectively. Among the most important predators are sphecid wasps (Hymenoptera: Sphecidae). Adult sphecids capture and paralyze nymphal and adult grasshoppers, bury them within

cells in the soil, and deposit an egg on the surface of the grasshopper. Upon hatching, the larva devours the paralyzed grasshopper. Predatory beetles (Coleoptera) attack the egg, nymphal, and adult stages of grasshoppers, and include ground beetles (Carabidae), tiger beetles (Cicindelidae), soldier beetles (Cantharidae), and blister beetles (Meloidae). Blister beetles are most important, though, because the grasshopper egg pod is the stage destroyed and the predatory activities are hidden below-ground, their effect is often not appreciated. Parker and Wakeland (1957) summarized the results of several studies on egg pod predation in western states; during the period 1938–40, for example, an average of 8.8% of egg pods were destroyed by blister beetles. Flies are also important predators, particularly robber (Asilidae) and bee flies (Bombyliidae). Robber fly larvae and Gryllus spp. field crickets (Orthoptera: Gryllidae) occasionally attack egg pods, and robber fly adults routinely attack nymphs and adults of grasshoppers, though other insects are also taken. Robber flies undoubtedly are important predators under rangeland conditions, but predation rates of grasshoppers in cropping systems have not been determined. Also, the propensity of robber flies to capture other predators such as sarcophagids significantly decreases their value (Rees and Onsager, 1982). Bee fly larvae are predatory on grasshopper eggs and on other insects. In western studies, an average of 6.2% of egg pods were destroyed by bee fly larvae (Parker and Wakeland, 1957). Birds are known to be important predators of grasshoppers. They are among the most important sources of food for many avian species due to their large size and abundance. The great abundance of grasshoppers in the spring coincides with the period when most birds are nesting. Birds forage freely on them in open areas such as grasslands, with some species consuming 65–150 grasshoppers per day (McEwen, 1987). Although avian predators significantly decrease the abundance of grasshoppers in grasslands, it is less certain that they forage freely in crops. Microbial pathogens can be quite important mortality agents, especially when weather conditions are suboptimal for grasshoppers, or when grasshoppers are very abundant. The principal microbial pathogens of grasshoppers are fungi, viruses, protozoans, and nematodes and nematomorphs. Several fungi (microsporidia) affect the migratory grasshopper. Species of Antonospora (Nosema) are most common, and A. (N.) locustae has been developed as a microbial insecticide. Antonospora (Nosema) spp. affect feeding, development, reproduction, and survival, and are transmitted by ingestion (Johnson, 1997). However, they infrequently appear at high levels in natural populations. Slamovits et al. (2004) suggested that this species be moved to the genus Antonospora. In addition, the fungus Entomophaga grylli causes “summit disease,” a behavior wherein grasshoppers ascend vegetation, cling to the uppermost point, and perish. In

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some areas, particularly near the bodies of water, many dead grasshoppers can be found attached to plants. Melanoplus spp. are susceptible to one pathotype of the fungus, and significant grasshopper population decreases have been linked to the incidence of this fungal disease. Infection normally occurs when nymphs contact spores that are sheltered in the soil. Spores produced in grasshoppers dying due to this disease remain in cadavers or soil for protracted periods of time. Attempts to manipulate this pathogen have met with mixed results (Carruthers et al., 1997). This is the only common naturally occurring fungus of grasshoppers. Several viruses called entomopoxviruses affect grasshoppers. One such virus, Melanoplus sanguinipes entomopoxvirus, affects the crop-feeding Melanoplus spp. and American grasshopper, Schistocerca americana (Drury). The virus disease spreads naturally by cadaver feeding. Infected grasshoppers are pale colored and lethargic, have prolonged developmental periods, and often perish. These diseases are quite rare in the field (Streett et al., 1997). Several types of protozoa are associated with grasshoppers, including amoebae, eugregarines, and neogregarines, but their impact under field conditions is not well known. When cultured at high densities in the laboratory, however, they can be important pathogens. Nematodes are important mortality factors of grasshoppers in South America, New Zealand, and Australia, but in North America only Mermis nigrescens, Agamermis decaudata, and Hexamermis spp. affect them. Mermis nigrescens is most important, and is unique in that nematodes crawl from the soil onto vegetation to deposit desiccation-resistant eggs. The eggs hatch when consumed by grasshoppers, and the resulting larvae kill the grasshoppers, return to the soil, and continue the life cycle. These nematodes apparently do not thrive in arid areas, as the adults only emerge during wet periods, and are more common in irrigated croplands than dry rangeland (Capinera, 1987). Sometimes they parasitize up to 70% of grasshoppers in an area. Nematomorphs, commonly called horsehair worms, resemble nematodes but tend to be much longer. They are rare, possibly because part of their life cycle must occur in water, but they attract considerable attention because of their large size (Baker and Capinera, 1997). Weather. Migratory grasshopper is greatly influenced by weather. Through most of its range longevity and reproduction are limited by shortage of warm weather. Thus, abnormally warm and dry periods of about 3 years stimulate increase in their numbers. Warm weather during spring and autumn is particularly important. Cool and cloudy weather in the spring inhibits feeding by young nymphs, and results in high mortality. Also, adults have the potential to be long lived and highly fecund, but their reproductive effort is normally terminated prematurely by the onset of cold weather. When summers are hot or prolonged, the development proceeds faster or longer, resulting in greater egg production. In southern areas, ­grasshoppers are less limited

by shortage of warm weather, but are more affected by shortages of food. Therefore, the occurrence of precipitation early in the season to provide luxurious foliage, especially broadleaf weed vegetation, is an important prerequisite for population increase (Capinera and Horton, 1989). Life Cycle and Description. In most areas of its range, migratory grasshopper produces a single generation, and overwinters in the egg stage. However, in southern portions of its range, two generations may occur annually. Eggs hatch relatively early, usually beginning in early June but about a week after twostriped grasshopper, Melanoplus bivittatus, begins to hatch. Hatching is protracted and may require up to 6 weeks in an area, resulting in asynchronous development of the population. Early hatching individuals mature early in the summer and have adequate time for reproduction, whereas l­ate-hatching individuals are handicapped by the onset of cold weather. Egg. Eggs are yellowish, elongate-elliptical, and slightly curved. One side convex and the opposite side concave, causing the egg to resemble a banana. They measure about 4.5 mm long and 1.2 mm wide. They are arranged in two columns within a frothy egg pod. Egg pods contain 18–25 eggs per pod, and are buried to a depth of about 35 mm. The pods are curved, about 25 mm long, and 3–4 mm in diameter. The upper portion contains only froth, allowing ready escape of the nymphs when they hatch. Females can produce 7–10 pods if fed high-quality diets; over 20 per female is recorded, usually at 2–3-day intervals. Thus, fecundity of over 300 eggs per female is possible. Natural habitats vary greatly in suitability, however, and reproductive potential is sometimes affected. In Montana, for example, Sanchez and Onsager (1988) estimated that females were able to produce only three to four egg pods annually during the 2 years of their study. Egg pods are inserted into the soil among the roots of plants. Migratory grasshopper is more likely to deposit pods within crop fields, particularly among wheat stubble, than other common crop-feeding Melanoplus spp. Under normal weather conditions about 80% of embryonic development occurs in the autumn before the onset of diapause, with the remainder of development in the spring after a period of cold weather terminates egg diapause. Nymph. Nymphs can develop over a range of about 22– 37°C, but early instars suffer high mortality at both extremes in temperature. Nymphal development requires about 42, 27, and 23 days when cultured at a constant temperature of 27, 32, and 37°C. Diet also affects the development rate. When reared at 30°C, favorable plants like tansymustard allow complete nymphal development in about 29 days, whereas nymphs developing on less suitable plants like western wheatgrass require about 42 days. Most field observations suggest that nymphal development requires 35–45 days. Normally, there are five instars, though if migratory grasshoppers are cultured at low temperature six instars is most common.

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The nymphs are tan or gray, occasionally greenish, throughout nymphal development. They bear a curved black stripe that extends from behind each eye across the pronotum. The lower edge of the stripe is bordered in white. The outer face of the hind femur is marked with an interrupted black stripe. Body length in instars 1–5 is 4–6, 6–8, 8–11, 11–16, and 16–23 mm, respectively. The number of antennal segments is 12–13, 15–17, 18–20, 21–22, and 22–24 in the corresponding instars. A detailed description of the nymphal stages is found in Shotwell (1930).

than the air temperature. However, they also tend to roost on tall vegetation at night, as this allows them to be warmed by sunlight early in the morning, thus extending their period of activity. Because of its importance as a field crop pest over wide areas of central North America, there is an extensive literature on M. sanguinipes. Important aspects of biology were given by Parker (1930, 1939), Shotwell (1941), Pfadt (1949), Parker et  al. (1955), Pfadt and Smith (1972), Onsager and Hewitt (1982), and many others. A very good synopsis was presented by Pfadt (1994b), who also pictured all stages of development. Melanoplus sanguinipes was included in many grasshopper keys, including those by Blatchley (1920), Dakin and Hays (1970), Helfer (1972), Capinera and Sechrist (1982), and Richman et al. (1993). Melanoplus sanguinipes was also included in a key to grasshopper eggs by Onsager and Mulkern (1963). A synopsis of migratory grasshopper, including keys to related Canadian Orthoptera, was given by Vickery and Kevan (1985). This species is included in a field guide to grasshoppers (Capinera et al., 2004). Rearing of Melanoplus species was described by Henry (1985).

Damage FIG. 11.8  Migratory grasshopper. (Photo by J. Capinera.)

Adult. The adult is a medium-sized species, measuring 20–26 mm long in males and 20–29 mm in females. They are grayish brown, and often tinged with reddish brown. A broad black stripe extends back from the eye and about twothirds of the length of the pronotum. The front wings are grayish brown or brown, usually with a row of brown spots centrally. The hind wings are colorless. The hind femora are not distinctly marked. The hind tibiae are greenish blue or red. The cerci of males are broad and flat, and turn dorsally at the tip. The subgenital plate is elongate, and bears a notch and grove apically. Females have a preoviposition period of 2–4 weeks. Adults normally live 60–90 days, though with good food and weather, and living under low-density conditions, longevity may be extended considerably. They can mate repeatedly. Nymphs and adults are affected by daily change in temperature. Activity levels at the soil surface, including feeding, are at their maximum when the air temperature is 18–25°C. This often results in a peak in feeding in late morning, followed by cessation of feeding at midday due to excessively hot temperature, and then perhaps a secondary peak in feeding in the afternoon as temperatures cool. Mass flights by adults take place only if air temperature is high, often about 29°C, but high densities and light wind are also required. When it is hot, grasshoppers tend to climb upward to escape the soil, which is usually considerably warmer

Migratory grasshopper is a defoliator, often completely removing leafy vegetation and leaving only stem tissue. Sometimes other tissue is eaten; heads of wheat may be clipped, for example. Migratory grasshopper thrives on rangeland that has a high density of broadleaf weeds, so it often moves from grazing land to nearby irrigated crops. In this behavior it differs from some other species, particularly differential grasshopper, Melanoplus differentialis (Thomas), and twostriped grasshopper, M. bivittatus (Say), which favor the taller undisturbed vegetation usually associated with fences and irrigation ditches, and usually do not develop high numbers on grazing land. Migratory grasshopper is often quite dispersive, and of course this behavior is the basis for its common name. When they are developing at high densities, the weather is abnormally warm, and a light wind is present, swarms of grasshoppers may disperse tens or even hundreds of kilometers and descend without warning to cause immense damage. With the availability of modern insecticides and aircraft for application, such potential disasters can be dealt with quickly and efficiently. When such insecticide and effective application technologies are not available, or where environmentally sensitive land or crops are concerned, grasshopper swarms can be disastrous. This species is the most important grasshopper pest in western North America.

Management Sampling. Grasshopper populations are usually assessed by visual observation. A sweep net is a useful tool to aid in collection, and its use is a prerequsite to identify the

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species complex. It is important to determine if grasshoppers collected from noncrop areas are crop-feeding species, because there are many nonpest grasshoppers that restrict their feeding to grasses or weeds. It is advisable to monitor nearby uncultivated land, particularly weedy areas, in addition to crop plants, due to the tendency of the pest species, to invade crops later in the season. Insecticides. Liquid formulations of insecticides are commonly applied to foliage to protect against damage. As grasshoppers rarely develop in crops, but instead invade from weedy areas, it is often the edges of crop fields that are most injured. Therefore, application of insecticide to the borders of crop fields is often adequate to protect an entire field. It is even better to apply insecticides to the developing grasshopper populations in weedy areas before they move to crops. This not only minimizes damage to crop plants, but also often results in younger grasshoppers being targeted for elimination. Younger grasshoppers are more susceptible to insecticides, with large nymphs and adults sometimes difficult to kill. Application of insecticide-treated bait is an effective alternative to foliar treatments for Melanoplus spp., because these grasshoppers spend considerable time on the soil where they come into contact with baits. Bait formulations are bulky and more difficult to apply than liquid products, so they are not often used, but have the advantage of limiting exposure of crops to insecticide residue and of minimizing mortality of beneficial insects such as predators and parasitoids due to insecticide exposure. Also, the total amount of insecticide active ingredient that is necessary to obtain control is usually considerably less when applied by bait, because the grasshoppers actively seek out and ingest the bait. Finally, for relatively expensive products that must be ingested to be effective, such as microbial insecticides, baits are the most effective delivery system. The attractant used most commonly for grasshopper bait is flaky wheat bran, though other products such as rolled oats are sometimes suggested. No additives, other than insecticide (traditionally 5% active ingredient, but decreasing concentrations of active ingredient now that more bioactive insecticides are available) are necessary because the wheat bran (or rolled oats) is quite attractive to Melanoplus grasshoppers. Other additives such as sawdust, water, vegetable or mineral oil, molasses, amyl acetate, salt, or sugar have been suggested, but provide little or no additional benefit over dry bran. The bait should be broadcast widely to maximize the likelihood of grasshopper contact, and be applied while they are in the late instars because the adults ingest less bait. Shotwell (1942) and Cowan and Shipman (1940) provided an excellent information on the formulation of grasshopper baits. Mukerji et al. (1981) gave an interesting perspective of bait use on rangeland. Cultural Practices. Elimination of weeds within, and adjacent to, crops is the most important cultural practice, and can have material benefit in preventing damage to crop

borders. However, during periods of weather when grasshoppers become numerous they may move long distances and invade crops. Tillage is an effective practice for the destruction of eggs, particularly in migratory grasshopper, which is especially likely to deposit eggs among crop plants. Deep tillage and burial are required, as shallow tillage has little effect. All the crop-feeding Melanoplus species deposit some eggs in crop fields, especially during periods of abundance, but it is fence row, irrigation ditch, field edge, and roadside areas that tend to be the favorite oviposition sites, so tillage is not entirely satisfactory unless other steps are taken to eliminate grasshopper egg pods from these areas that cannot be tilled. Although providing suppressive effects, deep tillage is not consistent with the soil and water management practices in many areas, so it may not be a good option. Row covers, netting, and similar physical barriers can provide protection against grasshoppers. This approach obviously is limited to small plantings, and can interfere with pollination. Also, grasshoppers are capable of chewing through all except metal screening, so this approach cannot guarantee complete protection. Biological Control. The opportunities for biological control are limited, but improving. Historically, poultry were found to consume many grasshoppers and could provide considerable relief if the grasshopper-infested garden was small to moderate in size, and the birds were plentiful. This remains a viable option for some people, and turkeys are usually considered the most suitable among poultry. The birds may also inflict some direct damage to plants, however, so introduction of poultry is probably most viable when grasshoppers are plentiful and threatening. The microsporidian fungus Antonospora (Nosema) locustae is well studied as a microbial control agent of Melanoplus spp. (Ewen and Mukerji, 1980; Johnson and Pavlikova, 1986; Johnson and Henry, 1987; Bomar et al., 1993), and is available commercially. Slamovits et  al. (2004) suggested that Nosema locustae be moved to the genus Antonospora. It is fairly stable, and easily disseminated to grasshoppers on bait. However, its usefulness is severely limited by the long period of time that is required to induce mortality and reduction in feeding and fecundity. Also, the level of mortality induced by the consumption of Antonospora (Nosema) is quite low (Johnson and Dolinski, 1997), often imperceptible. It is best used over very large areas, not just on individual farms, and should be applied at least 1 year in advance of the development of potentially damaging populations. More rapid suppression of grasshoppers is attainable by applying very high levels of Antonospora (Nosema) (Capinera and Hibbard, 1987), but this is not usually considered to be an economic approach, and commercial products are not prepared in this manner. Other species of Antonospora (Nosema) also exist naturally in North America, but have not been developed as bioinsecticides due to difficulties with mass production.

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Fungi have also been investigated for grasshopper suppression, and a grasshopper strain of Beauveria bassiana called GHA has been effective in some trials (Moore and Erlandson, 1988; Johnson and Goettel, 1993; Demirel and Cranshaw, 1996, 2006d; Jaronski and Goettel, 1997). Behavioral thermoregulation by grasshoppers, wherein they bask in the sun and raise their body temperatures, is potentially a limiting factor for use of fungi. Basking grasshoppers easily attain temperature in excess of 35°C (as can grasshoppers living in warm climates); such high temperature decreases or even prevents disease development in infected grasshoppers (Inglis et  al., 1996). Inconsistent quality control in the production of fungi also limits use of these organisms for grasshopper control. Products for grasshopper control that display minimal toxicity to mammals and other nontarget organisms are always desirable. In addition to microbes, insecticidal chemicals derived from plants and microbes offer potential in this regard (Foster et al., 1996; Sieglaff et al., 1997, 1998; Amarasekare and Edelson, 2004; Demirel and Cranshaw, 2006d; Capinera and Froeba, 2007; Sandoval-Mojica and Capinera, 2011). Grasshoppers display a reluctance to feed on plants treated with neem (azadirachtin, neem oil) products, which are botanically derived, because neem can also be insecticidal. However, neem does not persist in sunlight, so the benefits can be transient. Spinosad, a naturally derived insecticide produced by fermentation of the actinomycete bacterium Saccharopolyspora spinosa, is effective on grasshoppers, though it performs better by ingestion than by contact. This products does not cause rapid mortality, but after 2–4 days, mortality may be discernable.

Redlegged Grasshopper

Melanoplus femurrubrum (De Geer)

Southern Redlegged Grasshopper Melanoplus propinquus Scudder (Orthoptera: Acrididae)

Natural History Distribution. The redlegged grasshopper, Melanoplus femurrubrum (De Geer), is known from nearly all of the United States and southern Canada, and its distribution also extends south through most of Mexico. In parts of the southeast, on the coastal plain from eastern North Carolina to southern Mississippi and Louisiana, and including all of Florida, it is replaced by a very similar form, M. propinquus Scudder, which appears to be a separate species (Dakin, 1985). These are native insects. Host Plants. These species are polyphagous, feeding on a broad range of plants and apparently preferring

a ­dietary mixture over a single food plant, and broadleaf plants over grasses. The preferred habitat is tall vegetation in pastures, fence rows, along irrigation ditches and roadways, and in fallow agricultural fields which have become weedy. The redlegged grasshoppers are known throughout North America for damage to crops, attacking alfalfa, barley, birdsfoot trefoil, clover, corn, lespedeza, oat, orchardgrass, soybean, timothy, tobacco, and vetch in addition to vegetables. Among vegetable crops, bean, beet, cabbage, and potato seem to be most frequently injured, but nearly any crop may be fed upon. Among the uncultivated plants eaten are aster, Aster spp.; Kentucky bluegrass, Poa pratensis; brown knapweed, Centaurea jacea; cinquefoil, Potentilla argentea; ­dandelion, Taraxacum ­officinale; fleabane, Erigeron divergens; goldenrod, Solidago canadensis; kochia, Kochia scoparia; Russian thistle, Salsola kali; smooth brome, Bromus ­inermis; sweet clover, Melilotus officinalis; wavyleaf thistle, Cirsium ­undulatum; western ragweed, Ambrosia ­psilostachya; and likely many others. The weeds fed upon most frequently in North Dakota alfalfa fields were reported to be kochia, Kochia scoparia, field bindweed, Convolvulus arvensis; awnless bromegrass, Bromus inermis; and foxtail, Setaria spp. (Mulkern et  al., 1962). On prairie, redlegged grasshopper ate primarily Kentucky bluegrass, Poa ­pratensis; western ragweed, Ambrosia psilostachya; golden aster, Chrysopsis villosa; flixweed, Descurainia sophia; and leadplant, Amorpha canesens (Mulkern et al., 1964). Bailey and Mukerji (1976) were able to culture redlegged grasshopper on nearly all plants tested, though they completed development more rapidly when provided with plants on which they preferred to feed. Natural Enemies. The natural enemies of the cropfeeding Melanoplus spp. are quite similar. For information on natural enemies of redlegged grasshopper, see the section on natural enemies of migratory grasshopper, Melanoplus sanguinipes (Fabricius). Weather. Like most grasshoppers, redlegged grasshopper is favored by hot weather. Long-term periods of drought and hot weather favor population increase, especially in northern areas that normally are cooler. A certain amount of precipitation is necessary to provide adequate food for the grasshoppers, of course, but prolonged cool, wet weather, especially during the period of eggs hatch, is detrimental for survival. The late onset of winter can favor grasshopper population increase because it allows adults additional time to produce eggs. Life Cycle and Description. There is only a single generation per year throughout the range of these species, with the egg stage overwintering. Eggs hatch in late spring and adults are present from July until they are killed by heavy frost, though their numbers decrease steadily throughout the season. The following information has been derived from studies of M. femurrubrum, but likely applies equally well to M. propinquus.

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Egg. The eggs are elongate-cylindrical, and widest at the middle. They measure 4.1–4.6 mm long and 0.9–1.5 mm in diameter. Their color is yellowish brown or creamy white. They are deposited in structures called pods, which consist of two columns of eggs arranged in parallel rows. The pod, which is secreted by the female during oviposition, consists of frothy material secreted between, and covering, the eggs. The pod is a curved cylinder in form, and measures about 20–25 mm long and 3–5 mm in diameter. It is buried in the soil, and normally contains 20–26 eggs per pod. The upper portion of the pod consists solely of froth, and the young grasshoppers chew their way through this material to escape from the soil. Eggs are often deposited among the roots of grasses and weeds, particularly along the edges of crop fields. Females can produce 300 eggs during their life time. These are not early season grasshoppers. Hatching occurs about 3 weeks after hatching of twostriped grasshopper, Melanoplus bivittatus (Say), 2 weeks after migratory grasshopper, Melanoplus sanguinipes (Fabricius), and about the same time as differential grasshopper, Melanoplus ­differentialis (Thomas), with which redlegged grasshopper may cooccur. The period of hatching is extended, however, so nymphs can be found during most of the summer. Nymph. Development of the nymphal stage normally requires about 40 days, during which there usually are five to six instars. Throughout their development they are yellowish, but marked with a broad black stripe that extends across the face, eyes and prothorax, and onto the abdomen. They also bear a second, less discrete black stripe that is below, but parallel to the aforementioned stripe, and that arches across the lateral lobe of the prothorax. The outer face of the femora is marked with a broad black stripe. The underside is yellowish. The hind tibiae are yellow or gray, and bear black spines. The overall body length of nymphs is 4.0–5.6, 6.2–7.2, 7.4–9.7, 10.0–15.5, and 16.5–22.5 mm for instars 1–5, respectively. Antennal segment numbers increase from 12 to 14 to 15–16, 17–19, 22–24, and 24–26 in the corresponding instars. Bellinger and Pienkowski (1989) reported mostly six to seven instars in Virginia. They reported total nymphal development times for grasshoppers displaying a total of six instars to average 67, 42, 31, and 29 days when cultured at 26.5, 30, 35, and 38°C, respectively. Nymphs developing through seven instars required 5–10 additional days, depending on the rearing temperature. Mean instar development time was reported to be 3.6, 4.1, 4.7, 5.0, 5.8, and 7.9 days for instars 1–6 when cultured at 35°C, an optimal temperature. Nymphs change their behavior in response to temperature, with feeding commencing at 20–24°C, and nymphs moving to elevated perches to escape the heat when the air or soil temperatures get high. Like most grasshoppers, they tend to remain inactive, even at high temperature, in the absence of sunlight.

FIG. 11.9  Redlegged grasshopper. (Photo by J. Capinera.)

Adult. The adults are medium-sized grasshoppers, the males measuring 17–23 mm long, and the females 18–27 mm. They are reddish-brown or grayish-brown dorsally and yellow or yellowish-green ventrally. The front wings lack distinct markings; the hind wings are colorless. The lateral lobe of the pronotum is usually marked with a distinct black area. The outer face of the hind femora are yellowish but bear an indistinct dark stripe. The hind tibiae are almost always deep red with black spines, and are the basis for the common name of these grasshoppers. In the male, the tip of the abdomen is rather bulbous, with the subgenital plate bearing a broad “U”-shaped depression apically. The cerci narrow rapidly from the base, with the distal third narrow and the tip either angled (M. femurrubrum) or rounded (M. propinquus). The males, but not the females, of these two species are easily separated by the genitalia. In M. femurrubrum, the cerci are pointed at the tip, formed by an acute angle. The cerci at the midpoint are relatively broad, about one-half the maximum width at the base. The furcula in this species diverge from the base and then converge distally, the space between the arms of the furcula forms a “U” shape. In contrast, in M. propinquus the tips of the cerci are rounded, and the cerci are relatively narrow at the midpoint, about one-third the maximum width at the base. The furcula diverge more strongly, the space between the arms forms a “V” shape (Dakin, 1985). Adults normally roost at night on the tops of tall grasses and weeds. Early in the morning they crawl down the plant and resume feeding when the air temperature warms, often moving along the soil in search of food. As happens with nymphs, the adults ascend vegetation to escape high temperature. In the evening, they perch again on elevated roosts, and remain there until they are warmed by sunlight in the morning. Females have a preoviposition period of 9–15 days before they commence egg laying. Redlegged grasshopper is a fairly strong flier and can fly 10 m if disturbed.

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Despite its importance, the comprehensive treatment of M. femurrubrum biology is lacking. Some important aspects were given by Parker (1939), Shotwell (1941), and Bellinger and Pienkowski (1989). A very good synopsis was presented by Pfadt (1994d), who also pictured all stages of development. A summary of redlegged grasshopper biology, including keys to related Canadian Orthoptera, was given by Vickery and Kevan (1985). Melanoplus femurrubrum was included in many grasshopper keys, including those by Blatchley (1920), Dakin and Hays (1970), Helfer (1972), Capinera and Sechrist (1982), and Richman et  al. (1993). Blatchley (1920) includes M. propinquus; and Dakin and Hays (1970) treated the two redlegged grasshoppers as subspecies. Melanoplus femurrubrum was also included in a key to grasshopper eggs by Onsager and Mulkern (1963), and in a field guide to grasshoppers (Capinera et al., 2004). Rearing of Melanoplus spp. was described by Henry (1985).

Damage Redlegged grasshopper is a defoliator, often removing all leaf tissue and leaving only plant stems. Lower densities leave plant ragged or tattered. Redlegged grasshopper is a common component of the grasshopper complex that affects plants growing along the margins of fields, though it causes extensive damage only during periods of very high density. Redlegged grasshopper is capable of developing high densities and migratory tendencies during periods of drought, and may be found mixed into swarms of migratory grasshopper, Melanoplus sanguinipes (Fabricius).

Management Management of the various Melanoplus spp. grasshoppers is substantially the same. For information on redlegged grasshopper management, see the section on management under migratory grasshopper, Melanoplus sanguinipes (Fabricius).

Twostriped Grasshopper

Melanoplus bivittatus (Say) (Orthoptera: Acrididae)

Natural History Distribution. This native grasshopper is widely distributed in northern North America. In the United States, it extends from the Atlantic to the Pacific Oceans, and is absent only from the Gulf Coast region. In Canada, it occurs from Nova Scotia to British Columbia. Host Plants. This species is adaptable, and is found in a variety of habitats. However, it is most abundant in moist meadows, dense vegetation along water courses, and in disturbed, weedy areas. In the Great Plains region, it is abundant in moist tallgrass regions, but uncommon in

the drier ­shortgrass prairie. Twostriped grasshopper feeds on both grasses and broadleaf plants, but prefers the latter and fares poorly in habitats lacking broadleaf plants. Plants in the families Compositae and Cruciferae seem to be preferred. Among the uncultivated vegetation consumed is arrowleaved colt’s foot, Petasites sagittatus; burdock, Arctium lappa; dandelion, Taraxacum officinale; dog mustard, Eruscastrum gallicum; flixweed, Descurainia sophia; needleleaf sedge, Carex eleocharis; leadplant, Amorpha canescens; oxeye daisy, Chrysanthemum leucanthemum; pepperweed, Lepidium densiflorum; plantain, Plantago major; redtop, Agrostis alba; sand dropseed, Sporobolus cryptandrus; Canada thistle, Cirsium arvense; sunflower, Helianthus spp.; wavyleaf thistle, Cirsium undulatum; mustard, Brassica spp.; and others. Mulkern et  al. (1962) determined the plants consumed by twostriped grasshoppers in North Dakota alfalfa fields, and reported that the plants most often consumed, after alfalfa, were kochia, Kochia scoparia; wild oat, Avena fatua; awnless bromegrass, Bromus inermis; flixweed, Descurainia sophia; marsh elder, Iva xanthifolia; and quack grass, Agropyron repens. On prairie, however, the plants most often consumed were Kentucky bluegrass, Poa pratensis; leadplant, Amorpha canescens; and western ragweed, Ambrosia psilostachya (Mulkern et al., 1964). Survival rates and body weights of twostriped grasshopper are higher, and development times shorter, on mixed diets than on single hosts (MacFarlane and Thorsteinson, 1980). Bailey and Mukerji (1976) were able to culture twostriped grasshopper on nearly all plants tested, though they completed development more rapidly when provided with plants on which they preferred to feed. Twostriped grasshopper commonly infests vegetable and field crops, though most injury is limited to field margins. Among vegetable crops injured are beet, cabbage, chicory, corn, lettuce, onion, potato, and likely others. Field crops such as alfalfa, birdsfoot trefoil, clover, young barley and oat, timothy, vetch, and the immature seedheads of wheat are also fed upon. Flowers and ornamental plants likewise are attacked. Natural Enemies. The natural enemies of the crop-­ feeding Melanoplus spp. are quite similar. For information on natural enemies of twostriped grasshopper, see the section on natural enemies of migratory grasshopper, Melanoplus sanguinipes (Fabricius). Weather. Survival and population increase in grasshoppers are favored by hot weather. High numbers tend to occur after a period of years with abnormally hot and dry weather during the spring and summer months. This is especially true in northern areas, where it tends to be cooler. Enough precipitation is required to provide adequate food for grasshoppers, of course, but protracted periods of rainfall during egg hatch, especially if accompanied by cool weather, disrupt feeding by young grasshoppers and induce high mortality. Late onset of winter can favor grasshopper population increase because it allows adults additional time to produce eggs.

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Life Cycle and Description. Over most of its range, twostriped grasshopper displays one generation annually, with the egg stage overwintering. In Colorado, eggs begin to hatch in June, though hatching can occur over a 4–6week period. Nymphs may be present until September, but adults appear beginning in July. Oviposition commences in August and continues until adults are killed by cold weather. At higher elevations in British Columbia, a 2-year life cycle is reported (Beirne, 1972). Egg. The egg is elongate-cylindrical, with the ends tapering to blunt points. The eggs are olive in color and measure about 5.0 mm long and 1.2 mm in diameter. The reported values of number of eggs per pod and total fecundity vary considerably among studies. Drake et  al. (1945) reported 69.7 per pod and a mean total of 129 eggs per female, whereas Smith (1966) reported 43.3 eggs per pod and a total of 355. The number of pods produced per female ranges from 4 to 15, with a mean interval between oviposition of 4 days (Smith, 1966), but this assumes good weather. Eggs are arranged in columns of four within a frothy secretion; the egg structure is called an egg pod. The pods are curved, measure 30–38 mm long and 6–7 mm in diameter. They are inserted into the soil at a depth of 2–5 cm, and topped with a frothy plug. Favorite oviposition sites are along fence rows, ditch banks, and pastures with compact, undisturbed soil. Pods are often inserted among the roots of plants, and a soil moisture content of 10%–20% is preferred. The act of oviposition requires about 2 h. The egg is the overwintering stage, and typically persists in the soil for 7–8 months. Embryonic development begins in the summer and autumn after oviposition, and is 60%–80% complete before embryos enter diapause for the winter. However, they can be induced to hatch if exposed to about 5°C for 90 days. Nymph. Nymphal development normally requires 30– 50 days. Most nymphs display five to six instars, but seven instars occur occasionally. Nymphal development time when fed lettuce or alfalfa and cultured at 21°C is about 10, 8.5, 10, 11, and 14 days, respectively, for instars 1–5 (Langford, 1930). The young nymphs initially are dark brown or greenish, but gain a distinct dark stripe along the pronotum behind each eye at the third instar. The wing development is poor until instar 3, and the developing wings are pointed downward. At instar 4, the wing orientation is reversed, with the wings oriented upward, but also pointing posteriorly. In the fifth instar, the wings are quite evident and extend out to at least the second abdominal segment. The number of antennal segments is 12–13, 17–18, 19–22, 23–25, and 24–26 for instars 1–5, respectively. Corresponding body lengths are 5.0–6.6, 7.4–10.4, 9–14, 15– 21, and 20–17 mm. Nymphs are found on the soil and seek food each morning, but may ascend plants to escape the heat of the soil by noon. Like the adult stage, nymphs can perch on elevated roosts at night, and can sun themselves in the morning and in cool weather to attain optimal body temperature.

FIG. 11.10  Twostriped grasshopper. (Photo by J. Capinera.)

Adult. This is a fairly large and robust species. Males measure 23–29 mm long and females 29–40 mm. The general body coloration is olive or brownish-green dorsally and yellowish or yellowish-green ventrally. The head and pronotum tend to be darker, usually olive green. A narrow but distinct yellow stripe passes from behind each eye along the pronotum and forewings, extending nearly to the wing tips. The stripes are often bordered below with black, especially on the anterior portions of the body. The stripes come together posteriorly in the forewings, forming a “V” shape. It is this pair of yellow stripes that is the basis for the common name of this grasshopper. The front wings are usually uniform in color except for the stripes, and the hind wings colorless. The hind femora are yellow with a dark stripe along the outer face. The hind tibiae are variable, usually reddish but also greenish, yellowish, and purplish, and equipped with black spines. The male cerci are short, broad, and boot shaped. Adults seek crop borders and roadsides for oviposition. Mated females have a 7–14 day preoviposition period, after which they oviposit within the roots of grasses and weeds. Duration of the oviposition period is about 30 days (range 15–55 days). Aspects of the biology of twostriped grasshopper were treated by many authors, including Parker (1939), Shotwell (1941), Church and Salt (1952), and Smith (1966). An excellent synopsis was presented by Pfadt (1994e), who also pictured all stages of development. A summary of twostriped grasshopper biology, including keys to related Canadian Orthoptera, was given by Vickery and Kevan (1985). Melanoplus bivittatus was included in many grasshopper keys, including those by Blatchley (1920), Dakin and Hays (1970), Helfer (1972), Capinera and Sechrist (1982), and Richman et al. (1993). This species was also included in a key to grasshopper eggs by Onsager and Mulkern (1963), and in a field guide to grasshoppers (Capinera et  al., 2004). Rearing of Melanoplus spp. was described by Henry (1985).

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Damage Twostriped grasshopper consumes the leaves of numerous plants. Damage is greatest in areas adjacent to weeds, and along fence rows, irrigation ditches, roadsides, and fallow fields. Damage is exacerbated by drought, which apparently increases nymphal survival rates and decreases the amount of weed vegetation available to the grasshoppers. Although the grasshoppers feed at night if it is sufficiently warm, where nights are cool these grasshoppers tend to perch on elevated objects. This behavior allows them to be warmed by the light from the setting and rising sun, and maximizes their period of activity. This also results in nibbling on the resting substrate by the grasshoppers; these grasshoppers feed on the bark of bushes and young trees, and even damage shingles on buildings and eat holes in vinyl window screens while perching. The nymphs and adults are fairly dispersive, and walk tens or even hundreds of meters in the search for food. At high densities, they show propensity to swarm, which is expressed by band formation in the nymphal stage and flight by adults. The temperature threshold for flight is 30–32°C. Ascending to heights of 200–500 m, and flying with the wind, swarming adults can disperse long distances.

Management Management of the various Melanoplus spp. grasshoppers is substantially the same. For information on twostriped grasshopper management, see the section on management under migratory grasshopper.

FAMILY GRYLLIDAE—FIELD CRICKETS Fall Field Cricket

Gryllus pennsylvanicus Burmeister

Spring Field Cricket

Gryllus veletis (Alexander and Bigelow)

Southeastern Field Cricket Gryllus rubens Scudder (Orthoptera: Gryllidae)

Natural History Distribution. Field crickets are extremely difficult to distinguish based on the appearance, so for many years they have been grouped into one species, Gryllus assimilis (Fabricius). As the significance of the calling behavior (chirping) became known, some of the species have been distinguished, but much work remains. The biology and damage potential of the various field crickets is confused by the problems with identification. The most economically

significant North American species are fall field cricket, Gryllus pennsylvanicus Burmeister; spring field cricket, Gryllus veletis (Alexander and Bigelow); and southeastern field cricket, Gryllus rubens Scudder; though other species may be locally important. The fall and spring field crickets are most abundant in the northern states and southern Canada, whereas the southeastern field cricket is known from the southeastern states. These are native insects. Host Plants. These field crickets are found widely in grassy fields, pastures, weedy areas, roadsides, and lawns, and occasionally feed on foliage, flowers, and fruit of crop plants. Among the vegetable crops damaged are bean, beet, carrot, cabbage, cantaloupe, cucumber, lettuce, parsnip, pea, potato, pumpkin, squash, sweet potato, tomato, watermelon, and likely others. Other crops known to be injured by field crickets are alfalfa, barley, corn, cotton, flax, rye, strawberry, sweet clover, and wheat. Flowers and seeds of weeds are also suitable food for crickets, and among those known to be consumed are foxtail, Setaria spp.; lambsquarters, Chenopodium album; pigweed, Amaranthus spp.; ragweed, Ambrosia spp.; Russian thistle, Salsola kali; sunflower, Helianthus spp.; and wheatgrass, Agropyron spp. Russian thistle is reported to be particularly suitable, and foliage as well as flowers and seeds are eaten. The foliage of broadleaf plants is preferred over grasses. Natural Enemies. Many natural enemies of field crickets are known, though their importance is poorly documented. Eggs are parasitized by Ceratotelia marlattii Ashmead and Paridris brevipennis Fouts (both Hymenoptera: Scelionidae), with 20%–50% of the eggs in South Dakota reportedly parasitized by C. marlatti and 1%–5% by P. brevipennis. Nymphs and adults are killed by Exoristoides johnsoni Coquillett and Euphasiopteryx ochracea (Bigot) (both Diptera: Tachinidae), Sarcophaga kelleyi Aldrich (Diptera: Sarcophagidae), mermithid nematodes (Nematoda: Mermithidae), and horsehair worms (Nematomorpha). Gregarine parasites (Protozoa: Sporozoa) infest the guts of field crickets (Zuk, 1987), but have little effect on their host other than to extend the cricket developmental period. Among the predators that take advantage of cricket abundance are many birds, but particularly crows, Corvus bachyrhynchos brachyrhynchos Grehm; and ring-necked pheasant, Phasianthus torquatus Gmelin; as well as snakes, toads, and gophers. Life Cycle and Description. Field crickets differ in their life history, but the aforementioned species exemplify the common life cycles. Gryllus pennsylvanicus and G. veletis have one generation per year, whereas G. rubens has two generations annually. In South Dakota and most of its range, G. pennsylvanicus eggs overwinter, hatching occurs in the spring, nymphs develop in early summer, adults appear beginning in July, and eggs are deposited in August and September. The presence of adults in the late summer and autumn is the basis for its common name of “fall” field cricket. In Michigan, adults chirp from early August until mid-November (Alexander and Meral, 1967).

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In the same area, G. veletis nymphs overwinter in the later instars, the adult stage is attained in May–June, eggs are produced in May–June, hatching is completed by the end of August, and nymphs complete part of their development before the onset of winter. The presence of adults during spring and early summer is the basis for its common name of “spring” field cricket. In Michigan, adults chirp from mid-May until early August. Thus, though these two field cricket species occupy the same habitat, there is seasonal separation of life stages. In Louisiana and most of its range G. rubens overwinters as nymphs, the adult stage is attained by April when eggs are produced, and nymphs are matured by July. Secondgeneration adults begin to appear and produce eggs in late July–August, and nymphs develop until winter, failing to molt to the adult stage until the subsequent spring. Egg. Eggs are deposited in firm, damp soil, usually within the upper 2 cm of soil. Under dry conditions, crickets may deposit their eggs within the cracks formed as moist soil hardens. The eggs are elongate-cylindrical, with bluntly rounded ends. They are also slightly curved, with one side convex and the opposite side concave. Eggs normally measure about 3 mm long and 0.6 mm wide. They are light yellow or cream. As the embryo develops, it may change shape and color, and increase slightly in size. Such eggs become barrel-shaped, and measure about 4 mm long and 0.85 mm wide. They are deposited singly, but several are usually placed in close proximity. The number of eggs produced by each female ranges from about 150 to 400. Except for overwintering eggs, most hatch in about 2 weeks. Nymph. The young cricket hatching from an egg is faced with the difficult task of burrowing through soil to reach the surface. The form of the cricket that escapes the egg is called the “vermiform larva,” and differs from the following nymphal stages by being encased in a transparent membrane that immobilizes the appendages. Once the vermiform larva wriggles to the surface the membrane is shed, the legs are freed, and the young first instar cricket is able to walk and jump. The cricket undergoes several molts, growing at each stage. In G. pennsylvanicus and G. veletis, normally there are eight to nine instars, though there may be more under adverse conditions. Mean duration of instars under field temperature in South Dakota was reported to average about 8.5, 8.2, 8.9, 8.8, 9.3, 10.7, 11.9, 12.0, and 14.6 days for instars 1–9, respectively. In G. rubens, the number of instars averages 10, but the same pattern of latter instars requiring more time for development is apparent. Total nymphal development time usually requires 80–90 days in the cooler northern environments inhabited by G. pennsylvanicus and G. veletis, and 70– 80 days in the warmer environments inhabited by G. rubens. The nymphs initially are brownish, but marked with black and sometimes yellow in the thoracic area. They resemble the adults, and bear long antennae and cerci, but

lack wings. With each succeeding molt, the nymphs become darker. The ovipositor begins to appear in instars 3–4, the wing pads in instar 6. By instar 8, both the ovipositor and wing pads are apparent. Body length is about 3, 3.3–3.9, 4.2–5.0, 5–6, 7–8, 7.3–8.5, 9–12, 12–18, and 13.5–20 mm for instars 1–9, respectively.

FIG. 11.11  Southeastern field cricket. (Photo by L. Buss.)

Adult. The adult cricket is mostly shiny black, though the front wings may be brownish black. The front wings usually cover the abdomen or extend slightly beyond the tip. The hind wings, however, are variable, and wing length determination has both environmental and genetic components (Walker, 1987). In shortwinged forms, at least half of the abdomen is covered, but such insects are incapable of flight. Longwinged forms tend to be a small component of the population (Veazey et  al., 1976; Harrison, 1979), and though they are capable of flight, they do not often fly (Walker and Sivinski, 1986; Walker, 1987). Body length of adults is 15–26 mm. Females bear a long ovipositor, which often equals the length of her body. The width of the head is greater than the pronotum in G. pennsylvanicus and G. veletis, whereas in G. rubens the head is narrower than the pronotum. Males of these cricket ordinarily space themselves in a field and remain rather sedentary for all of their adult lives. They produce a song that attracts roving females. The song of G. pennsylvanicus and G. veletis is intermittent, which consists of about 150–240 discrete chirps per minute at 29°C, with chirps comprised of four pulses at about 25 pulses per second. In contrast, G. rubens produces a nearly continuous song, consisting of about 60 pulses per second at 29°C, with a considerably louder song than the other species. Females usually mate within 1–4 days of attaining the adult stage, and normally they mate repeatedly during the oviposition period. Most females commence oviposition from 7 to 14 days after attaining the adult stage, though some require up to 30 days. Egg production continues for the life of the cricket, with 50–60 days considered to be about average adult longevity.

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Crickets are usually active at night, though they may venture forth during daylight during cloudy weather, and in late afternoon. Males normally call at night, but in cold weather may call during warmer daylight hours. Typically, crickets hide beneath debris during the day, but sometimes seek shelter in soil cracks or excavate small chambers in the soil. They sometimes appear to be gregarious, because they are found clustered, but this is simply a reflection of preference for a habitat with limited availability. Alexander (1957) discussed the taxonomy of eastern field crickets and included keys to separate most species based on morphology; however, the fall and spring field crickets were treated as spring and fall broods of “pennsylvanicus.” Nickle and Walker (1974) provided keys to the southeastern species. An excellent treatment of fall and spring field cricket biology (as G. assimilis) was provided by Severin (1935), with similar treatment of (probably) G. rubens (as G. assimilis) by Folsom and Woke (1939). Calling behavior of G. pennsylvanicus and G. veletis was described by Alexander and Meral (1967), and of G. rubens by Doherty and Callos (1991). Synopses of fall and spring field crickets, including keys to related Canadian Orthoptera, were given by Vickery and Kevan (1985). These crickets are included in a field guide (Capinera et al., 2004) to North American Orthoptera. Culture of field crickets was described by Winewriter and Walker (1988).

bait ­formulations ­usually eliminate molasses and other additives, employing only coarse bran and insecticide. Insecticides can also be applied to sawdust and distributed broadcast or in a band around the margin of a field; in such cases, crickets perish from contact rather than ingestion of the bait, and this approach is most appropriate when cricket densities are very high (Blank et al., 1985). Foliar applications of insecticides can also be effective if they are residual, but are less selective. Soil tillage can destroy eggs, and clean cultivation deprives overwintering insects of shelter. Crickets can be attracted to broadcast sounds that approximate the calling of cricket species (e.g., Walker, 1993), providing a selective mechanism of population sampling. However, not all crickets fly, as some species lack long wings and some have both short and longwinged morphs. Thus, it is difficult to use this technology for quantitative estimates of population density.

FAMILY GRYLLOTALPIDAE—MOLE CRICKETS Shortwinged Mole Cricket

Neoscapteriscus abbreviatus (Scudder)

Southern Mole Cricket Damage Field crickets are omnivorous, and feed on a variety of plant and animal matter. They may consume the roots, stems, leaves, flowers, fruits, and seeds of plants, but the flower, fruit, and developing seeds are preferred. Once mature, seeds are no longer suitable. They also eat nearly any dead insect they encounter, sometimes are reported to be cannibalistic, and gnaw on animal and plant products such as fur, wool, linen, and cotton. They are rarely considered to be serious plant pests, and only when exceedingly abundant. Spring field cricket, in particular, is questionable as a plant pest, because it is not very gregarious. Field crickets also display predatory behavior which offsets their occasional tendency to feed on valuable plants. For example, grasshopper eggs, flea beetle adults, fly puparia, caterpillar pupae, and insects from spider webs are among the documented animal material eaten by nymphs and adults (Burgess and Hinks, 1987).

Management Suppression of crickets is rarely necessary, but the application of insecticide-treated bran bait is effective if needed. Often its application can be limited to the edge of fields, where crickets dispersing into crops would encounter and ingest the poison bait. A formula for preparing baits was given by Severin (1935), but modern

Neoscapteriscus borellii (Giglio-Tos)

Tawny Mole Cricket

Neoscapteriscus vicinus (Scudder) (Orthoptera: Gryllotalpidae)

Natural History Distribution. These mole crickets were inadvertently introduced to the southeastern United States in about 1900. Shortwinged mole cricket, Neoscapteriscus abbreviatus Scudder, was first observed at Tampa, Florida in 1899, but separate introductions were discovered near Miami in 1902 and Brunswick, Georgia in 1904. Southern mole cricket, Neoscapteriscus borellii Giglio-Tos (known until recently as S. acletus Rehn and Hebard), was similarly introduced to major seaports, beginning with Brunswick in 1904, and followed by Charleston, South Carolina in 1915, then Mobile, Alabama in 1919, and finally Port Arthur, Texas in 1925. Tawny mole cricket, Neoscapteriscus vicinus Scudder, was first observed at Brunswick, Georgia in 1899. The origin of these crickets is uncertain, but Argentina and Uruguay are likely sources, because they occur in these areas of southern South America. In the years since introduction to the United States, the Neoscapteriscus spp. have expanded their ranges, but they differ considerably in their current distribution. Shortwinged mole

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cricket, which is flightless, remains fairly confined to the southern Florida and southern Georgia-northeast Florida introduction sites, though it also occurs in Puerto Rico and the Virgin Islands. It has been redistributed in southern Florida, but is largely found in coastal areas. In contrast, southern mole cricket is now found from North Carolina to eastern Texas, including the northern regions of Georgia and Alabama and the entire peninsula of Florida, and recently was detected in Yuma, Arizona. Tawny mole cricket is somewhat intermediate in its spread; it occurs from North Carolina to Louisiana, and throughout Florida, but thus far remains restricted to the southern coastal plain. These are not the only mole crickets found in North America, but they are most damaging. For example, a native species, the northern mole cricket, Neocurtilla hexadactyla (Perty), is widely distributed in the eastern states west to about South Dakota and Texas, and including southern Ontario, but is not a pest. European mole cricket, Gyllotalpa gryllotalpa (Linnaeus), has been introduced from Europe into the northeastern states, but is of minor significance. Changa, Neoscapteriscus didactylus (Latreille), invaded Puerto Rico from South America before 1800, and has caused considerable damage to crops on this island. Host Plants. Mole crickets are omnivorous, feeding on animal as well as plant material. Several studies have indicated that when provided with grass or collected from grassdominated habitats, southern mole cricket is less damaging than tawny mole cricket. Southern mole cricket feeds mostly on other insects, whereas tawny mole cricket is principally herbivorous (Matheny, 1981; Matheny et  al., 1981; Walker and Ngo, 1982). Both species are associated with tomato and strawberry fields (Schuster and Price, 1992). Among vegetable crops reported to be injured are beet, cabbage, cantaloupe, carrot, cauliflower, collards, eggplant, kale, lettuce, onion, pepper, potato, spinach, sweet potato, tomato, and turnip. Other plants injured include chufa, turf, and pasture grasses, peanut, strawberries, sugarcane, tobacco, and such flowers as coleus, chrysanthemum, and gypsophila. Among the turf grasses, bahiagrass and bermudagrass are commonly injured by tawny mole cricket, whereas St. Augustine grass and bermudagrass are favored by shortwinged mole cricket. Mole crickets also feed on weeds such as pigweed, Amaranthus spp. Natural Enemies. Few natural enemies of Scapteriscus mole crickets exist naturally in North America. Among the natural enemies are amphibians such as toads, Bufo spp.; birds such as sandhill cranes, Grus canadensis; and mammals such as armadillos, Dasypus novemcinctus. They, and the few predatory insects that attack crickets such as tiger beetles (Coleoptera: Cicindelidae), are not effective. Therefore, several natural enemies have been introduced from South America (Parkman et  al., 1996). The most effective introduced beneficial insect is the parasitoid Ormia depleta (Wiedemann) (Diptera: Tachinidae), which was imported from Brazil. This fly is attracted to the calls of male mole crickets. Its release has resulted in reduced mole cricket injury in southern Florida

(Frank et al., 1996). A less effective parasitoid is Larra bicolor Fabricius (Hymenoptera: Crabrionidae), which was imported from Bolivia but seems to be constrained by availability of suitable adult food sources in the United States (Frank et al., 1995). An entomopathogenic nematode, Steinernema scapterisci, was introduced from Uruguay (Nguyen and Smart, 1992). It is fairly specific to mole crickets, persists readily under Florida’s environmental conditions, and is dispersed by crickets. Field collections consistently show infection levels of 10% or greater (Parkman et al., 1993a,b; Parkman et al., 1996), and infected crickets die within 10–12 days. Together, these natural enemies have suppressed mole crickets to much lower population densities in most locations where they are well established. Life Cycle and Description. Southern and tawny mole cricket are quite similar in appearance and biology. Shortwinged mole cricket differs in appearance owing to the short wings but also in behavior, because it has no calling song and the short wings render it incapable of flight. Typically, the eggs of these three species are deposited in April–June, and nymphs predominate through August. Beginning in August or September some adults are found, but overwintering occurs in both the nymphal and adult stages. Maturity is attained by the overwintering nymphs in April, and eggs are produced at about this time. A single generation per year is normal, though in southern Florida there are two generations in southern mole crickets and an extra peak of adult flight activity in the summer, resulting in spring, summer, and autumn flights from the two generations (Walker et al., 1983). Egg. The eggs are deposited in a chamber in the soil adjacent to tunnels. The chamber is usually constructed at a depth of 5–30 cm below the soil surface. It typically measures 3–4 cm in length, width, and height. The eggs are oval to bean-shaped, and initially measure about 3 mm long and 1.7 mm wide. The eggs increase in size as they absorb water, eventually attaining a length of about 3.9 mm and a width of 2.8 mm. The color varies from gray or brownish. They are deposited in a loose cluster, often numbering about 25–60 eggs. Duration of the egg stage is about 10–40 days. Total fecundity is not certain, but over 100 eggs have been obtained from a single female, and the mean number of egg clutches produced per female was reported to be 4.8 (Hayslip, 1943). Nymph. Hatchlings are whitish initially but turn dark within 24 h. They may consume the egg shell or cannibalize siblings, but soon dig to the soil surface. The juvenile stages resemble the adults, but nymphs have poorly developed wings. The number of instars is variable, probably 8–10 (Hudson, 1987). Nymphs and adults create extensive below-ground tunnel systems, usually within the upper 20–25 cm of soil. When the soil is moist and warm they tunnel just beneath the surface, but crickets tunnel deeper if the weather becomes cooler or the soil dries. They come to the surface to forage during the evening, usually appearing shortly after dusk if the weather is favorable.

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large spots dorsally, and smaller spots dorsolaterally. These crickets measure 22–29 mm long. The two dactyls on the foretibiae are slightly divergent, and separated at the base by a space equal to at least half the basal width of a dactyl. Shortwinged mole cricket makes no calling song, producing only a weak 1–5 pulse chirp during courtship.

FIG.  11.12  Comparative images of mole crickets: shortwinged (left), tawny (middle), and southern (right). (Photo by L. Buss.)

Adult. Mole crickets have peculiar enlarged forelegs that are used for digging in the soil. The foretibiae have large blade-like projections, called dactyls, and the number and arrangement of dactyls are diagnostic. These crickets also bear antennae which are shorter than the body. Females lack a distinct ovipositor. Both sexes have elongate cerci at the tip of the abdomen. The male produces a courtship song that is attractive to females; they normally call during the night. Except for the shortwinged mole cricket, the male enlarges the entrance to his burrow, forming a horn-shaped opening, in preparation for calling. This increases the volume of the call, and allows flying females to locate males. Mating occurs within the male’s burrow in the soil, and apparently the female may usurp the burrow after mating.

FIG. 11.14  Southern mole cricket. (Photo by P. Choate.)

Southern mole cricket has long hind wings that extend beyond the tip of the abdomen. The front wings are longer than the pronotum, about two-thirds the length of the abdomen. They are broad and rounded at the tips. This cricket is brown, with the dorsal surface of the pronotum often quite dark. As with shortwinged mole cricket, in southern mole cricket the two dactyls on the foretibiae are separated at the base by a space equal to at least half the basal width of a dactyl. Thus, these two species can be distinguished by the wing length. Southern mole cricket produces a calling song that consists of a low-pitched ringing trill at about 50 pulses per second. It is usually emitted during the first 2 h after sunset.

FIG. 11.13  Shortwinged mole cricket. (Photo by P. Choate.)

Shortwinged mole cricket bears front wings that are shorter than the pronotum. The front wings cover the hind wings, which are minute. The body is mostly whitish or tan in color, though the pronotum is brown mottled with darker spots. Also, the abdomen is marked with a row of

FIG. 11.15  Tawny mole cricket. (Photo by P. Choate.)

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Tawny mole cricket is quite similar to southern mole cricket in general appearance, with moderately long front wings and long hind wings, a yellowish-brown body, and a dark pronotum. It can be distinguished from southern mole cricket by dactyl form. The tibial dactyls are nearly touching at the base, separated by less than half the basal width of a dactyl. Tawny mole cricket produces a loud, nasal trill at about 130 pulses per second during the first 90 min after sunset. Summaries of mole cricket life history were given by Worsham and Reed (1912), Thomas (1928), Hayslip (1943), and Walker (1984), though biology of shortwinged mole cricket is poorly documented. Keys to North American and Caribbean area mole crickets were provided by Nickle and Castner (1984). These crickets are also included in a field guide (Capinera et al., 2004).

Damage The crickets usually damage seedlings, feeding aboveground on foliage or stem tissue, and below-ground on roots and tubers. Girdling of the stems of seedling plants at the soil surface is a common form of injury, though young plants are sometimes severed and pulled below-ground to be consumed. Additional injury to small plants is caused by soil surface tunneling, which may dislodge seedlings. Southern mole cricket does much more tunneling injury than tawny mole cricket.

Management Sampling. Various approaches to population estimation have been developed. A commonly used, but less reliable technique, is the assessment of population density by the frequency of soil surface tunneling. Tunneling is affected by soil moisture levels, and is most appropriate for assessing nymphs. A more consistent, but labor intensive, approach for estimation of nymph and adult abundance is flushing with about a 0.5% aqueous solution of dishwashing soap. Soil flushing is affected by soil moisture conditions, with greater cricket extraction efficiency as the soil approaches field capacity (Hudson, 1989). Flushing with synergized pyrethrin insecticide solution is equally effective (Hudson, 1988). A soil washing apparatus also has been developed to separate crickets from soil (Fritz, 1983). Adults can be captured with sound traps that use electronic sound synthesizers to lure crickets to a catching device, usually a large funnel. Insecticides. Liquid and granular formulations of insecticides are commonly applied to the soil to suppress mole crickets. Insecticide application should be followed by irrigation, because the insecticide needs to enter the root zone of the plants to be most effective. Bait formulations are also useful. Various baits have proven effective, but most contain wheat bran, cottonseed meal, or some other grain product

plus 2%–5% toxicant. Also, addition of 5%–15% water and 2%–5% molasses to the grain-toxicant mixture are sometimes recommended (Thomas, 1928; Walker, 1984). Cultural Practices. Most injury to vegetable transplants occurs on small plants, so placement of larger plants is suggested as a strategy to avoid injury (Schuster and Price, 1992). Crickets can quickly invade crop land that has been fumigated or otherwise cleared of crickets, so isolation from sources of crickets, or planting in large blocks of land with proportionally little edge, is desirable (Poe, 1976). Host Plant Resistance. Efforts have been made to find turf and pasture grass varieties that are resistant to attack by mole crickets. If grass varieties contain antibiotic properties, or otherwise limit the reproductive abilities of mole crickets, this can translate into fewer crickets seeking food within vegetable crops, because grass-containing fields are a principal source of mole crickets. Thus far, strains have been identified which are fairly tolerant of feeding or which are not preferred by mole crickets, primarily the finer textured grass selections, but considerable improvement in these grasses is needed before they can affect cricket population biology. Biological Control. Biological control of mole crickets can be enhanced by the application of the entomopathogenic nematode Steinernema scapterisci (Nguyen and Smart), which was imported from Uruguay and established in Florida. It is inconsistently available from commercial producers of biological control agents, but when available it can be suspended in water and applied to soil in the same manner as an insecticide for long-lasting suppression of mole crickets. It is more effective when applied to adults than when applied to nymphs. The other imported biological control agents can also be relocated, though it is uncertain how they will cope with climatic zones different from Florida.

FAMILY TETTIGONIIDAE—SHIELDBACKED KATYDIDS Mormon Cricket

Anabrus simplex Haldeman

Coulee Cricket

Peranabrus scabricollis (Thomas) (Orthoptera: Tettigoniidae)

Natural History Distribution. Despite the name “cricket” applied to these species, they are not members of the superfamily Grylloidea, which contains the crickets. Rather, they are members of superfamily Tettigonioidea and the family Tettigoniidae, the katydids. Most interesting is that they are in the subfamily

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Tettigoniinae, the predaceous katydids. The dark coloration of these insects (at least when they occur abundantly) and flightless condition likely accounts for their names, because (superficially) they have a cricket-like appearance. These similar insects are native to western North America. Mormon cricket occurs widely, with a range that includes southern British Columbia to Manitoba in the north, and south to northern California and northern New Mexico. As a persistent pest, however, its range is limited to the Rocky Mountain and Great Basin regions. Coulee cricket is even more limited in occurrence, and is known from Montana, eastern Washington, northeast Oregon, and southern British Columbia. Indeed, of the 122 species in the subfamily Tettigoniinae found in the United States, all but 10 occur west of the Great Plains. Thus, this group is clearly centered in western North America. Host Plants. Over 400 species of grasses, forbs, trees, and shrubs were reported by Swain (1944) to be eaten by Mormon cricket. Most of these records occurred during the arid “dust-bowl” era of the 1930s when cricket densities were extremely high and vegetation was scarse. Thus, these observations are not typical of cricket feeding behavior. These crickets are often considered to be omnivorous, but despite their wide potential host range they display some specific preferences unless confronted by starvation. Among plants consumed, crickets often feed preferentially on the flowers and seed heads, ignoring the leaf material. Seed-head consumption is especially pronounced in grasses, though grasses are a minor component of cricket diet. Forbs such as bitterroot, Lewisia rediviva; wild onion, Allium spp.; arrowleaf balsamroot, Balsamorhiza sagittata; wild mustard, Brassica spp.; tumble mustard, Sisymbrium altissimum; and lupine, Lupinus spp.; are preferred by nymphs. In the adult stage, crickets eat mostly big sagebrush, Artemisia tridentata (MacVean, 1987; Redak et al., 1992). In an analysis of Mormon cricket diet in Colorado, Ueckert and Hansen (1970) reported that the diet consisted of forbs 50%, arthropods 21%, fungi 16%, grasses 6%, clubmoss 5%, and grasslike plants 2%. These crickets actively prey on other insects, including cicadas, ants, aphids, and beetles if they have the opportunity to catch them. They are also quick to consume injured or dead Mormon crickets as well, and one of the distinctive characteristics about these crickets is their tendency to stop and feed on comrades that have been injured in some manner or crushed on roadways by vehicles. As the healthy crickets remain on the highways to feed on fallen crickets, they may also become crushed by ongoing vehicular traffic, resulting in long, dark, greasy road slicks consisting of pulverized crickets. Among the vegetables reported to be damaged by crickets are bean, beet, cabbage, cantaloupe, carrot, cauliflower, Chinese cabbage, corn, lettuce, onion, potato, pumpkin, radish, rutabaga, salsify, spinach, tomato, turnip, and likely

others. Not readily eaten are pea and mustard. Other crops susceptible to injury are alfalfa, barley, clover, flax, millet, oat, sugar beet, sweet clover, timothy, and wheat. Due to the nature of the cropping systems in the areas inhabited by these crickets, alfalfa and wheat are the crops most often injured, and most consumption of vegetation is limited to rangeland plants. Natural Enemies. Predators are perhaps the best-known mortality factor associated with crickets. Destruction of crickets in 1848 by California gulls, Larus californicus, saved the early Mormon settler’s grain crops; a fact commemorated by a large statue of the gulls in Salt Lake City. Gulls are not the only vertebrates attracted to these insects when they become numerous, and among the avian predators most commonly observed feeding on crickets are crows, Corvus brachyrhynchos; hawks, Falco spp. and possibly others; meadowlarks, Sturnella magna; and blackbirds, various species (Wakeland, 1959). Mammals such as coyotes, Canis latrans; ground squirrels, Citellus spp.; and kangaroo rat, Dipodomys spp. also feast on crickets when they are abundant. Also, the wasps Palmodes laeviventris (Cresson) and Tachysphex semirufus (Cresson) (both Hymenoptera: Sphecidae) capture crickets and feed them to their young. Despite the frequency at which predation is observed, there is little evidence that predators are normally effective at maintaining crickets at low densities, or capable of suppressing crickets during periods of population outbreak. Parasitism is surprisingly uncommon in cricket populations. Only the egg stage is parasitized with any degree of frequency, and though levels of up to 50% parasitism have been reported, it is usually quite low. The parasitoids responsible for attacking eggs are Sparaison pilosum Ashmead (Hymenoptera: Scelionidae) and Oencyrtus anabrivorus (Hymenoptera: Encyrtidae). A fly, Sarcophaga harpax Pandelle (Diptera: Sarcophagidae), has been reared from adult crickets, but occurs infrequently. Pathogens vary greatly in their effect on crickets. The microsporidian fungus Heterovesicula (Vairimorpha) cowani can naturally infect substantial proportions of Mormon cricket populations, and causes rapid mortality when young crickets ingest spores (MacVean and Capinera, 1991, 1992; Lange et al., 1995). Heterovesicula appears to be the most important pathogen of crickets. The report that the g­ rasshopper-infesting microsporidian fungus Antonospora (Nosema) locustae can infect Mormon cricket (Henry and Onsager, 1982) seems to be premature (MacVean and Capinera, 1992). Slamovits et  al. (2004) suggested that this species be moved to the genus Antonospora. The nematode Agamaspirura anabri (Nematoda) (Christie, 1930) and the horsehair worm Gordius robustus (Nematomorpha) (Thorne, 1940) have also been observed in crickets. Gordius was reported to be quite common in crickets near standing water, because part of the horsehair worm’s life cycle takes place in water; however, water is not plentiful in the habitat of these crickets.

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Life Cycle and Description. The life cycle and description of Mormon cricket and coulee cricket are nearly identical. Normally, there is one generation per year, though there are reports of eggs at high altitudes remaining in diapause for an entire year, resulting in a 2-year life cycle. They overwinter, with egg hatch occurring in March–May, often while snow remains on the ground. The nymphs are present until June when adults begin to emerge and start egg production. Adults usually perish by late August, often earlier. About 100 days is required for the nymphal and adult stages to be completed. Mormon cricket and coulee cricket are differentiated by the texture of the dorsal surface of the pronotum; it is punctate or rough in coulee cricket but smooth in Mormon cricket.

the ovipositor becomes increasingly obvious. Mean duration (range) of development period of crickets cultured at 21–26°C is 9.5 (7–12), 7.4 (4–14), 5.1 (4–7), 6.5 (4–10), 5.6 (3–9), 5.6 (5–7), and 10.4 (5–15) days, respectively, for instars 1–7. Thus, the mean total development time of nymphs is estimated at 50 days (range 43–58 days) but weather, and probably density, can significantly affect development rate. Mature nymphs have small wings but the wings do not protrude from beneath the pronotum. The crickets tend to aggregate, seeking shelter together beneath bushes and debris during inclement weather and at night. Once they reach the third or fourth instar crickets the aggregations begin to move long distances, with numerous crickets coalescing into groups, which move in bands. The density of crickets in bands may be 10–30 per m2, but sometimes much lower. The width of a band is often 300 m or more, but only 10 m deep, with crickets moving in the same direction along the entire width. The crickets all seem to move independently and consistently in the same direction, often at 1 km per day. There is no indication that they follow one another, and the basis of orientation is unknown. Bands moving in different directions sometimes converge and then emerge without loss of individual band integrity. Key elements in the movement of cricket bands are high density and recurring contact with conspecifics; both are important in maintaining the integrity of the bands (Sword, 2005).

FIG. 11.16  Eggs of Mormon cricket. (Photo by J. Capinera.)

Egg. The egg is elliptical and measures about 7–8 mm long and 2.0–2.5 mm wide. Initially, brown in color, it soon turns whitish and then gray. They are deposited in the soil singly or in small clusters at a depth of 6–25 mm during the summer, where they remain until spring. One end of the egg, where the head of the embryo is located, swells slightly before egg hatching. Sometimes they are deposited around the base of plants, but more often bare soil is favored, including the mounds of ants. Females deposit, on average, about 85 eggs, but up to 160 per female has been observed. They complete their embryonic development in the summer and autumn, before entering diapause. Thus, they are ready to emerge early in the spring, and begin to hatch when soil temperature attain about 5°C, a much lower temperature than the threshold of development for hatch of grasshoppers. Nymph. Upon hatching from the soil, the nymphs are dark, resemble the adults, and measure about 6 mm long. There are seven instars, and by the time they attain the last instar they are about 30 mm long. The initial instars are black with white along the lateral edge of the posterior end of the pronotum, and the ovipositor of the female is not apparent. As they attain the fourth instar, however, they acquire green, red, purple, or brown color and in the female

FIG. 11.17  Female of adult Mormon cricket. (Photo by J. Capinera.)

Adult. The adult is very similar to the mature nymph in form and color but larger, measuring 35–45 mm long. Also, the sword-shaped ovipositor of the adult female is much longer, and the short wings of the adult male protrude from beneath the pronotum and are used as a stridulatory organ. As happens with nymphs, the adults may cluster under shelter both during the evening and inclement weather. They can also climb into bushes to escape the hot soil during excessively warm weather. Adults continue to move in bands in the same manner as nymphs, stopping only to eat and oviposit.

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Reproduction commences 10–14 days after attaining the adult stage. Males call from perches on vegetation during the morning hours. Females compete for the attention of males, mount the males, and are inseminated. Males are selective in their choice of partners, often choosing the largest female with which to mate. During insemination the male provides the female, attached to the sac containing sperm, a large proteinaceous mass that protrudes from her genital opening. While the sperm is draining into the female’s reproductive system, the proteinaceous mass provides a meal for the female. This is a significant investment on the part of the male, as the sperm and its accompanying protein meal represents up to 25% of his body weight (Gwynne, 1984). Breeding usually occurs in hilly areas where vegetation is sparse. Mormon cricket (and presumably coulee cricket) exists in solitary and gregarious forms. The aforementioned description applies mostly to the gregarious form. Only this damaging form has been thoroughly studied and the solitary forms are rarely observed. The solitary form occurs at low density between periods of population outbreak and in areas where crickets do not become numerous. In contrast to the gregarious form, solitary crickets are green, and do not aggregate or form bands. Sexual selection behavior is also reversed, with females choosing among males. This change in mating behavior seems to be related to better nutrition of crickets when they are not at high densities (Gwynne, 1993). The gregarious and solitary forms are quite distinct in morphology and in production of calling (mating) songs, likely because they also tend to remain in different geographic areas. However, the two forms can mate successfully, so they are not completely reproductively isolated (Bailey et al., 2007). Biology of Mormon cricket was described by Cowan (1929), and of coulee cricket by Melander and Yothers (1917). The impact of Mormon cricket was summarized by Wakeland (1959), and a modern review was given by MacVean (1987). A key to Mormon and coulee crickets, and their near relatives, was published by Rentz and Birchim (1968). Pfadt (1994c) provided a concise summary of Mormon cricket biology and pictures all stages of development. Synopses of Mormon cricket and coulee cricket, including keys to related Canadian Orthoptera, were given by Vickery and Kevan (1985). Mormon cricket and some other shield-back katydids are shown in a field guide by Capinera et al. (2004).

Damage Mormon and coulee crickets normally occur in arid sagebrush rangeland, and generally cause little injury unless they move into irrigated cropland. On rangeland, Mormon cricket herbivory often is largely restricted to sagebrush and

forbs, with no loss in understory plant biomass at densities of eight crickets per square meter, and with little dietary overlap between crickets and cattle (MacVean, 1989; Redak et  al., 1992) In earlier times, when settlers had to be nearly self-sufficient, vegetable gardens were critically important to ranchers, and crop losses caused by Mormon and coulee crickets were a significant threat to the existence of western communities. Now, however, vegetable production is usually not a commercial enterprise and not essential for continued existence of ranchers in these arid lands, and control technologies have improved markedly, so cricket importance has declined. Crickets remain a threat, however, and when bands of crickets invade lush crop vegetation, usually irrigated grains and alfalfa, they can cause serious defoliation.

Management Sampling. Cricket bands are easily detected when they cross roads, and their presence in an area rarely is a surprise. However, they move rapidly and their course of travel is unpredictable, so when crickets are discovered control efforts are usually directed at the bands before they enter cropgrowing areas. Some areas, usually mesas, seem to support continuously breeding populations, and serve as a source of crickets for nearby regions. Insecticides. Persistent insecticides are sometimes applied by aircraft to foliage in areas supporting nymphal populations or migrating bands. An alternative is to apply insecticide-treated bait. The preferred bait is flaky wheat bran, and it may be applied dry or with 10% water, but other additives such as molasses do not increase effectiveness (Cowan and Shipman, 1940). Cultural Practices. In earlier times, a common practice to prevent invasion of crop fields by cricket bands was to surround the crop with ditches possessing steep sides; crickets falling into such ditches had great difficulty regaining the soil surface. Similarly, vertical barriers of metal topped by a deflector served to prevent crickets from entering areas surrounded by such “cricket fences.” Biological Control. Although highly desirable, presently there are no effective means of implementing biological control of Mormon crickets. The microsporidian fungus Heterovesicula (Vairimorpha) cowani offer the greatest potential for long-term suppression because it persists naturally in Mormon cricket populations, but it is not commercially available. The microsporidian fungus Antonospora (Nosema) locustae was reported to affect Mormon crickets (Henry and Onsager, 1982) but this was not verified in later tests (MacVean and Capinera, 1992). Similarly, tests with Metarhizium spp. fungi for Mormon cricket control have been disappointing (Keyser et al., 2017).

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Chapter 12

Order Thysanoptera—Thrips

American Bean Thrips

Caliothrips phaseoli Hood

Bean Thrips

Caliothrips fasciatus (Pergande) (Thysanoptera: Thripidae)

Natural History Distribution. Caliothrips fasciatus and C. phaseoli are apparently native species that have spread to many other parts of the world, including Asia, Europe, and South America. In North America, C. fasciatus is known principally from the western United States, including Oregon, Idaho, and Wyoming south to California and Arizona, and south into Mexico. It is most damaging in the dry interior valleys of California, and arid regions of nearby states. Caliothrips phaseoli occurs in Southwest America as well, but is relatively more common in the southeastern states. Caliothrips phaseoli occurs widely in the Caribbean and South America, and is known to be damaging in Brazil, though it is not well studied. Host Plants. These species are reported from numerous plants, though some are incidental hosts, on which the adult can feed but not reproduce successfully. As suggested by the common names, these thrips are most often damaging on legumes, but have been reported to severely damage bean, cantaloupe, lettuce, and pea. Vegetable crops from which bean thrips have been reported include asparagus, bean, beet, cabbage, cantaloupe, carrot, cauliflower, corn, fennel, garlic, kale, leek, lettuce, melon, onion, pea, pepper, potato, radish, Swiss chard, tomato, and turnip. Other hosts include field crops such as alfalfa, clover, cotton, and hops; fruit such as apple, avocado, fig, grape, orange, peach, pear, persimmon, plum, prune, and tangerine; and ornamentals such as canna, California poppy, geranium, gladiolus, hollyhock, iris, nasturtium, and sunflower. Many wild hosts are known, including trees, grasses, and weeds. Among common weeds known to support bean thrips are Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00012-1 © 2020 Elsevier Inc. All rights reserved.

field bindweed, Convolvulus arvensis; milkweed, Asclepias spp.; mallow, Malva parviflora; mullein, Verbascum virgatum; redroot pigweed, Amaranthus retroflexus; common sow thistle, Sonchus oleraceus; and prickly lettuce, Lactuca scariola. In California, the abundance of prickly lettuce is one of the most important elements in the ecology of C. fasciatus. Bailey (1937) provided a more complete list of host plants. Natural Enemies. Field studies suggest that only about 40% of C. fasciatus thrips attain the adult stage. Much of the mortality results from the feeding habits of general predators such as minute pirate bug, Orius tristicolor (White) (Hemiptera: Anthocoridae), the larvae of the predatory thrips Aelothrips fasciatus (Linnaeus) and A. kuwanae Moulton (Thysanoptera: Thripidae), the lacewing Chrysopa californica Coquillette (Neuroptera: Chrysopidae), and the convergent lady, Hippodamia convergens Guerin-Meneville (Coleoptera: Coccinellidae). An important natural enemy of C. fasciatus seems to be the internal parasitoid Ceranisus russelli (Crawford) (Hymenoptera: Eulophidae). Levels of parasitism by C. russelli observed in the field have been quite variable rates, but up to 70% have been noted. In Brazil, Orius insidiosus (Say) (Hemiptera: Anthocoridae) is an important predator of C. phaseoli (Mendes and Bueno, 2001). An unspecified nematode was found in C. fasciatus nymphs, but the importance of this observation is uncertain. Weather. Caliothrips fasciatus, like most thrips, suffer direct mortality due to the weather conditions. They are largely restricted to areas with mean winter temperature of about 20°C or greater. However, during the winter, the adults can survive temperature as low as − 9°C for short periods of time. Rainfall is also important. Heavy rains dislodge larvae from the plant, washing them to the soil where they perish. Summer rainfall in excess of about 10 cm is detrimental, but winter rainfall is also detrimental to overwintering adults. Thus, rainfall is likely the principal factor restricting C. fasciatus damage to arid regions of western states. On the other hand, in Brazil, C. phaseoli is reported to be relatively insensitive to weather (Boiça et al., 2015). 581

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This is likely due to the more uniform weather conditions as well as inherent differences between the Caliothrips spp. Weather also indirectly affects thrips populations. Caliothrips benefit from an abundance of early-season and late-season weeds if they are to attain high densities. In the absence of spring or autumn rains, the thrips populations are checked by the shortage of suitable host plants. Thus, a critical balance of rainfall is required; enough early and late in the season to assure adequate food, but not so much in the summer and winter to cause direct mortality from being crushed and drowned. Life Cycle and Description. These species display the developmental patterns typically seen in the vegetable crop-infesting thrips: egg, larva I, larva II, propupa, pupa, and adult. As with many other thrips, several overlapping generations occur annually. Six annual generations of C. fasciatus are estimated from California. These thrips apparently overwinter in the adult stage. In California, overwintered C. fasciatus adults generally produce the first generation on weed hosts in March. Overwintered adults usually perish by late April or May. Adults from the first and second generations may find the weeds suitable, or may move to alfalfa to feed. By late June, however, thrips begin dispersal to numerous cultivated crops, and weeds contained within the crops, where additional generations develop. As crops mature, thrips disperse again to late-planted crops, where an additional generation develops. The first generation in the spring requires about 6 weeks for completion, but as the weather warms the generations are completed in about 3 weeks. Egg. The egg is bean shaped, bluntly rounded at both ends, with one side concave and the opposite side converse. The egg measures about 0.2 mm long and 0.1 mm wide. The egg is white and is inserted by the female into the leaf tissue. The fecundity of females has been poorly studied, but females commonly produce five or more eggs per day when held in captivity. Larva. There are two larval stages in bean thrips. They are quite small in size, measuring about 0.3 and 0.9 mm long, respectively. Duration of the first instar is about 5.4 and 4.3  days at 21°C and 32°C, respectively. Duration of the second instar is about 8.2 and 3.8  days at 21°C and 32°C, respectively. Thus, larval development requires about 13.6  days at relatively cool temperature and only about 8.1 days at relatively warm temperature. The larvae of the bean thrips tend to be reddish yellow or pink. When they first hatch, however, they are nearly translucent white, and as they mature they gain more red coloration, becoming almost entirely crimson in color at the end of the larval stage. Larvae lack wings, and are found feeding on foliage, often in association with adults. The larvae typically carry a drop of liquid excrement at the tip of their slightly upturned abdomen. In most respects, they resemble the adult stage.

After completing the two larval instars, the insect drops to the soil and burrow to depths up to 35 cm. Most are found at depths of 7–15 cm, with burrowing behavior affected by soil type, moisture level, and shading. Dry, fine, and sandy soil is not conducive to thrips burrowing and survival. Propupa and pupa. Two additional immature stages of development, the propupa (often called prepupa) and pupa are passed in the soil. In general appearance, they greatly resemble the larval and adult stages. They differ from both, however, in having partly developed wings, or wing pads, and by not feeding. The propupa and pupa are orange, with crimson markings on the thorax and abdomen. They measure about 1.0 mm long during the propupal stage, but shrink slightly to about 0.8 mm during the pupal stage. Also, they can be differentiated from each other in that the wing pads are much larger in the pupal stage. Duration of the propupal stage is only 1–2 days, with 1.6 days the normal development period at 21°C, and 0.9 days at 32°C. The pupal stage is longer, requiring 9.3 and 2.4 days at 21°C and 32°C, respectively. Adult. The adult is minute, the female measuring only about 1.0 mm long and the male 0.9 mm. The body color is grayish black, and the front wings are banded with two alternating light and dark areas. The hind wings are entirely dark. Both sets of wings are fringed, as is normal for insects of this order. When the wings are folded against the body, which is the usual condition, the body appears to bear two whitish bands across the central region. The eight-­ segmented antennae and the legs are also banded with alternating light and dark areas. Adults begin to copulate and commence oviposition 2–4  days after emergence. A sex ratio of about two females per one male is normal for this species. The adult overwinters within curled leaves, on the underside of pubescent foliage, under the old coverings of scale insects, and in other sheltered locations. Plants that remain green throughout the winter are favorite overwintering locations. If disturbed, the adults slowly move, but little or no feeding occurs during this period. The most complete treatment of C. fasciatus thrips biology was provided by Bailey (1933a). However, a later treatment by Bailey (1937) and an earlier report by Russell (1912) also contain useful information. Caliothrips spp. are included in keys by Palmer et  al. (1989), Mound and Kibby (1998), Mound and Marullo (1996), and Hoddle et  al. (2012). Also, Caliothrips fasciatus and C. phaseoli are included in a key to common vegetable-infesting thrips in Appendix A. Nakahara (1991) published a key to the 10 Nearctic species of Caliothrips.

Damage Damage results from larvae and adults feeding on the underside of foliage. In the process of feeding, the thrips extrude

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their stylets, puncture cells, and drain the liquid contents. Apparently, the larvae feed more extensively and inflict more damage than do adults. The insects feed gregariously, so there tends to be a great amount of tissue destruction localized on individual leaves, and these leaves turn whitish and often are shed by the plant. The underside of the foliage becomes covered with black spots of fecal material. Injury tends to be concentrated on leaves of intermediate age, though adults tend to move to newer tissue as it becomes available. As the foliage is depleted, thrips move to bean pods and stem material to feed. The scar resulting from insertion of the eggs into foliage seems to cause little injury.

Management Sampling. Thrips populations are best assessed by close visual observation of susceptible crops and suitable alternate hosts, such as early-season weeds. Boiça et al. (2015) indicated that higher thrips populations were found on the lower portions of the bean plants. Insecticides. Formerly a serious pest of several crops, bean thrips have been reduced to minor status by the introduction of modern insecticides. Foliar contact and systemic insecticides are generally effective, but because the propupal and pupal stages are sheltered in the soil, persistent insecticides or multiple applications are necessary. Thorough coverage of the lower surface of the leaves is also important. Cultural Practices. The most important cultural practice is weed management. Elimination of early-season weeds in, or adjacent to, crops can reduce thrips populations. Irrigation ditches and other sites where weeds may survive during winter and periods of drought should be examined for thrips, and be treated with insecticides if necessary. Overhead irrigation can decrease bean thrips densities, and reduce the need for insecticides. In Brazil, C. phaseoli cultivars varied among bean varieties during the winter season, but not during the dry season.

Chilli Thrips

Scirtothrips dorsalis Hood (Thysanoptera: Thripidae)

Natural History Distribution. Though native to the Indian subcontinent, S. dorsalis has been redistributed to many other areas of southern and eastern Asia. It is also found in parts of Africa, Middle East, Oceania, Central and South America, and the Caribbean. It has very limited distribution in Europe, and in North America it is known mostly in the warm-weather states of Florida, Georgia, Texas, and Hawaii, but also New York and Massachusetts. It was first found in the United States in Florida, beginning in 1991. It is expected eventually to spread to all the southern and Pacific Coast states.

In greenhouses, it could survive colder climates, and likely escape and inoculate nearby areas annually. Host Plants. Chilli thrips display a wide host range, feeding on more than 100 species in over 40 plant families. Some of these plants may allow survival but not support reproduction, so as often is the case with host range observations, the number of true or reproductive hosts is somewhat less than its dietary breadth. The principal plant association of chilli thrips is the family Fabaceae, but it is associated with numerous crops. Among vegetable crops attacked are bean, corn, eggplant, melon, various peppers including chilli pepper, pumpkin, watermelon, squash, cucumber, and sweet potato. Other food crop hosts include banana, citrus, cocoa, grape, kiwi, litchi, longan, mango, peanut, blueberry, and strawberry. Cotton and tea are hosts, as are many ornamental plants such as rose, viburnum, camellia, bottle brush, coleus, snapdragon, zinnia, and coreopsis. They do not feed on flower pollen. Seal et al. (2010) allowed adults to select from several prospective vegetable crops for oviposition by moving from infested cotton plants. The largest numbers of dispersing adults were found on pepper and rose, whereas low numbers were found on tomato, squash, beans, and eggplant. Larvae and pupae resulting from the dispersing adults were abundant on pepper and rose, found in small numbers on squash and tomato, and absent from bean and eggplant. Thus, although the purported host range is broad, these data demonstrate that not all “hosts” are equally attractive. Interestingly, the suitabilities of these different hosts, as judged by insect size and duration of the developmental stages when confined to a single host plant, were not very different among the different host plants. This suggests that a large number of vegetable crops can support chilli thrips, though not equally well. Localized specialization has also been reported, wherein one plant species is favored in one area whereas another species is favored in another location. The presence of cryptic species in a “Scirtothrips ­dorsalis species complex” is one possible explanation for this phenomenon. Natural Enemies. Predators are important natural enemies of all thrips, most notably minute pirate bugs (Hemiptera: Anthocoridae). Predatory mites such as Neoseiulus cucumeris (Oudemans) and Amblyseius ­swirskii (Athiot-Henriot) (both Acari: Phytoseiidae) have been demonstrated to provide suppression on pepper (Arthurs et al., 2009; Dogramaci et al., 2011). In some locations, the entomogenous nematodes Thripinema spp. (Tylenchida: Allantonematidae) infect thrips. Although they are not pathogenic, infection causes sterilization of females. Life Cycle and Description. This species displays the developmental pattern typically seen in vegetable cropsinfesting thrips: egg, larva I, larva II, propupa, pupa, and adult. Chilli thrips have four to eight generations per year in temperate areas of Japan, and overwintering occurs as

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pupae in the soil, but under warmer conditions more generations can occur, possibly up to 18. Development time varies with environmental conditions, but under constant temperature conditions of 26°C on pepper, the duration of the egg stage is about 6.8 days, the combined larval instars require about 8.4  days, the combined propupal and pupal instars require about 2.6 days, and the preoviposition period of the adults is about 3.6  days. Thus, about 21  days are needed for a complete generation (from egg to onset of oviposition) (Seal et al., 2010). They can develop at temperatures from 9.7°C to 33°C. They do not overwinter successfully if exposed to temperatures of − 4°C for more than a few days. Like many thrips, they are favored by dry conditions. Duraimurugan and Jagadish (2011) and Kumar et al. (2013) provide information on chilli thrips biology. Egg. Adults insert eggs into above-ground plant tissue, on or near leaf veins of tender leaves. The eggs are kidney shaped or oval, and creamy white. The eggs measure about 0.23–0.26 mm long and 0.10–0.12 mm wide. Females can deposit 60–200 eggs over the course of its life. When developing at 26°C in pepper tissue, the mean (± SE) duration of this stage is 6.8 (± 0.2) days, with a range of 4–8 days at normal temperatures.

FIG. 12.1  Head and thorax of chilli thrips. (Photo by J. Capinera.)

Larva. As with other pest thrips, there are two larval instars. First instars are pale or creamy white in color, becoming more yellow-orange in the second. Both instars have seven-segmented antennae. They tend to gather near the mid-vein or borders of the damaged portions of the leaf. When cultured at 26°C on pepper tissue, the mean (± SE) duration of the combined first and second larval instars was 8.4 (± 0.2) days, or 8–10 days at normal temperatures. The mean (± SE) length and width of the first instar were reported to be 0.38 (± 0.02) and 0.10 (± 0.01), respectively. The corresponding values for the second instar were 0.70 (± 0.01) and 0.19 (± 0.01), respectively. Propupa and pupa. The propupa is yellow, with shortwing pads reaching only the third abdominal segment, and

short antennae. The duration of the propupa is 1.0 (± 0.23) days (range of 0.75–1.5 days). The size of the propupa averages about 0.59 mm (range 0.59–0.61 mm) in length and 0.22 mm (range of 0.21–0.24 mm) in width. The pupa is dark yellow with pinkish eyes. The antennae are longer in the pupa, reaching over the head and thorax. The wing pads are also longer, extending to the eighth abdominal segment. When cultured at 26°C on pepper tissue, the mean (± SE) duration of the combined propupal and pupal periods was 2.6 (± 0.2) days; about 24 h of this development time is represented by the propupa. The mean (± SE) length and width of the pupa were reported to be 0.8 (± 0.01) and 0.21 (± 0.01), respectively. These thrips normally drop to the soil for pupation and pupae are found in the soil or leaf litter, but sometimes they become wedged in the leaf axils or curled leaves or other tight spots. Adult. The adult is pale yellow to yellowish orange, about 1.2 mm long, and bears dark-brown antecostal ridges. Antecostal ridges are a thickening on the anterior margin of each abdominal body segment. Dark spots forming an incomplete dorsomedial stripe are found on the abdomen. The wings are shaded dark centrally, but the shading does not form dark bands. The postocular setae are equal in length. As with larvae, the adults tend to gather near the mid-vein or borders of the damaged portions of the leaf. Adult flight and oviposition activities are higher in the afternoon (1200– 1400 h) (Seal et al., 2010). When the immature stages were cultured at 26°C and fed pepper, the mean (± SE) duration of the resultant adult stage was 15.6 (± 0.4) days. The adults average about 1.2 mm in length, the males averaging smaller than the females. Females can reproduce without mating. Identification of thrips is difficult, and most often left to experts; good sources of identification include Mound and Marullo (1996), Mound and Kibby (1998), Hoddle et  al. (2012), and Cluever and Smith (2017).

Damage Chilli thrips typically feed on young, expanding plant tissue of stems, leaves, flowers, and fruit. They do not feed on mature tissue. They feed on individual cells of the plant tissue, extracting their contents. Initially, the death of the cells may cause a speckled appearance (silvering), but as large numbers of cells die tissue necrosis develops. Leaf curling and other forms of tissue deformation may occur. Initially, the thrips feed on the lower (abaxial) surface of leaves, but as they become abundant they may disperse to the upper (adaxial) surface. Heavy infestation results in leaf drop. Feeding by S. dorsalis can also result in transmission of plant viruses. Among the viruses known to be transmitted by chilli thrips are chilli leaf curl virus, peanut necrosis virus, peanut yellow spot virus, tobacco streak virus,

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­ atermelon silver mottle virus, capsicum chlorosis virus, w and melon yellow spot virus.

Management Sampling. The distribution of chilli thrips on plants can be affected by the host species, plant age, and thrips age. However, infestations typically begin with young tissue, so these sites should be monitored especially closely. Population assessment is usually accomplished by visual counts, although the infested plant material can be examined on the plant or removed for closer examination. Often, plant material is washed in 70% alcohol to facilitate removal, and then sieved through 300 a mesh sieve to separate the thrips. In addition, the attraction of insects to traps of various colors can be used to concentrate insects, and if adhesive is applied to the colored surface, the insects are retained for easy tabulation. In the case of chilli thrips, yellow, blue, white, or green are attractive colors (Chu et al., 2006; Kumar et al., 2013). Insecticides. Insecticides are the principal method of chilli thrips suppression, and the insecticides used include products from several mode of action classes. Because insecticide resistance is quickly developed in thrips species, rotation of products from different classes of insecticides is recommended. Many products are applied to the foliage, but some systemic products can be applied to seed at planting time, or as soil drenches after the plants are partly grown. The bioinsecticides Beauveria bassiana, Metarhizium anisopliae, and azadirachtin (neem) provide some short-term protection. Cultural Practices. Thrips are easily dispersed by infected plant material, so sanitation is important. Intercropping favored plants with less suitable plants seems to diminish the negative effects of the thrips. Partially resistant varieties of chilli peppers have been identified in India, and in the United States, 14 out of 158 cultivars evaluated were found to be resistant (Kumar et al., 2013).

Common Blossom Thrips

Frankliniella schultzei (Trybom) (Thysanoptera: Thripidae)

Natural History Distribution. Frankliniella schultzei is found in tropical and subtropical areas throughout the world. In the United States, it is limited principally to Florida and Hawaii, but it also occurs elsewhere in greenhouses. It also occurs nearby in Bermuda and many islands in the Caribbean, and seems likely to spread further within North America. It is an important pest in South America, and growing in importance in Florida. Host Plants. Common blossom thrips is known principally as a pest of cotton, peanuts, bean, and pigeon pea.

However, it has a wide host range, and is found at least on 83 species from 35 families of plants. Among vegetable crops affected are tomato, eggplant, peppers, onion, sweet potato, watermelon, melon, pumpkin, lettuce, spinach, lentil, bean, and cowpea. In a survey of selected vegetable crops in southern Florida, F. schultzei was most abundant in blossoms of tomato, squash, and cucumber, relative to bean and pepper (Kakkar et al., 2012). Cucumber supported the largest number of immature thrips, documenting its suitability for reproduction. Tree fruit, strawberry, and some flower crops also support F. schultzei. Natural Enemies. This insect is not yet well studied in North America. As with most thrips, predatory mites (Phystoseiidae) and minute pirate bugs (Anthocoridae) are often suggested as important biological control agents. However, when Kakkar et  al. (2016) assessed Neoseiulus cucumeris (Oudemans) (Phytoseiidae) and Amblyseius swirskii Athias-Henriot (Phytoseiidae) for suppression of blossom thrips they were not effective, whereas they were effective for Thrips palmi Karny (Thysanoptera: Thripidae). See the sections on onion thrips, Thrips tabaci Lindeman, or western flower thrips, Thrips occidentalis Pergande, for additional discussion of natural mortality factors. Life Cycle and Description. This species is somewhat unusual in having two distinct color forms. In the continental areas of the United States, as well as Central and South America, the dark form occurs, whereas in Hawaii the light form is found, and in Puerto Rico both color forms exist. The dark form is also found in Europe, whereas in Africa and Asia both color forms exist. Pinet and Carvalho (1998) studied the biology of F. schultzei on tomato in Brazil when cultured at 24.4°C. A summary of biology and characters used for identification is given by Kakkar et al. (2017). This species seems not to be as well known as many others. It displays the typical development pattern found among thrips that are vegetable crop pests: egg, larva I, larva II, propupa, and pupa before the adult stage. In the warm environments where they typically occur, there are several overlapping generations annually. Egg. Females deposit their eggs in the blossoms of host plants. Pinet and Carvalho (1998) reported that the embryonic period averaged 4.3 days when adults had been fed tomato foliage and maintained at about 26°C. Larva. Pinet and Carvalho (1998) reported that average development time of the first and second instars each was 2.5 days when fed tomato foliage and maintained at about 26°C. Propupa and pupa. The larval instars are followed by a propupal and pupal stage. Pinet and Carvalho (1998) reported that average development time of the propupa and pupa were 1.2 and 2.1  days when larvae had been fed

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t­omato foliage and maintained at about 26°C. Unlike the larval instars, the propupa and pupa do not feed. Adult. Pinet and Carvalho (1998) reported that average longevity of males and females was 13.1 and 13.6  days when fed tomato foliage and maintained at about 26°C. The preoviposition period averages 1.7  days, the oviposition period 9.8  days, and the postoviposition period 3.0  days. Frankliniella schultzei reproduce both sexually and parthenogenetically. Keys to the genera of thrips can be found in Palmer et al. (1989). Additional useful keys containing F. schultzei are found by Mound and Kibby (1998), Mound and Marullo (1996), Hoddle et al. (2012), and Cluever and Smith (2017). Also, a key to common vegetable-infesting thrips is found in Appendix A.

Damage Common blossom thrips feed on pollen, flowers, and leaves, which can lead to blossom drop and plant stunting. Given the choice between feeding on the pollen, petal, and leaf tissues of wax mallow, Malvaviscus arboreus Cav. (Malvaceae), the insects chose petal tissue (Milne et  al., 1996). Discoloration of blossoms can be indicative of a large thrips infestation. Fruit is often deformed by thrips feeding early in the fruit’s development. According to Pereira et al. (2016), these thrips have the greatest impact on the vegetative stage (of watermelon); the flowering and fruiting stages were less susceptible to damage. The thrips can also vector Tospovirus plant diseases such as tomato-spotted wilt virus, tomato chlorotic spot virus, and groundnut ringspot virus. The dark- and light-colored forms of this species are reported to differ in their ability to transmit tospoviruses (Kakkar et al., 2017).

Management Sampling. Visual examination of blossoms can be used to estimate abundance of this thrips, and insecticide treatment only when needed can reduce insecticide use as compared to regularly scheduled treatment. The thrips tend to be concentrated in the upper portions of plants, and Pinto et al. (2017) noted that the best location for sampling watermelon was the most apical leaves. Bacci et  al. (2008) also recommended sampling terminal leaves of cucumber, though Kakkar et  al. (2011) reported that in cucumbers, most of the thrips were found in the blossoms, with few found elsewhere. In lettuce, however, leaf beating over a white tray was recommended for population monitoring on lettuce (Silva et  al., 2017). Pereira et  al. (2016) indicated that sequential sampling plans are more efficient than standard population sampling, especially at high thrips densities. Sticky traps can be used to capture adults in flight. Interestingly, females prefer red, whereas although males

are not strongly attracted to any one color, they seem to favor yellow (Yaku et al., 2007). Insecticides. Insecticides are often used as a means of population regulation. Insecticide classes should be rotated to reduce the rate of resistance development in treated populations of thrips. Cultural Practices. Because these thrips can transmit tospoviruses, crops should be initiated with virus-free planting material. Tall plants such as corn can be planted around susceptible crops to serve as a barrier to entry of dispersing thrips. The thrips can shelter there, so such crops should be treated periodically to eliminate the insects. Marigolds can serve as a trap crop. Weed elimination is helpful.

Florida Flower Thrips

Frankliniella bispinosa (Morgan) (Thysanoptera: Thripidae)

Natural History Distribution. Florida flower thrips is a native species that occurs commonly in Florida, southern Georgia, Alabama, and likely in adjacent areas. It is also found in Bermuda and the Bahamas. In Florida, it was formerly a problem only in southern Florida, but it has also caused damage in northern Florida in the recent years. Host Plants. Florida flower thrips is associated with a large number (at least 100) host plants, where it feeds mostly on blossoms and pollen. However, not all the plants fed upon by adults are also good reproductive hosts, so the truly important species are a subset of the host list. According to Oetting et al. (1993), F. bispinosa favors flowers with open structure, allowing ready access to stamens and pistils. Florida flower thrips is often very abundant on citrus, oaks, Quercus spp., and pines, Pinus spp., during the bloom period, but then most disperse, which is often when they become abundant in vegetable crops. In southern Florida they often are the dominant thrips in vegetable crop fields (Childers and Beshear, 1992), though when some of the new invaders such as melon thrips (Thrips palmi Karny) or blossom thrips (Frankliniella schultzei) are abundant the latter usually are more damaging. Among the crops infested are corn, cucumber, eggplant, pepper, tomatillo, tomato, watermelon, strawberry, blueberry, avocado, and peanut. Also supporting F. bispinosa are flower crops such as rose, snapdragon, gloxinia, chrysanthemum, iris, Persian violet, and baby’s breath (Frantz and Mellinger, 1990; Oetting et al., 1993). Trees, shrubs, and weeds infested by F. bispinosa include oaks; pines, pygmy date palm, Phoenix robelenii; trumpet tree, Tabebuia sp.; palmetto, Sabal sp.; elderberry, Sambucus sp.; nightshade, Physalis sp.; wild cherry, Prunus serotina; Spanish needles, Bidens pinosa; Aster sp.; and cattail, Typha domingensis (Childers, 1999).

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Natural Enemies. The natural enemies of Florida flower thrips are not well documented, although as with many other trips, the minute pirate bug Orius insidiosus (Say) (Hemiptera: Anthocoridae) is a very important predator. Funderburk et  al. (2000) and Ramachandran et  al. (2001) documented the importance of O. insidiosus in Florida. Life Cycle and Description. Like most vegetable-­ infesting thrips, the development of F. bispinosa includes the egg stage, then two larval instars, followed by the propupal and pupal instars, followed by the adult stage. This thrips is yellow in color. Generation time is estimated at 20–30 days (Tsai et al., 1996), so in southern Florida 12–15 generations are believed to occur annually. Apparently, both sexual and parthenogenetic reproduction can occur. Egg. The eggs are kidney-shaped, pale yellow, and approximately 0.4 mm long. Eggs are inserted into plant tissue. When fed cattail (Typha domingenisis) pollen, fecundity was an average of 187 eggs per female thrips. In contrast, mean fecundity was 85.5, 47.7, and 17.5 eggs per female when fed pollen from slash pine, Spanish needle, and pygmy date palm, respectively. According to Tsai et al. (1996), the incubation period of eggs averaged 4.9 days regardless of pollen source provided to females. Larva. The larvae are yellow and elongate oval in shape. They resemble adults but lack wings. The two larval instars are feeding stages. When cultured on cattail pollen, larval development time averaged 4.8  days; other pollens produced similar development time values (Tsai et al., 1996). For pupation, most larvae drop to the soil after the larval stage. Propupa and pupa. The propupal and pupal stages do not feed. They resemble adults but have wing buds instead of wings. The antenna is bend back over the head during the propupal stage but protrude forward during the pupal stage. The combined propupa and pupal development time averaged 4.1 days on cattail pollen, similar to other pollen diets.

FIG.  12.2  Head and thorax of Florida flower thrips. (Photo by J. Capinera.)

Adult. The body is elongate, and averages about 1.25 mm in length. The males tend to be slightly shorter than the average, and the females lightly longer. These thrips are yellow with brown setae, though the males are lighter in color than the females. Also, females often have gray bands or blotches on each segment of the abdomen. Both sexes are winged. The antennae are eight-segmented and bear stout, brown spines on the second segment. Newly emerged adults seek food, mate, and commence laying eggs in about 2.5 days. Average longevity of female F. bispinosa reared on cattail pollen was 31.1  days, similar to culture on Spanish needle and slash pine pollen. When reared on pygmy date palm, however, mean longevity was less, only 10.3 days. Keys to the genera of thrips can be found in Palmer et al. (1989) and Mound and Kibby (1998). Additional useful keys are presented by Oetting et  al. (1993), Hoddle et  al. (2012), Cluever and Smith (2017), and a key to common vegetable-infesting thrips in Appendix A.

Damage Larvae and adults of F. bispinosa will feed on buds, flower tissue, and leaves in addition to pollen. When densities are high, they may also feed on developing fruit. The young tissue (new growth) of the plant is most affected. Plant tissue fed upon by these insects may turn black or brown, shrivel, and drop prematurely. Discoloration of strawberry fruit is often referred to as “bronzing,” whereas in roses the petal discoloration is called “scarring.” These thrips will also oviposit in fruit, causing small dimples. Generally, this is not a problem unless the number of thrips is very high. Florida flower thrips are not as damaging as some other thrips species, in part because they are not efficient at transmission of plant disease-causing organisms. They can transmit tomato-spotted wild virus, but western flower thrips (Frankliniella occidentalis [Pergande]) is a much more efficient vector, so Florida flower thrips is of much less concern (Funderburk, 2009). Although western flower thrips is more efficient as a vector of plant pathogens, Florida flower thrips is better adapted to the high humidity levels typically found in the southeastern United States. Florida flower thrips is a good competitor with F. occidentalis (Paini et al., 2008) in this environment. Fecundity of F. bispinosus was greater, and the period of egg incubation is shorter, than those of F. occidentalis when cultured at high levels of relative humidity (70% and 85%) (Garrick et  al., 2016). During the periods of high humidity (typically summer and fall), Florida flower thrips become more abundant. The competitive ability of Florida flower thrips seems to suppress populations F. occidentalis at high humidities except when insecticides favor survival of insecticide-resistant F. occidentalis populations.

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Management These thrips usually are sampled by direct observation or by sampling blossoms and then flushing the thrips from the blossoms with alcohol. Insecticides are not normally applied for suppression of Florida flower thrips because the damage they cause is usually minor, they compete for food resources with more damaging pest such as western flower thrips, and they are a good source of food for predators that will also consume the more damaging species. Ramachandran et  al. (2001) reported that although these thrips may be abundant during the spring, by summer the native predators have suppressed their populations.

Grass Thrips

Anaphothrips obscurus (Müller) (Thysanoptera: Thripidae)

Natural History Distribution. This is the most common and widespread of the thrips species found on corn. It occurs throughout the United States and southern Canada. Host Plants. Among vegetable crops, only corn is affected. Other crops infested are barley, oat, timothy, and wheat. Numerous forage and weed grasses known to be suitable hosts include Agrostis spp., Arrhenatherusm sp., Avena sp., Bromus spp., Elymus spp., Festuca spp., Lolium sp., Panicum spp., and Poa spp. Natural Enemies. The natural enemies of grass thrips are poorly known because this minor pest has been studied infrequently. See the sections on onion thrips, Thrips tabaci Lindeman, or western flower thrips, Thrips occidentalis Pergande, for discussion of natural mortality factors. Life Cycle and Description. Adult females overwinter at the base of grass stems just above the soil. In the spring, as grass begins to grow, egg deposition begins. The number of annual generations is estimated at eight or nine in Massachusetts, with a generation requiring 2–4 weeks. The development is typical of vegetable crop-infesting thrips: the egg stage followed by larva I, larva II, propupa, pupa, and the adult. Egg. The eggs are deposited in the tender leaf tissue. The duration of the egg stage is 10–15 days during spring, but only about 7 days during summer. Females are thought to produce, on average, 50–60 eggs during their life span, but individuals produce over 200 eggs. The egg laying continues throughout spring and summer until freezes occur, but they do not overwinter successfully. Larva. The larvae are active, but tend to seek sheltered places to feed such as beneath leaf sheaths or within flowers. Larvae are long and slender in form and wingless. There are two larval instars. Larvae increase in size from about

0.25 mm at hatching to about 1.0 mm at maturity. Larvae resemble adults but have a smaller head, much narrower than the thorax. They also have shorter antennae. They are brownish or pinkish. Propupa and pupa. There are two pupal instars, the propupa (sometimes called prepupa) and pupa. They occur in a sheltered location, often at the base of a leaf sheath. The pupal stages are sluggish and do not feed. The antennae are folded back over the head during the propupal stage. The thorax bears wing cases that are long for pupae destined to produce winged adults, but short if giving rise to wingless individuals. Adult. Adults feed more openly than do larvae, often feeding on the leaves rather than within the leaf sheaths. Adults have both winged and wingless forms. Most overwintering females are wingless, but the winged forms soon predominate, accounting for about 90% of the thrips during the spring months. The proportions shift over the course of the summer, with wingless forms common late in the season, accounting for about 98% of the thrips. The winged individuals are larger, about 1.5 mm long, bear two pairs of fringed wings, and are brown. The wingless form is shorter, measuring about 1.0 mm long. The wingless forms are pink. The antennae of adults protrude forward, as in the larvae, but the head is relatively wide, nearly as wide as the thorax. Cary (1902) indicated that the males are infrequently found, with the females generally reproducing parthenogenetically. He also indicated that the males are not as slender as the females, the eyes are located more dorsally, and the tip of the abdomen bears paired copulatory structures that are absent from the females. In contrast, Palmer et al. (1989), Mound and Kibby (1998), and Mound and Marullo (1996) indicated that males were unknown. However, Hoddle et al. (2012) acknowledged the existence of males, reportedly in Iran. Females oviposit for a period of 4–6 weeks. The biology of grass thrips was given by Fernald and Hinds (1900) and Cary (1902), but important observations were made by Kamm (1972). A key to thrips genera was published by Palmer et al. (1989). This species is included in Mound and Marullo (1996), Mound and Kibby (1998), Hoddle et al. (2012), and in the key to common vegetableinfesting thrips in Appendix A.

Damage Larvae and adults puncture individual cells and remove the sap. This kills the cells, and imparts a silvery appearance to the tissue. The growing point or top of the plant is most often affected, usually while still immature, resulting in a condition called “silvertop.” This thrips is said to be highly mobile and very destructive to grasslands in Oregon (Kamm, 1971, 1972). However, grass thrips typically do not persist in corn, and cause little damage. Usually, thrips

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d­ isperse into young corn in June and within a generation they again disperse. Thrips injury to corn usually occurs only if the corn is also under moisture stress.

Management Management. Grass thrips tend to be a transient problem, and rarely require action. If corn is rapidly growing, recovery from thrips feeding is likely. Foliar insecticides are effective for suppression. Sampling. Reisig et  al. (2010) assessed the efficiency of different types of sampling methods for A. obscurus in timothy. Absolute sampling methods (per-unit area basis; e.g., beat cup) were very consistent. Relative estimates (e.g., sticky cards) are more affected by wind speed and thrips phenotype (relative abundance of winged and wingless forms). Cultural Practices. The presence of grasses, including small grains, may predispose a corn crop to infestation. Destruction of grasses in the autumn or winter may eliminate the overwintering stage. Corn cultivars differ in their susceptibility to grass thrips (Bing et al., 1990).

Melon Thrips

Thrips palmi Karny (Thysanoptera: Thripidae)

Natural History Distribution. Until the mid-1970s, the distribution of melon thrips was limited to Southeast Asia. In the recent years, it has spread throughout Asia, and to many Pacific Ocean islands, North Africa, Australia, Central and South America, and the Caribbean. In the United States, it was first observed in Hawaii in 1982, Puerto Rico in 1986, and Florida in 1990. It has the potential to infest greenhouse crops widely, but under field conditions melon thrips likely will be limited to tropical areas. Host Plants. Melon thrips is a polyphagous species, but is best known as a pest of Cucurbitaceae and Solanaceae. Among vegetables injured are bean, cabbage, cantaloupe, chilli, Chinese cabbage, cowpea, cucumber, bean, eggplant, lettuce, melon, okra, onion, pea, pepper, potato, pumpkin, squash, and watermelon. Tomato is reported to be a host in the Caribbean, but not in the United States or Japan. Tsai et al. (1995) reported that cucurbits were more suitable than eggplant, whereas pepper was less suitable than eggplant. In a survey of selected vegetable crops in southern Florida, T. palmi was abundant in blossoms of bean, squash, cucumber, and pepper, but absent in tomato (Kakkar et al., 2012). Other crops infested include avocado, carnation, chrysanthemum, citrus, cotton, hibiscus, mango, peach, plum, soybean, tobacco, and others. Natural Enemies. Natural enemies, particularly predators, are quite important in the ecology of melon thrips.

There is strong indication that melon thrips abundance and damage are increased by the application of some insecticides (Etienne et  al., 1990). Among the most important predators observed in Hawaii were the predatory thrips Franklinothrips vespiformis (Crawford) (Thysanoptera: Aeolothripidae) and especially the minute pirate bug, Orius insidiosus (Say) (Hemiptera: Anthocoridae). Other predators in Hawaii were Curinus coeruleus (Mulsant) (Coleoptera: Coccinellidae), Rhinacoa forticornis Reuter (Hemiptera: Miridae), and Paratriphleps laevisculus Champion (Hemiptera: Anthocoridae). Other predators and parasitoids are known in Asia (Hirose, 1991; Hirose et al., 1993; Kajita, 1986; Nagai, 1990). The parasitoid, Ceranisus menes Walker (Hymenoptera: Eulophidae), shows particular benefit in many Asian studies, and this wasp has been introduced to Florida (Castineiras et al., 1996a). Fungi known to affect melon thrips include Beauveria bassiana, Neozygites parvispora, Verticillium lecanii, and Hirsutella sp. and Lecanicillium muscarium (Verticillium lecanii) (Castineiras et  al., 1996b; North et  al., 2006). The entomopathogenic nematode Steinernema feltiae (Filipjev) (Steinernematidae) can induce mortality in T. palmi, though it is more effective on juvenile thrips than on adults (North et al., 2006). Life Cycle and Description. The development of melon thrips is typical of vegetable pest-infesting thrips: egg, larva I, larva II, propupa, pupa, and adult. A complete generation may be completed in about 20 days at 30°C, but it is lengthened to 80 days when the insects are cultured at 15°C. Melon thrips are able to multiply during any season that crops are cultivated but are favored by warm weather and suppressed by senescent crops. In southern Florida, they were damaging on both autumn and spring vegetable crops (Seal and Baranowski, 1992; Frantz et al., 1995). In Hawaii, they also became numerous on vegetables during the summer growing season (Johnson, 1986). Egg. The eggs are deposited in leaf tissue, in a slit cut by the female. One end of the egg protrudes slightly. The egg is colorless to pale white, and bean shaped in form. The duration of the egg stage is about 16 days at 15°C, 7.5 days at 26°C, and 4.3 days at 32°C. Larva. The larvae resemble the adults in general body form though they lack wings and have a smaller body size. There are two instars during the larval period. Larvae feed gregariously, particularly along the leaf midrib and veins, and usually on older leaves. Larval development time is determined principally by the suitability of temperature, but host-plant quality has also an influence. Larvae require about 14, 5, and 4  days to complete their development at 15°C, 26°C, and 32°C, respectively. At the completion of the larval instars, the insect usually descends to the soil or leaf litter, where it constructs a small earthen chamber for a pupation site.

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Propupa and pupa. There are two instars during the “pupal” period. The propupal instar is nearly inactive and pupal instar is inactive. Both instars are nonfeeding stages. The propupae (sometimes called prepupae) and pupae resemble the adults and larvae in form, except that they possess wing pads. The wing pads of the pupae are longer than that of the propupae. The combined propupal and pupal development time is about 12, 4, and 3 days at 15°C, 26°C, and 32°C, respectively. Adult. Adults are pale yellow or whitish, and lack dark pigmentation but bear numerous dark setae on the body. A black line, resulting from the juncture of the wings, runs along the back of the body. The population is heavily weighted toward females. The slender fringed wings are pale. The hairs or fringe on the anterior edge of the wing are considerably shorter than those on the posterior edge. The thrips measure 0.8–1.0 mm in body length, with females averaging slightly larger than males. Unlike the larval stage, the adults tend to feed on young growth, and so are found on new leaves. Adult longevity is 10–30 days for females and 7–20 days for males. The development time varies with temperature, with mean values of about 20, 17, and 12 days at 15°C, 26°C, and 32°C, respectively. Females produce up to about 200 eggs, but average about 50 per female. Both mated and virgin females deposit eggs. Careful examination is required to distinguish melon thrips from other common vegetable-infesting species. The Frankliniella spp. are easily separated from Thrips spp. because their antennae consist of eight segments, whereas in Thrips species there are seven antennal segments. To distinguish melon thrips from onion thrips, Thrips tabaci Lindeman, it is helpful to examine the ocelli. There are three ocelli on the top of the head, in a triangular formation. A pair of setae is located near this triangular formation, but unlike the arrangement found in onion thrips, the setae do not originate within the triangle. Also, the ocelli bear red pigment in melon thrips, whereas they are grayish in onion thrips. In general, the basic body color of adult melon thrips is yellow, but in onion thrips it is yellowish gray to brown. The geographic range of melon thrips in North America is quite restricted, so this also should aid in diagnosis. The most complete summary of melon thrips biology and management was presented by Girling (1992), although Cannon et al. (2007) provide an updated summary of management options. A detailed description of melon thrips is found in Bhatti (1980); Layland et al. (1994) also provided some diagnostic characters. Developmental biology was given by Tsai et al. (1995). Keys for identification of common thrips were presented by Palmer et al. (1989), Oetting et al. (1993), Mound and Marullo (1996), Mound and Kibby (1998), Hoddle et al. (2012), and Cluever and Smith (2017). Also, this species is included in a key to common vegetableinfesting thrips in Appendix A.

Damage Melon thrips cause severe injury to infested plants, particularly bean, eggplant, melon, and potato. Leaves become yellow, white, or brown, and then crinkle and die. Heavily infested fields sometimes acquire a bronze color. Damaged terminal growth may be discolored, stunted, and deformed. Densities from 1 to 10 per cucumber leaf have been considered to be the threshold for economic damage in some Japanese studies. However, studies in Hawaii suggested a damage threshold of 94 thrips per leaf early in the growth of the plant (Welter et al., 1990). Feeding usually occurs on foliage, but on pepper, a less suitable host, flowers are preferred to foliage. As the melon thrips prefer foliage, they are reported to be less damaging to cucumber fruit than western flower thrips, Frankliniella occidentalis (Pergande) (Rosenheim et  al., 1990). Nevertheless, fruits may also be damaged; scars, deformities, and abortion are reported. In Hawaii, thrips were observed to attain higher densities on cucumber plants infected with watermelon mosaic virus, but it was not determined whether the plants were more attractive to adults, or more suitable for survival and reproduction (Culliney, 1990). In addition to direct injury, melon thrips are capable of inflicting indirect injury by transmitting some strains of tomato-spotted wilt virus and bud necrosis virus.

Management Sampling. Larvae and adults are collected from foliage. Adults tend to move toward young foliage, with nymphs tending to be clustered on foliage inhabited by adults several days earlier. Adults can also be sampled with sticky and water pan traps. Blue and white are attractive colors for thrips, and have been used to trap melon thrips (Layland et al., 1994; Kawai and Kitamura, 1987). However, yellow has also been suggested to be an attractive color (Culliney, 1990). Insecticides. Foliar insecticides are frequently applied for thrips suppression, but at times it has been difficult to attain effective suppression. Various foliar and drench treatments, alone or combined with oil, have achieved some success (Seal and Baranowski, 1992; Seal et  al., 1993; Seal, 1994) though it is usually inadvisable to apply insecticides if predators are present. The eggs, which occur in the foliar tissue, and the pupae, which reside in the soil, are relatively insensitive to insecticide application. Cultural Techniques. Several cultural practices apparently affect melon thrips abundance, but few have been evaluated in the context of North American agriculture. Physical barriers such as fine mesh and row cover material can be used to restrict entry by thrips into greenhouses, and to reduce the rate of thrips settling on plants in the field (Kawai and Kitamura, 1987). High levels of atmospheric

Order Thysanoptera—Thrips Chapter | 12  591

carbon dioxide can be used along with elevated temperatures to disinfest produce and plants (Seki and Murai, 2012). Organic mulch is thought to interfere with the colonization of crops by winged thrips. Plastic mulch is also reported to limit population growth, but it is uncertain whether this is due to reduced rates of invasion or denial of suitable pupation sites. Crop stubble was not an effective deterrent (Litsinger and Ruhendi, 1984). The effects of intercropping potato with onion on melon thrips populations were evaluated by Potts and Gunadi (1991). Although aphid and aphid-borne disease incidence were decreased in such potato plantings, the density of thrips on potatoes was increased. Thus, the benefits of such cropping practices are largely a function of which pests are likely to be most important in an area. Heavy rainfall is thought to decrease thrips numbers (Etienne et al., 1990). However, there seems to be no evidence that overhead irrigation is an important factor in survival. Biological Control. The predatory mite Neoseiulus cucumeris (Oudemans) has been investigated for suppression of melon thrips (Castineiras et al., 1997). The mite density is correlated with thrips density, but within-plant distribution differs among the two species, suggesting that though the mites may increase in numerical abundance they are unlikely to drive the thrips to extinction. Host-Plant Resistance. Nuessly and Nagata (1995) reported that susceptibility to injury varied among pepper cultivars. They reported that though sweet and jalapeno types were sensitive to foliar injury, cubanelle and cayenne types produced acceptable size and quality fruit. This is the reverse of injury susceptibility to western flower thrips, so in areas with mixed thrips populations growers cannot rely solely on plant selection to avoid damage. Bean cultivars have also been shown to differ in susceptibility, but none are highly resistant (Cardona et al., 2002; Frei et al., 2003).

Onion Thrips

Thrips tabaci Lindeman (Thysanoptera: Thripidae)

Natural History Distribution. Onion thrips is believed to have originated near the eastern end of the Mediterranean Sea, or perhaps India. It was first observed in North America in 1872, and by the early 1900s had spread throughout the United States and southern Canada. It is easily transported on plant material, and redistribution of onion thrips occurs frequently with commercial shipment of bulbs and plants. It is now found throughout the world. Host Plants. Onion thrips has a wide host range, reportedly feeding on over 300 plants. In Hawaii, for example, 66  plants from 25 families were found to support onion

thrips (Sakimura, 1932). It has been found to infest such vegetables as asparagus, bean, beet, cabbage, cantaloupe, carrot, cauliflower, celery, cowpea, cucumber, garlic, kale, leek, mustard, onion, parsley, pea, pepper, pigeon pea, ­potato, pumpkin, spinach, squash, sweet potato, tomato, and turnip. Under field conditions, the most serious problems occur on onion, followed by cosmetic injury to cabbage and edible-podded pea. In greenhouse cultivation of vegetables, onion thrips sometimes cause severe injury to tomato and cucumber. Field crops such as alfalfa, cotton, oat, soybean, sugarbeet, wheat, and tobacco may also support onion thrips. Ornamental crops such as rose and carnation may be injured, especially when grown under greenhouse conditions. Many common weeds support onion thrips, including amaranth, Amaranthus palmeri; dandelion, Taraxacum ­officinale; mullein, Verbascum thapsus; goldenrod, Solidago canadensis; ragweed, Ambrosia spp.; kochia, Kochia scoparia; sage, Salvia sp.; sunflower, Helianthus annuus; smartweed, Polygonum spp.; and yellow nutgrass, Cyperus esculentus (Chittenden, 1919; Doederlein and Sites, 1993). Onion thrips is the most important insect pest of onion, and the dominant thrips species on onion. However, it is not the only species attacking onion, and in the southern states it is sometimes a relatively small component of the thrips fauna, being supplanted by western flower thrips, Frankliniella occidentalis (Pergande) (Bender and Morrison, 1989; Doederlein and Sites, 1993), and possibly by tobacco thrips, Frankliniella fusca (Hinds) (D.G. Riley, personal communication). Natural Enemies. No natural enemies of significance are known. Numerous species of lady beetles (Coleoptera: Coccinellidae), lacewings (Neuroptera: Chrysopidae), and flower flies (Diptera: Syrphidae) have been observed attacking onion thrips, but none regularly are effective enough to provide suppression. Insidious plant bug, Orius insidiosus (Say), and minute pirate bug, O. tristicolor (White) (both Hemiptera: Anthocoridae), are among the most effective of the predators, because their small size allows them to pursue the thrips between the closely appressed leaves of the onion plant, but these predators are rarely abundant enough to suppress thrips populations. Parasitoids have been introduced from Southeast Asia, where they parasitize a high proportion of onion thrips and other thrips species. In Hawaii, Ceranisus brui (Vuillet) (Hymenoptera: Eulophidae) was successfully introduced, but introductions of other Ceranisus spp. failed (Clausen, 1978). Weather. Weather is reported by many authors to be important in determining thrips abundance and damage. The combination of abnormally high temperature and low precipitation stimulates thrips reproduction and/or enhances survival. High temperature speeds up the life cycle, increasing the biotic potential of populations. Heavy rain is considered to be an important mortality factor, and is easy to observe large decreases in thrips abundance following

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s­ ignificant rainfall events (Harding, 1961), though the population decrease is temporary (Liu, 2004). Life Cycle and Description. The developmental stages of onion thrips are consistent with other vegetable pestinfesting thrips: the egg stage is followed by the two plantfeeding larval instars, then the nonfeeding propupa and pupa, and finally the adult. The life cycle of onion thrips is completed rapidly. Based on the field work conducted in Iowa, the development time from the egg to adult stages is estimated at only 10 days in July and 20 days in September. Similar values were obtained by Watts (1934) in South Carolina. Sakimura (1932) studied this insect in Hawaii, and observed longer developmental periods, but these studies were conducted during the relatively cool winter months. In any event, the potential number of generations is great under ideal conditions. The actual number of generations is estimated at 10 per year in southern climates but only two in the north. In temperate areas, overwintering occurs in the adult stage. Adults survive on winter wheat, alfalfa, clover, other crops, and weeds, but not on soil in the absence of living plants (Chambers and Sites, 1989). Some of these crops, particularly wheat, are also favorable for oviposition and larval development in the spring, and provide inoculum for nearby vegetables (Shirck, 1951; North and Shelton, 1986b). In warmer climates, nymphs may survive the winter, or reproduction may continue throughout the year. Egg. The female cuts slits in the foliage and deposits eggs just beneath the epidermis, with one end of the egg protruding slightly. The egg is colorless or yellowish white. It measures about 0.26 mm long and 0.12 mm wide, a size that is quite large considering the small size of the female. The egg is bean shaped, with one side being strongly concave, whereas the opposite side is strongly convex. The duration of the egg stage is estimated at 5–7 days when held at 21°C, but only 3 days at 25°C. Larva. The larvae of thrips resemble the adults in appearance, though the larvae are colorless or yellowish white. There are two instars. The first instar measures about 0.4 mm long, whereas the second instar measures about 0.9 mm long. The larvae can be differentiated by the tarsal arrangement—first instars bear tarsi with two claws, but these are absent from second instars. A less reliable, but more convenient, character for distinguishing the instars is their color; first instars tend to be colorless to white, whereas second instars tend to be yellowish. Both the first and second instars bear antennae with four segments. Larval duration varies from 3 to 11 days, with about 8–10 days required for the development at 21°C and 5–6 days required at 25°C. Watts (1934) reported the development times of 2.0 and 2.8 days, respectively, for the first and second instars. Larvae usually confine their feeding on

onion to the youngest foliage at the center of the plant, but are more widely distributed on cabbage. Propupa and pupa. After feeding for the first two instars, the larva almost always leaves the foliage and descends to the soil, where it constructs a small earthen chamber. Occasionally, it descends only to some vegetative structure such as a leaf axil. The active larval stages are followed by relatively inactive propupal (sometimes called prepupal) and pupal instars, which may last about 1.0 and 2.5 days, respectively, at warm temperature. This nonfeeding period requires about 5–7 days at 21°C, and 4–6 days at 25°C. These stages are colorless to yellow, and measure about 0.7 mm long. Setae are much more abundant on the propupal and pupal stages than in the larvae, and their wing pads are visible. The wing pads of the propupae extend back to the second abdominal segment. The wing pads of the pupae are considerably larger, extending back to the eighth abdominal segment. Antennae of the propupal stage project forward, whereas those of the pupal stage orient back over the thorax.

FIG. 12.3  Head and thorax of onion thrips. (Photo by J. Capinera.)

Adult. Females are about 1.0–1.2 mm long, and yellowish or yellowish-brown. Patches of dark brown occur on the thorax and abdomen. The overall body color is lighter in the summer and darker in the winter. The wings are colorless, and extend back to about the sixth abdominal segment. Male onion thrips are slightly smaller than females, and have nine abdominal segments instead of the 10 found in females. A preoviposition period in adults of about 3 days was noted by Watts (1934), but it was estimated at 5–7 days by Sakimura (1932). Adult females live about a month, and typically deposit an average of 30–40 eggs (maximum of about 100 eggs) at 1.5–2.5 per day. The adults do not occur in an equal sex ratio. Male thrips are relatively rare, and seem to be more plentiful in the autumn (Sakimura, 1932) and in the western hemisphere (Kendall and Capinera, 1990). Females generally reproduce parthenogenetically. There are three modes of

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p­ arthenogenetic reproduction found in onion thrips: thelytoky (unfertilized eggs develop into females), arrhenotoky (unfertilized eggs develop into males and fertilized eggs develop into females), and deuterotoky (unfertilized eggs develop into either males or females) (Nault et al., 2006). The type of reproduction in a given location can change over the course of the season and from year to year. Differentiation of onion thrips from other common ­vegetable-infesting thrips requires close examination. The adult female of onion thrips has seven antennal segments, a character that is useful to distinguish this species from co-occurring Frankliniella thrips, have eight antennal segments. To distinguish onion thrips from melon thrips, which also has seven antennal segments, it is helpful to examine the ocelli. There are three ocelli on the top of the head, in a triangular formation. In onion thrips, a pair of setae originate from within this triangular formation, unlike the arrangement found in melon thrips, where the setae do not originate within the triangle. The ocelli also tend to be gray in onion thrips, but bear red pigment in melon thrips. Although the body color of onion thrips is quite variable, some gray or brown is usually present on the body of adults in addition to yellow, whereas in melon thrips the body is uniformly yellow. Melon thrips has a very restricted geographic range, and so should infrequently be confused with onion thrips. Good accounts of onion thrips biology were given by Chittenden (1919), Horsfall and Fenton (1922), and by Gill et  al. (2015). Developmental parameters were given by Sakimura (1932), Watts (1934), and Edelson and Magaro (1988). Quaintance (1898b) provided a useful morphological description. Keys for identification of common thrips are given by Palmer et al. (1989), Mound and Marullo (1996), Mound and Kibby (1998), Hoddle et al. (2012), and Cluever and Smith (2017). This species is also included in a key to common vegetable-infesting thrips in Appendix A.

Damage The principal form of damage caused by onion thrips results from the piercing of cells and removal of cell contents by larvae and adults. In onions, this leads to an irregular or blotchy whitening of the leaves, a condition sometimes termed “blast.” Heavy levels of feeding injury disrupt the hormonal balance of the plant, causing the leaves to curl and twist, and the foliage to be stunted (Kendall and Bjostad, 1990). Such damage decreases onion bulb size, and may even lead to death of the plant. Silvering or whitening of the pods on edible-podded peas is also attributed to onion thrips (Shelton and North, 1987). On cabbage, feeding by thrips causes a bronze discoloration and rough texture, and the cabbage heads may fail fresh market standards (North and Shelton, 1986a). However, Liu and Spark (2003) reported that most of the feeding on red cabbage occurs on the outer five leaves, which can be removed easily from large heads of cabbage.

Crops such as cabbage may also fail processing standards due to contamination of products such as sauerkraut with thrips bodies (Shelton et al., 1982). As happens with onion, the thrips feed in sheltered locations, here between the leaves that form the cabbage head. The tips of asparagus spears may be heavily infested with thrips following dispersal from nearby weeds or crops. Careful management of weeds and crop borders can alleviate such problems (Banham, 1968). The relationship between thrips numbers and onion yield has been the subject of much study. Although Mayer et  al. (1987) found no relationship between thrips abundance and yield of dry onions in Washington, studies in Colorado (Kendall and Capinera, 1987), Texas (Edelson et  al., 1986, 1989), and Quebec (Fournier et  al., 1995) demonstrated yield decreases associated with thrips feeding. For thrips injury to be significant, feeding must occur during the mid-season period of rapid bulb expansion; early- and late-season feeding has little or no effect on yield. Also, there is a threshold effect. Thrips densities of 1–2 per leaf or 30 per plant generally must be reached for injury to occur. To complicate matters further, however, onion responses are modified somewhat by weather, particularly moisture, and by onion variety. Sweet onions are particularly susceptible to bulb size reduction. Unlike the situation with storage onions, there is virtually no tolerance of thrips on scallions (green onions) because the tops as well as the developing bulb are marketed and consumed (Kawate and Coughlin, 1995). Onion thrips may also affect plant disease incidence. The fungus Alternaria porri causes a foliar disease of onions called purple blotch. Although the fungus does not depend on thrips for transport or inoculation, feeding wounds can serve as a penetration site for the purple blotch fungus, and disease incidence is increased in the presence of thrips (McKenzie et al., 1993). Onion thrips is also implicated in the transmission of tomato-spotted wilt virus to several vegetable crops (Greenough and Black, 1990). Tomato-spotted wilt virus is acquired by thrips during the larval stage, but the insects remain infected and capable of transmitting the virus for the duration of their life (Sakimura, 1963). Recently, onion thrips has been implicated in transmission of Pantoea ananatis (Serrano), the bacterium that causes center rot of onion (Grode et al., 2017).

Management Sampling. Edelson et  al. (1986) studied the dispersion of thrips among onion plants and reported a clumped distribution. The distribution within plants is also nonuniform. Thrips densities are normally determined by visual examination of plants. Although there is variation among counts attributable to different observers (Theunissen and Legutowska, 1992), there is a strong correlation between

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visual estimates and actual population densities (Edelson, 1985a; Liu and Chu, 2004). Onion thrips is normally found at the basal area of onion leaves or in the leaf folds, and it is necessary to pull the leaves apart slightly to observe the entire population. This is especially true early in the season and early in thrips development, while later the thrips are more prone to move apically. Sunny weather, maturity, and the need for flight are reputed to account for the change in thrips distribution (Sites et al., 1992). Sequential sampling (Shelton et al., 1987) and binomial (the presence-absence) sampling (Fournier et  al., 1994) plans have been developed to reduce the effort associated with thrips sampling on onion. Thrips are active and are readily dispersed by wind when they fly. Because thrips readily disperse from crop to crop, it is useful to employ sticky traps to monitor thrips flights, or to monitor their densities in crops that provide potential inoculum. White- or yellow-sticky traps can be used for monitoring thrips in flight, and thrips on foliage can also be sampled by using heat to drive than from the foliage, as with a Berlese funnel (Shelton and North, 1986; Doederlein and Sites, 1993). Onion thrips are active fliers during the daylight hours, but not at night. Also they are active between 20°C and 28°C, but not above 30°C or when the windspeed exceeds 3.8 m/s (13.7 kph) (Smith et al., 2016). Insecticides. Insecticides are frequently used for thrips suppression, especially during the period of rapid bulb expansion when plants are most susceptible to injury, or late in the season when thrips have the potential to reach very high levels of abundance and feeding injury is obvious. Foliar applications are made as frequently as twice per week in commercial onion production. It is difficult to obtain an excellent control because of the cryptic nature of thrips feeding. Many onion and cabbage producers, concerned that they will be unable to eliminate thrips from sheltered feeding locations, apply insecticides as a preventive measure, in advance of potential problem development. Especially for cabbage, once the leaves begin to cup and form a head, effective insect control is difficult. Insecticides that produce toxic fumes are desirable, because they penetrate into crevices where thrips hide, but these materials are limited in availability. On occasion, insecticides are applied to the soil or plastic mulch beneath plants because the thrips descend to the soil to pupate, where they contact the insecticide (Pickford, 1984). Some, but not all, systemic insecticides effectively suppress thrips for at least part of the season (Getzin, 1973; Sinha et al., 1984). Insecticide resistance occurs in many locations. Biological Control. Introduction of exotic eulophid parasitoids has had some success (see above, natural enemies), but this approach to biological control has not been fully exploited. Release of predators that are easily cultured, particularly lacewings (Neuroptera: Chrysopidae), has not been very successful in the field. Predatory mites (Acari:

Phytoseiidae) have been found to provide suppression of onion thrips in greenhouse culture (Bakker and Sabelis, 1989), as have releases of Orius spp. (Hemiptera: Anthocoridae). It is important to release adequate numbers of predators at the first sign of thrips infestations. Cultural Practices. Crop management can influence the nature of thrips injury in several ways. For example, the proximity of susceptible crops to thrips sources is important. Damage sometimes occurs when thrips disperse in large numbers into susceptible crops. This often results when an early-season crop such as oats or wheat reaches maturity, or when a crop is cut at mid-season, as is the case with alfalfa and clover (Banham, 1968; Shelton and North, 1986). Mulches can influence the abundance of thrips and the transmission of plant viruses. In studies conducted in Louisiana, aluminum-surfaced mulch reduced the incidence of tomato-spotted wilt transmission by thrips to tomato and pepper by about 60%–80% (Greenough and Black, 1990). This approach toward disease management has been studied much more with aphid vectors (see section on Melon Aphid), but most of this technology is probably applicable to thrips-transmitted diseases. Intercropping can have some benefit for onion thrips management. Despite its wide host range, there are clearly preferred hosts, principally onion. For example, Uvah and Coaker (1984) alternated rows of onion with various ratios of carrot rows, and found that the presence of carrots decreased abundance of thrips. This occurred despite the fact that carrot is a nominal host of thrips. Sanitation is very important. Long ago, Horsfall and Fenton (1922) noted the ability of thrips to disperse from contaminated overwintering plants left in the field, or from transplanted onion bulbs taken from storage, to newly seeded onions. Soil and weeds also harbor onion thrips during the winter months (Larentzaki et al., 2007). With the increased availability of rapid transportation, thrips are often moved with plant material, and then inadvertently inoculated into fields. For example, Schwartz et al. (1988) found that nearly all batches of onion transplants shipped from Texas to Colorado were contaminated with thrips. Sporadic incidence of insecticide resistance among Colorado onion fields apparently was related to different sources of onions and thrips, and different pesticide exposure histories. Also, in some northern areas, greenhouses are a source of thrips in the spring. High levels of atmospheric carbon dioxide can be used along with elevated temperatures to disinfest produce and plants (Seki and Murai, 2012). Host-Plant Resistance. The cryptic nature of thrips feeding on onion has long made chemical control difficult, and has stimulated the search for resistant varieties. Characteristics associated with resistance are round leaves and open or spreading plant architecture, attributes sometimes found in white onion varieties. It has been speculated

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that this plant architecture affords less opportunity for thrips to hide between leaves, hastening their predation by other insects (Jones et al., 1935; Coudriet et al., 1979). However, differences in plant chemistry have also been suggested (Saxena, 1975) to account for this difference. Indeed, differences in the ratio of adult and larval thrips among onion varieties (Coudriet et al., 1979) could indicate physiological differences in suitability for thrips growth and survival. Some studies, however, report that the basis for resistance to onion thrips is feeding tolerance by some onion cultivars. Resistance has also been identified among cabbage varieties (Shelton et  al., 1983a, 1988, 1998; Hoy and Kretchman, 1991), but the basis for resistance is uncertain, and no clear patterns have emerged that would allow prediction of resistant types of cabbage. It is interesting to note, however, that some “resistant” cultivars support as many thrips as the susceptible cultivars, but on resistant plants the thrips feed principally on the outer leaves that are discarded at harvest, thus causing little damage (Stoner and Shelton, 1988). Whereas insecticide treatments alone sometimes fail to keep thrips from damaging thrips injury-susceptible cabbage in New York, when insecticides are used with varieties moderately susceptible to thrips injury, the combination is effective at preventing damage (Shelton et al., 1998).

Tobacco Thrips

Frankliniella fusca (Hinds) (Thysanoptera: Thripidae)

Natural History Distribution. Tobacco thrips is widely distributed in eastern Canada and the United States, west to about the Rocky Mountains. However, it is most abundant, and most often recorded as a pest, in the southeastern states. It is a native species. Host Plants. Vegetable hosts include bean, beet, cantaloupe, carrot, corn, cowpea, cucumber, onion, pea, pepper, potato, tomato, and watermelon. Because tobacco thrips can vector tomato-spotted wilt virus, among vegetable crops it is known principally as a pest of tomato. Also, note that in a survey of vegetable crops conducted in southern Florida, F. fusca was found in tomato, but absent in cucumber, pepper, bean, and squash (Kakkar et  al., 2012). Tobacco thrips is better known as a field crop-infesting insect, infesting alfalfa, barley several types of clover, cotton, lespedeza, peanut, rye, tobacco, vetch, wheat, and occasionally corn and oats. Winter grains such as rye and wheat, and volunteer peanut, apparently are suitable overwintering hosts. Several weeds have been reported to support tobacco thrips, such as Bermudagrass, Cynodon dactylon; blue toadflax, Linaria canadensis; broomsedge, Andropogon virginicus; buttercup, Ranunculus sp.; cocklebur, Xanthium sp.; crabgrass, Digitaria sp.; cutleaf evening primrose, Oenothera laciniata;

dandelion, Taraxacum officinale; dog fennel, Eupatorium capillifolium; false dandelion, Pyrrhopappus carolinianus; feathergrass, Leptochloa filiformis; Johnsongrass, Sorghum halepense; little barley, Hordeum pusillum; rabbit tobacco, Gnaphalium obtusifolium; sand blackberry, Rubus cuneifolius; shepherdspurse, Capsella bursa-pastoris; spiny sow thistle, Sonchus asper; wild lettuce, Lactuca sp.; wild radish, Raphanus raphanistrum; wood sorrel, Oxalis spp.; and a grass, Brachiaria extensa. Natural Enemies. The natural enemies of tobacco thrips have not been well documented, but likely are the same as those associated with western flower thrips, Frankliniella occidentalis. A nematode, Thripenema fuscum (Tylenchida: Allantonematidae), was observed to parasitize up to 68% of thrips in Florida, suggesting that this may be an important mortality factor in some cropping systems (Tipping et al., 1998). As percent parasitism of thrips increased, the number of F. fusca in terminals and buds of peanut decreased (Funderburk et al., 2002). Insidious flower bugs, Orius insidiosus (Say) (Hemiptera: Anthocoridae), also have been observed to be important, and heavy rainfall is detrimental. Life Cycle and Description. Tobacco thrips display the developmental pattern typically found among vegetable crop-infesting thrips pests: egg, larva I, larva II, propupa, pupa, and adult. The propupa and pupa do not feed. Tobacco thrips tend to be abundant in a crop during the spring and summer (McPherson et  al., 1992). In Florida, tomato blossoms are infested during April-June (Salguero Navas et al., 1991a), but the thrips are abundant later further north. Several generations are present annually in Florida, including about three during the winter months (Toapanta et  al., 1996). Eddy and Livingstone (1931) reported five generations annually from South Carolina. Unlike the situation in Florida, where reproduction occurs during the winter months, in Georgia, South Carolina, and Louisiana overwintering occurs only in the adult form. The life cycle requires about 15–21 days for completion. Egg. The egg is inserted into the foliage with an end protruding slightly. The egg measures about 0.25 mm long, is white, and bean shaped. Mean duration of the egg stage is 6.7 days (range 3–10 days). Larva. The first instar measures about 0.23 mm long, the second instar 0.60–1.17 mm. Both instars are yellowish or whitish. Mean duration of the first larval instar is about 1.1 and 1.0  days at 25°C and 35°C, respectively. Mean duration of the second larval instar is about 4.7 and 2.6 days at 25°C and 35°C, respectively. Larvae tend to feed in cryptic habitats such as blossoms and terminal growth, and rarely in open, exposed areas. Propupa and pupa. At the completion of the larval instars, the mature larva drops to the soil to pupate. The ­propupa

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(sometimes called prepupa) is 0.5–0.6 mm long, the pupa 0.6–1.2 mm. The color of these latter instars is yellow, but unlike the larval instars they display wing pads. Mean duration of the propupa is about 1.1 and 0.8 days at 25°C and 35°C, respectively. Mean duration of the pupa is about 1.4 days at both 25°C and 35°C.

thrips the hairs at the center of the posterior edge are shorter or absent. Tobacco thrips is distinguished from common blossom thrips by the length of the anteromarginal setae: in tobacco thrips they are distinctly shorter than the anteroangular setae, whereas in blossom thrips they are only slightly shorter. The biology of tobacco thrips was presented by Hooker (1907) and Eddy and Livingstone (1931), but additional observations were made by Newsom et al. (1953), Chamberlin et  al. (1992), and Chellemi et  al. (1994). The developmental data were given by Watts (1934) and Lowry et  al. (1992). Culture methods were described by Kinzer et  al. (1972). Keys for identification were included by Palmer et al. (1989) and Oetting et al. (1993), Mound and Marullo (1996), Hoddle et al. (2012), and Cluever and Smith (2017). Also, this species is included in a key to common vegetableinfesting thrips in Appendix A.

Damage FIG. 12.4  Head and thorax of tobacco thrips. (Photo by J. Capinera.)

Adult. The adult stage is variable but often has two pairs of fringed wings. The fringe on the anterior edge of the wings is markedly shorter than the fringe on the posterior edge. The adults measure 1.0–1.3 mm long. The head and thorax are light brown or yellowish brown, but the abdomen is dark brown. The antennae consist of eight segments. Long-winged (macropterous) and short-winged (brachypterous) forms occur, with brachypterous forms dominant during the winter months. The wings of the macropterous form usually do not attain the tip of the abdomen, but wing length is quite variable. Wings of the brachypterous forms barely extend to the first abdominal segment. Feeding behavior of the adults is similar to that of the larvae. The mean preoviposition period is about 2.6  days (range 2–6  days). Lowry et  al. (1992) reported adult female longevity of about 6–10 days, and fecundity of 13–24 eggs per female, but these values seemed small and might reflect a poor host or suboptimal rearing conditions. When fed cotton, mean longevity of mated females is 27–47  days and fecundity about 20–60 eggs per female (Eddy and Livingstone, 1931). Fertilized females produce both female and male offspring, though females are favored; unfertilized females produce only males. Distinguishing tobacco thrips from other vegetableinfesting thrips requires careful examination. Antennal structure can be used to separate the Thrips spp. because their antennae consist of seven segments, whereas in Frankliniella there are eight segments. Separation of western flower thrips from tobacco thrips is accomplished by examining the eighth dorsal plate on the abdomen. In western flower thrips, there is row of short hairs of approximately equal length along the posterior edge, whereas in tobacco

Vegetable seedlings can be damaged by this thrips when they disperse to young annual crops from maturing perennial crops such as alfalfa or clover. Thrips feed and deposit eggs into the young tissue, causing young leaves to curl upward and older leaves to acquire a silvery or speckled, and crinkled, appearance (Webb, 1995). Buds and other young tissue may be killed, giving the seedling a scorched or burnt appearance. Destruction of terminal growth may disrupt apical dominance, producing an excessively bushy, branched growth form. Tobacco thrips may be found in blossoms, but unlike its co-occurring species western flower thrips, it is primarily a leaf feeder. Direct feeding injury has been studied best in peanuts, where insect suppression has been shown to increase yields slightly or not at all (Tappan and Gorbet, 1981; Tappan, 1986; Lynch et al., 1984a). Similarly, direct injury to vegetables is infrequent, but because tobacco thrips now transmits tomato-spotted wilt virus, its importance as a vegetable pest has escalated greatly (Frantz and Mellinger, 1990). As happens with western flower thrips, virus acquisition occurs in the larval stage. After a latent period of 4–18 days, adults remain capable of transmitting tomato-spotted wilt throughout their life (Sakimura, 1963). The temporal occurrence of both thrips and tomato-spotted wilt varies among locations and years (Groves et al., 2003). Weeds are important in the overwintering of both the thrips and virus (Hobbs et al., 1993; Johnson et al., 1995).

Management Sampling. Distribution of tobacco thrips in tomato blossoms was studied by Salguero Navas et  al. (1994). Populations were aggregated or randomly dispersed, varying with year of sampling. A binomial (the presence or the absence) sampling protocol was developed and shown to be useful for populations with less than an average of two

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thrips per blossom. These authors recommended collecting 16–18 flowers to assess the need for population suppression. Washing of plant material with a dilute sodium hypochlorite and soap solution produced higher thrips estimates than visual searches (Burris et al., 1990). Management. Management of this species is normally accomplished in conjunction with western flower thrips suppression because Frankliniella occidentalis is generally more numerous and damaging (see the discussion under western flower thrips for more detail), they are similar in ecology, and because they occur together in mixed populations. Other thrips species, particularly flower thrips, Frankliniella tritici (Fitch), also may be present, but because they are not an important vectors of tomato-spotted wilt virus F. tritici are considered to be of little importance. Reflective mulches help to deter invasion of susceptible crops by dispersing thrips. Some of the benefit can be lost as the canopy of the plants expands, but there is considerable benefit to be gained by delaying the inoculation of plant pathogens by thrips. For example, Riley and Pappu (2004) documented yield benefit associated with use of reflective mulch, and the additive effects of using reflective mulch plus resistant plants or thrips-effective insecticide treatments.

Western Flower Thrips

Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae)

Natural History Distribution. Formerly restricted mostly to the western United States and Canada, by 1980 this native thrips had spread east to Georgia. Subsequently, it has spread throughout the United States and into southern Canada, and to other continents. It has also become a serious pest in Hawaii. Range expansion has undoubtedly been enhanced by movement of ornamental plants and vegetable seedlings from southern nurseries. It survives best in warm climates, and overwinters outdoors on growing plants along the west coast and throughout the southeastern states. Normally, it is not thought to overwinter in very cold climates, but to reinvade these areas annually from greenhouses, or via introduction of seedlings from southern areas. However, the report of overwintering in Pennsylvania under leaf debris and in bare soil (Felland et al., 1993) suggests a significant degree of cold hardiness. Host Plants. Western flower thrips apparently has an exceedingly wide host range. However, plant suitability varies seasonally and even geographically. Also, from an economic perspective, the most important hosts are those that support both thrips reproduction and virus disease multiplication. Western flower thrips occurs on several

vegetable crops, including cucumber, onion, pepper, potato, lettuce, and tomato. In a survey conducted in southern Florida, F. occidentalis was found on cucumber, bean, squash, and tomato, but absent in pepper (Kakkar et  al., 2012). Tomato is most seriously injured directly by the thrips, through oviposition, but both lettuce and tomato are seriously damaged by tomato-spotted wilt virus transmitted by thrips. Under greenhouse conditions, cucumber and pepper are also readily damaged. Field crops on which western flower thrips occurs include alfalfa, canola, crimson and white clover, millet, peanut, rye, vetch, and wheat. Several fruit crops have been reported to serve as hosts, such as apple, blackberry, blueberry, peach, pear, and plum. Among the weeds that serve as good hosts are such common species as black nightshade, Solanum nigrum; cheese weed, Malva palviflora; daisy fleabane, Erigeron annuus; dandelion, Taraxacum officinale; false dandelion, Pyrrhopappus carolinianus; jimson weed, Datura stramonium; galinsoga, Galinsoga parciflora; lambsquarters, Chenopodium album; lantana, Lantana camara; pigweed, Amaranthus spp.; prickly lettuce, Lactuca serriola; sorrel, Oxalis spp.; sow thistle, Sonchus oleraceus; and wild radish, Raphanus raphanistrum (Stewart et  al., 1989; Yudin et  al., 1986; Chamberlin et  al., 1992; Bautista and Mau, 1994; Chellemi et al., 1994), but numerous other species can also serve as hosts. In Hawaii, the blossoms of woody legumes growing near cultivated fields serve as a major source of thrips. Natural Enemies. Considering the abundance of western flower thrips and the severity of their injury to plants, surprisingly little is known about natural enemies. Minute pirate bugs, particularly Orius tristicolor (White) and O. insidiosus (Say), feed voraciously on western flower thrips, and there is good evidence that they suppress thrips populations in vegetable crops (Salas-Aguilar and Ehler, 1977; Letourneau and Altieri, 1983; Funderburk et  al., 2000). Ceranisus spp. (Hymenoptera: Eulophidae) parasitize the immature stages, but are not generally abundant. Fungal epizootics caused by Lecanicillium (Verticillium) and Entomophthora have been observed in moist climates. Nematodes in the genus, Thripenema (Howardula) (Nematoda: Allantonematidae), appear to be frequent associates of western flower thrips (Wilson and Cooley, 1972; Heinz et  al., 1996), and reduce feeding by, and induce sterility in, their hosts (Poinar, 1979; Arthurs and Heinz, 2003). Impact of these nematodes has been inadequately studied, but an incidence of 88% has been reported from California (Heinz et al., 1996). Life Cycle and Description. This species displays the developmental pattern typically seen in vegetable cropinfesting thrips: egg, larva I, larva II, propupa, pupa, and adult. In mild climates, these thrips readily overwinter as adults and nymphs on many crops and weeds. However,

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in relatively cold climates such as northern Texas they overwinter on hardy crops such as alfalfa and winter wheat (Chambers and Sites, 1989). As noted above, they apparently can also survive in leaf debris and soil under the cold-weather conditions of Pennsylvania (Felland et  al., 1993). Toapanta et  al. (1996) estimated three to five generations per year in north Florida, with populations highest in spring and a smaller peak in autumn. They can complete one generation in 15 days, so under ideal conditions many more generations are possible. A temperature of about 30°C seems to be optimal for population growth. FIG. 12.5  Head and thorax of western flower thrips. (Photo by L. Buss.)

Egg. The eggs are deposited in young vegetative tissue, with one end protruding slightly. They are shaped like a bean, white in color, and measure about 0.25 mm long. Duration of the egg stage is reported to require 5–15 days in the field, but mean duration is only 2.6  days at 25°C. Females were reported by Gaum et  al. (1994) to produce about 9–10 eggs during their life span when cultured on cucumber, and Lowry et al. (1992) reported that fecundity was 14–23 eggs per female when fed peanut. Because western flower thrips have preferred hosts, and usually include pollen in their diet, the aforementioned estimates of egg production may be artificially low. Thus, Lublinkhof and Foster (1977), recorded 43–95 eggs per female cultured on bean, and Trichilo and Leigh (1988) observed fecundity of about 130 eggs per female on cotton but 190 eggs on cotton supplemented with pollen. Larva. The development of the immature thrips requires two larval instars (also sometimes called nymphs), which are feeding stages, followed by nonfeeding propupal and pupal stages. The temperature threshold for larval development is about 9.4°C. Larval development time may require 9–12  days in the field, extending to 60  days during the winter. However, when reared at a constant temperature of 25°C, the first and second larval instars require only 2.3 and 3.7 days, respectively. Larvae and adults are somewhat gregarious, often feeding together in small groups. At maturity, the larvae drop to the ground to pupate. Propupa and pupa. The propupal and pupal stages are reported to require 1–3 and about 3–10 days, respectively, under field conditions. At a constant temperature of 25°C, however, thrips complete their pupal development in an average of 1.1 and 2.7  days, respectively. The propupa is distinguished by the presence of short-wing pads and erect antennae. The pupae have long wing pads that reach almost to the tip of the abdomen, and antennae that are bent backward along the head. Propupae and pupae may be found on the surface, under debris, or in cracks and crevices to a depth of 7–10 cm.

Adult. The adults have fully formed, fringed wings and measure 1.2–1.9 mm long, averaging 1.5 mm. The fringe along the anterior edge is markedly shorter than the posterior edge. Body color varies from yellow to brown. The antennae bear eight segments. The adults commonly live about 20–30 days, but some persist providing 40–70 days. Females may mate, or reproduce parthenogenetically. Unmated females produce only males, whereas mated females produce both sexes. The offspring of mated females are female biased, usually on the order of 2:1. Females mate immediately upon emergence, and repeatedly over the course of their life. Dispersal by adults usually occurs when their food plants become unsuitable, which commonly results from drought, maturity, or harvesting. Distinguishing western flower thrips from other vegetable-infesting thrips requires careful examination. ­ Antennal structure can be used to separate the Thrips spp., because their antennae consist of seven segments, whereas the antennae in Frankliniella bear eight segments. Separation of western flower thrips from tobacco thrips, Frankliniella fusca (Hinds), is accomplished by examining the eighth dorsal plate on the abdomen. In western flower thrips, there is row of short hairs of approximately equal length along the posterior edge, whereas in tobacco thrips the hairs at the posterior edge of the plate are shorter or absent centrally. Both nymphs and adults produce an alarm pheromone, and respond to it by moving away from the source of the pheromone, and usually by dropping from the plant. The pheromone is released in droplets of anal fluid (Teerling et al., 1993). Thrips biology was given by Bailey (1933b), Bryan and Smith (1956), Lublinkhof and Foster (1977), Gaum et  al. (1994), and van Rijn et  al. (1995). Rearing techniques were described by Teulon (1992) and Doane et al. (1995). Keys that included western flower thrips were presented by Palmer et  al. (1989), Mound and Marullo (1996), Oetting et al. (1993), Hoddle et al. (2012), and Cluever and Smith (2017). Also, this species is included in a key to common

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vegetable-infesting thrips in Appendix A. Useful s­ ummaries of pest management practices and potentials in different cropping systems and geographic locations were published by Demirozer et al. (2012) and Mouden et al. (2017).

Damage This species, as its common name suggests, prefers an interstitial habitat such as within flowers or in leaf clusters; only rarely is it found in exposed locations. It typically feeds on pollen grains and on the ovary of flowers, resulting in malformed, stunted, or discolored fruit. In cucumber, for example, western flower thrips feeding causes silvery, web, or streak-like scarring, which may be accompanied by fruit malformation (Rosenheim et al., 1990). When they feed on foliage, they cause distortion of expanding leaves and mottling or speckling of mature leaves. On onion, their feeding injury is similar to the effects of feeding by onion thrips, Thrips tabaci Lindeman. Thrips also deposit eggs in small fruits, inducing deformities. In the absence of flowering plants, however, they oviposit readily on such plants as nonflowering lettuce. Salguero Navas et  al. (1991a) documented the damage to tomato fruit caused by oviposition, typically reflected by a dimple or indentation surrounded by a light-colored halo. Western flower thrips was much more damaging than some other tomato-infesting thrips at comparable densities. In southern Texas, western flower thrips has been reported to damage onions along with onion thrips (Bender and Morrison, 1989). These thrips are also found associated with onion blossoms, where they enhance pollination and seed set. Only at very high densities, approximately 9000– 10,000 thrips per onion seed head, is damage likely to occur (Carlson, 1964). Shipp et al. (1998, 2000) calculated economic injury levels for greenhouse-grown sweet peppers and cucumber. There is a strong association between the prevalence of western flower thrips and tomato-spotted wilt virus (Riley and Pappu, 2004. In this thrips species, oviposition preference is as important as feeding preference, because only nymphs are capable of virus acquisition. Thus, selection of oviposition sites by females determines the likelihood of the thrips developing into a virus vector. Once infected, the thrips remain capable of transmitting the virus for the remainder of their life. Tomato-spotted wilt virus-infected weeds are the major host of virus in vegetable fields unless susceptible crops are cultivated continuously (Cho et  al., 1986, 1987). Western flower thrips has become a particularly serious pest of vegetable and ornamental greenhouse crops. Its short development time, wide host range, cryptic feeding habits, and particularly its tendency to evolve insecticide resistance rapidly, make it well suited for inhabiting commercial greenhouses. In high-value crops or in crops with a

high threat of virus transmission, there is virtually no tolerance for this insect (Mouden et al., 2017). Interestingly, Joost and Riley (2005) showed that F. occidentalis was not much affected (as judged by settling and probing behavior) by the insecticide imidacloprid, whereas F. fusca (Hinds) was negatively affected as judged by these same behaviors. Despite the severity of the western flower thrips virus disease problem on some crops, these thrips are not entirely detrimental. Thrips also feed on mites, and serve as important alternate hosts for some larger predators. Thus, in cropping systems that are not particularly susceptible to thrips or virus injury, the presence of low to moderate numbers of thrips can be beneficial (Gonzalez et al., 1995a).

Management Sampling. Thrips densities in blossom samples are often made in the field by visual examination of the plant or by shaking the blossom or other vegetative material over a tray, but the precision of this type of population estimate is quite low. A better estimate is gained by submerging the plant sample in 70% ethanol, or in sodium hypochlorite and soap solution, and shaking it to dislodge the insect. Rummel and Arnold (1989), for example, found that thrips counts were five to six times higher when sampled by washing vegetation. A sample unit of 10 blossoms from each of five areas in a field is considered optimal (Cho et  al., 1995). Thrips densities can also be estimated with the use of sticky traps. Yellow, white, and blue traps are generally most attractive to western flower thrips (Yudin et al., 1987; Vernon and Gillespie, 1990; Roditakis et al., 2001). Trap efficiency is increased by highly contrasting background color; yellow in front of a violet background, for example, is highly attractive (Vernon and Gillespie, 1995). The position of the trap, especially the height above the soil, also affects thrips capture (Roditakis et al., 2001). Thrips can also be captured in water traps, and higher captures are made when certain volatile chemicals are added to the trap (Teulon et al., 1993). Sampling has been reviewed by Shipp (1995). In tomato, western flower thrips are most abundant in blossoms on the upper half of plants, and at field margins. Nymphs are more abundant in blossoms in the lower regions of plants (Salguero Navas et al., 1991a). Thrips populations, especially nymphal populations, are aggregated. A binomial, or the presence-absence, sampling program has been developed for tomato when thrips densities average less than 1.4 per blossom; 16–18 blossom samples are used to estimate abundance and suppression is initiated only when greater than 50% of the blossoms are infested (Salguero Navas et al., 1994). Insecticides. Insecticides are commonly applied to the foliage and blossoms of vegetables to minimize feeding and oviposition damage and to limit disease transmission. However, insecticide resistance is a widespread

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­phenomenon (Zhao et  al., 1995). The severity of the resistance problem in the field is exacerbated by the ability of thrips to infest and escape from greenhouses, where insecticide use is frequent. Rotation of insecticide classes is frequently recommended to forestall development of resistance. If insecticides are used, a common practice is to apply two treatments about 5 days apart because the eggs are within the plant tissue and the propupal and pupal stages are beneath the soil, and thus relatively immune. Cultural Practices. Plant culture affects thrips abundance, and thereby the incidence of plant disease. For example, high rates of nitrogen fertilization increase the abundance of thrips on tomato (Funderburk et  al., 2002). Also, sanitation is an important element in thrips management. Weeds can serve as important alternate hosts of both thrips and virus diseases, and their presence should be minimized. Also, if seedlings are used to initiate a crop, care should be taken to assure that they are free from thrips. The proximity of greenhouses is another consideration, as this may be a principal source of crop infestation by thrips, especially in cold climates where overwintering success by thrips is limited. Barriers are sometimes recommended for insect exclusion, including such small species as thrips. However, the small size of western flower thrips (width of males is about 184 μm and width of females is about 245 μm) requires extremely fine screen if thrips are to be denied access to plants. This fact excludes most standard materials from consideration as screens (Bethke and Paine, 1991). However, low barriers can be used under field conditions to limit thrips dispersal and invasion of crops (Yudin et al., 1991). Also, walk-in tunnels or greenhouses covered with ultraviolet light-absorbing plastic are less infested by thrips than ultraviolet light-reflecting coverings, apparently due to reduced attraction of plants grown in filtered light or to modified thrips feeding behavior (Antignus et  al., 1996). High levels of atmospheric carbon dioxide can be used along with elevated temperatures to disinfest produce and plants (Seki and Murai, 2012).

Reflective mulches help to deter invasion of susceptible crops by dispersing thrips (Stavisky et al., 2002). Some of the benefit can be lost as the canopy of the plants expands, but there is considerable benefit to be gained by delaying the inoculation of plant pathogens by thrips. For example, Riley and Pappu (2004) documented yield benefit associated with use of reflective mulch, and the additive effects of using reflective mulch plus resistant plants or thrips-­ effective insecticide treatments. Biological Control. Considerable emphasis has been placed on the development of biological control agents for thrips-infesting greenhouses, including western flower thrips. Such beneficial organisms as the parasitic wasp Ceranisus menes (Hymenoptera: Eulophidae) (Loomans et  al., 1995); the minute pirate bug, Orius laevigatus (Hemiptera: Anthocoridae) (Chambers et  al., 1993); the foliage-dwelling predatory mites Neoseiulus (Amblyseius) cucumeris (Oudemans) Amblyseius degenerans Berlese, A. swirskii Athias-Henriot, and others in this group (Acari: Phytoseiidae) (van Houten and van Stratum, 1995; Mouden et al., 2017); the soil-dwelling predatory mites Geolaelaps sp. (Acari: Laelapidae) (Gillespie and Quiring, 1990); and entomopathogenic nematodes (Nematoda: Heterorhabditidae and Steinernematidae) (Chyzik et  al., 1996) have been studied. Some beneficial organisms, particularly the mites N. ­cucumeris and A. swirskii, are used in commercial greenhouse vegetable production. Factors such as temperature, photoperiod, and crop type sometimes limit success, even under greenhouse conditions. For example, mite predators are much more effective in suppressing western flower thrips on pepper than on cucumber, presumably due to interference with mite searching behavior by the numerous trichomes found on cucumber leaves (Shipp and Whitfield, 1991). Several common entomopathogenic fungi such as Beauveria bassiana, Metarhizium anisopliae, Paecilomyces fumosroseus, and Lecanicillium lecanii (Verticillium lecanii) also have been used, but the level of suppression is only moderate. The expense and dispersal tendencies of most beneficial organisms thus far have limited their use to greenhouses.

Chapter 13

Other Invertebrate Pests

CLASS ACARI—MITES Banks Grass Mite

Oligonychus pratensis (Banks) (Acari: Tetranychidae)

Natural History Distribution. This species occurs principally in the southern United States, particularly in the southern Great Plains. However, it also can occur in northern areas, and in the Great Plains region it commonly causes injury as far north as Nebraska. In the northwestern states, it is sometimes called the timothy mite. Banks grass mite is also known from Hawaii, Puerto Rico, Central America, and Africa. Apparently, it is native to North America. Host Plants. This mite feeds predominately on wild grasses and on grasses grown as field crops. The crops injured include Bermudagrass, bluegrass, corn, sorghum, and wheat. However, dates are reported to be injured in southern California. Among vegetables, only sweet corn is damaged, principally in the southern Great Plains and southern Rocky Mountain areas. Among the many wild grasses that serve as hosts for Banks grass mite are wheatgrass, Agropyron spp.; gramagrass, Bouteloua spp.; bromegrass, Bromus spp.; wildrye, Elymus spp.; panicum, Panicum spp.; and sorghum, Sorghum spp. Natural Enemies. Predators and disease agents both can be important in population biology, though predators are considered most important. The most important predator is the predatory mite Neoseiulus fallacis (Garman) (Acari: Phytoseiidae), which can consume an average of 15 mites per day. However, other phytoseiids such as Amblyseius scyphus Schuster and Pritchard may assume great importance at times, or in certain locations. Ganjisaffar and Perring (2015) have suggested that the phytoseid Galendromus flumenis (Chant) could be useful as an augmentative biological control agent. Pickett and Gilstrap (1984) have provided

Handbook of Vegetable Pests. https://doi.org/10.1016/B978-0-12-814488-6.00013-3 © 2020 Elsevier Inc. All rights reserved.

information on the phytoseiid complex attacking Banks grass mite in Texas, including a key to the important species. Also important as predators of Banks grass mite are the small black ladybird beetles Stethorus spp. (Coleoptera: Coccinellidae); the sixspotted thrips, Scolothrips sexmaculatus (Thysanoptera: Thripidae); the lacewings Chrysoperla spp. (Neuroptera: Chrysopidae); and minute pirate bugs Orius spp. (Hemiptera: Anthocoridae). The fungi Neozygites spp. are common pathogens at times, especially when conditions are moist and cool. Dick and Buschman (1995), working in Kansas, have reported that though Neozygites sp. appeared frequently, the fungus required an ambient humidity of at least 80% and often occurred too late in the season to prevent crop injury. Life Cycle and Description. The number of life cycles varies with location. In the northern states, six to seven generations are common, though more occur in the south. The life cycle requires about 8–25 days depending on temperature. Although adults may overwinter and become active during the winter in mild areas of the country, eggs are not produced until spring. Egg. The egg is pearly white when first deposited, but eventually turns yellowish-brown. They normally are laid singly on the underside of the leaf. They also may be deposited on the mite webbing that covers the leaf. Just before hatching, the eyespots of the developing larvae become evident. Duration of the egg stage is 2–5 days under summer conditions, but may require up to 36 days in Washington. Larva. The larva is whitish or pinkish-white initially. As it matures, it becomes green. The six-legged larval stage typically persists for 2–3 days under good conditions, but may require up to 17 days in northern locations. Nymph. The protonymph and deutonymph stages each may require only 1–2 days for development, but under cool

601

602  Handbook of Vegetable Pests

conditions development time for each instar may extend to 13–18  days. The eight-legged nymphal stages are pale green or bright green, depending on their food.

plants. With continued feeding, leaves turn yellow and then necrotic and brownish as damage increases. The mite infestation progresses upwards on the plant, with older leaves suffering premature death. Young leaves may be deformed, and leaf tissue can be contaminated with webbing. Serious infestation may occur relatively early in the season, once the plant is half grown, whereas twospotted spider mite tends to attain high densities on corn only after flowering has commenced. Damage is also found in dry areas, where wheat and range grasses serve as overwintering hosts. In contrast, twospotted spider mite infestations often occur in more moist locations such as along rivers, where broadleaf plants such as alfalfa may serve as overwintering hosts.

Management

FIG. 13.1  Banks grass mite. (Drawing by J. Capinera.)

Adult. The eight-legged adult stage is greenish, with darker pigmentation laterally. Banks grass mite is quite similar to twospotted spider mite, Tetranychus urticae Koch, in appearance, but T. urticae has dark pigmentation mostly in the anterior half of the body whereas in O. pratensis the pigmentation extends back to the posterior tip of the body. Banks grass mite is slightly smaller and its body is more flattened than twospotted spider mite. Banks grass mite measures about 0.4 mm in length, with the females averaging larger, the males smaller. The abdomen tapers to a point in males whereas in the female it is bluntly rounded. Overwintering forms are orangish or pinkish. Adults live for about 7–23 days under warm conditions, and up to 48 days under cool conditions. The preoviposition period is only 1–2  days, with peak oviposition at about sixth day and a steady decrease thereafter. Females produce 7–14 eggs per day, with a total fecundity of 75–150 eggs. Fecundity is higher when mites feed on mature corn or moisture-stressed corn (Feese and Wilde, 1977), and Banks grass mite thrives under hot, dry conditions. Optimum temperature is about 36–37°C. Light and low humidity favor passive aerial dispersal by adults, which are aided in their dispersal by production of silk threads (Margolies, 1987). The biology of Banks grass mite was given by Malcom (1955) and Tan and Ward (1977). Temperature relations were described by Perring et al. (1984). A useful synopsis was provided by Jeppson et al. (1975). Banks grass mite is included in a key to vegetable-feeding mites in Appendix A.

Damage Damage is typical of mite infestations. Feeding initially causes the appearance of minute yellow spots on the ­undersides of leaves, particularly the lower leaves on

Sampling. Mite infestations should be checked regularly, especially if mites are detected early in the season. Populations usually are monitored visually, though a hand lens with 10 × magnification is generally needed for accurate assessment, and even greater magnification is helpful for species confirmation. Populations of natural enemies should also be considered in determining the need to suppress mite populations. Cultural Practices. Drought stress commonly favors mite population increase, so proper irrigation is an important preventative treatment. Late planting of corn alleviates infestation in some areas, and excessive fertilization with nitrogen should be avoided. The presence of mites in winter-grown wheat, an important overwintering host, is positively related to subsequent infestation levels in corn (Holtzer et al., 1984). Insecticides. The application of insecticides to foliage is sometimes necessary to prevent damage. In areas where both Banks grass mite and twospotted mite occur, it is important to determine the pest species present before the application of insecticides because they differ in their susceptibility to various products. Banks grass mite is resistant to insecticides in some areas, but is generally considered easier to suppress than twospotted spider mite. Sometimes Banks grass mite population increases following the application of insecticides for other insect pests. If possible, it is desirable to suppress caterpillar pests with microbial insecticide Bacillus thuringiensis, either as a liquid application or by genetically altered corn, because Bacillus thuringiensis does not cause destruction of mite predator populations and increases in mite abundance.

Broad Mite

Polyphagotarsonemus latus (Banks) (Acari: Tarsonemidae)

Natural History Distribution. Broad mite is found throughout the tropical regions of the world, and is a greenhouse pest in many

Other Invertebrate Pests Chapter | 13  603

temperate areas. In the United States, it is a minor citrus pest in Florida, California, and Hawaii, and occasionally injures greenhouse and field-grown vegetables in those states and in Georgia and Texas. Its origin is likely southern Asia. Host Plants. The host range of broad mite is extensive with plants from over 60 families known to be injured (Gerson, 1992). The principal exception is the grasses, which are not fed upon. Among the important crop plants injured are cotton, citrus, mango, papaya, passion fruit, rubber, tea, and tobacco. Among the vegetables reported to be injured are beet, bean, chard, cucumber, eggplant, pepper, potato, and tomato. Damage is especially severe in bell pepper. Ornamental plants also are affected, including azalea, balsam, begonia, chrysanthemum, gerbera, dahlia, fuchsia, zinnia, and likely many others. Weeds such as spiny pigweed, Amaranthus spinosus; beggartick, Bidens spp.; jimsonweed, Datura sp.; and galinsoga, Galinsoga sp. serve as important reservoirs for mites during crop-free periods. Broad mites also can develop on plant pollen (Pena, 1992). Natural Enemies. Predators have been observed to be an important elements of field ecology. The natural enemies are listed by Gerson (1992), but Amblyseius spp. and Typhlodromus spp. (both Acari: Phytoseiidae) are most important (Badii and McMurtry, 1984; Pena et al., 1989). Sometimes these predators are able to maintain suppression of mite populations on crops, including pepper and citrus. Among diseases, only the fungus Hirsutella nodulosa has been noted as a naturally occurring pathogen. Life Cycle and Description. The complete life cycle requires only about 7  days under favorable conditions. Optimal conditions for growth and reproduction are about 24°C and high humidity. Reproduction does not occur below about 13°C and above about 34°C. Under such favorable conditions 20–30 generations may occur annually. They are found reproducing throughout the year in southern Florida, but are most abundant during the summer months. Temperature of about 25°C and humid conditions are optimal. Hot, dry weather in the autumn and early winter are disruptive to broad mite populations in California. Diapause is unknown in this species. Egg. The eggs are laid singly and are found on the lower surface of young apical leaves and in flowers. The eggs are elongate-oval and translucent. The upper surface is covered with rows of hemispherical projections, whereas the ventral, flat surface is attached to the substrate. The projections are whitish, though the egg itself is nearly transparent. Hatching occurs after 2 days when maintained at 25°C and high humidity. Female fecundity averages 40–50 eggs (range 30–75 eggs). Larva. Larvae are small, flattened, and oval. They are whitish and bear six legs. Larvae measure about 0.12– 0.18 mm in length. The general appearance is similar to

that in the adult stage. The larval stage feeds for 1 day, then molts to a quiescent stage usually called the pupa. Pupa. The pupal (also called nymphal) stage is similarly brief, lasting only about 1–2 days, and occurs within the old larval cuticle. The body form is similar to that of the larva and adult, though the legs are eight in number and are larger than in the larval stage.

FIG. 13.2  Broad mite. (Drawing by J. Capinera.)

Adult. Adults also are flattened and broadly oval, particularly the females. They possess four pairs of legs, the anterior two pairs being widely separated from the posterior pairs. The legs are well equipped with spines. The females bear relatively short legs, usually measuring less than the width of the body, but males bear longer legs, with some about as long as the length of the body. The posterior end of the body bears a few hairs, but lacks long or stout spines. The color of the mites varies, but often is whitish, yellow or yellowish-green. These organisms are quite small; body length is 0.2–0.3 mm in females, and males only half as large. Females usually have an indistinct white stripe along the center of the back, though males lack this character. Adult males emerge before females. Males grasp female pupae with their genital organs and carry them upwards in the plant to younger tissue. Therefore, males effectively select the oviposition sites of females. Females normally copulate immediately upon emerging from the pupal stage, though they may mate later. Adult males persist for about 7 days, whereas females survive for about 10  days. The ratio of males to females varies among populations, but usually is weighted toward females, often at a ratio of 4:1. Unmated females produce only males, which may then mate with their mother, assuring production of females. Both sexes are very active, though they do not use their hind legs for walking. Adults disperse short distances within and among plants by walking, but disperse long distances by being blown by wind, and by attaching to, and “hitch-hiking” on winged insects, especially whiteflies. Descriptions of broad mite were given by Lavoipierre (1940) and Lindquist (1986). Gerson (1992) provided an excellent review of broad mite biology and management.

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Temperature and humidity responses were studied by Jones and Brown (1983). A simple key to common vegetablefeeding mites is presented in Appendix A.

Damage The damage caused by broad mite varies among host plants. In general, the mite seems to secrete a plant growth regulator or toxin, and it induces plant deformities as it feeds. Among the common symptoms are distortion of apical tissue, shortening of internodes, stem swelling, darkening, blistering, shriveling, and curling of leaves. Fruit and flower development may be inhibited, or fruit may be deformed, split, or russeted. Broad mite injury is commonly confused with herbicide toxicity, micronutrient deficiency, and virus diseases (Cross and Bassett, 1982; Pena and Bullock, 1994). Damage persists for some time even after the mites are controlled. The presence of only a few mites may induce injury. As few as 10 mites per plant may damage pepper, potato, and bean. Young pepper plants support higher rates of mite population increase, and are more easily damaged (de CossRomero and Pena, 1998). Damage may result at some distance from the site of feeding, and may persist for weeks after the mites are removed by the application of insecticide. These characteristics suggest the presence of toxins secreted by the mites.

Management Sampling. Mites are found on the lower surface of young foliage and in flowers. Populations often are highly aggregated. Insecticides. Various insecticides and acaricides are effective against broad mite (Schoonhoven et al., 1978), but some classes of pesticides such as pyrethroids are not satisfactory and may cause population increases. Pesticides normally are applied to the foliage, and often at short intervals due to rapid population growth. Sulfur is commonly recommended for suppression of broad mite infestations (Pena, 1988). Cultural Practices. Broad mites cannot survive long in the absence of suitable plants. Thus, clean cultivation between crops decreases the incidence of infestation, as does the elimination of nearby weeds. The wide host range of broad mite, however, makes it difficult to eliminate mites through cultural practices. Biological Control. The fungi Beauveria bassiana, Hirsutella thompsonii, and Paecilomyces fumosoroseus have been shown to infect and kill broad mite (Pena et al., 1996). The predatory mites Neoseiulus californicus (McGregor), and N. barkeri (Hughes) (both Acari: Phytoseiidae) were studied as inoculative agents by Fan and Petitt (1994) and Pena and Osborne (1996), and found to have potential for broad mite population regulation, particularly under

g­ reenhouse conditions. Neoseiulus cucumeris (Oudemans) was reported to suppress broad mite on greenhouse peppers in Israel (Weintraub et al., 2003). Repeated releases of predators often are required to maintain suppression of broad mite. Host Plant Resistance. Plant resistance has not been well studied for managing broad mite. Nevertheless, several chilli cultivars in India are resistant, and partial resistance was reported from sweet pepper in Cuba (Depestre and Gomez, 1995; Rao and Ahmed, 2001). Echer et al. (2002) have also reported the occurrence of broad mite resistance in two accessions in Brazil.

Bulb Mites

Rhizoglyphus echinopus (Fumouze and Robin) Rhizoglyphus robini Claparede (Acari: Acaridae)

Natural History Distribution. Several species of mites are associated with the bulb mite complex. Misidentification of mites is common, so the validity of many reports is uncertain. However, Rhizoglyphus echinopus (Fumouze and Robin) and R. robini Claparede are the species most commonly reported to injure vegetables (Manson, 1972; Díaz et  al., 2000). These species are found throughout the world including North America, where it widespread, but do not appear to be native to North America. Host Plants. Bulb mites are associated with bulbs and roots of numerous plants. They are most commonly reported to be associated with bulbs of flowering crops such as dahlia, hyacinth, iris, lily, narcissus, and tulip. Among vegetable crops, onion, garlic, and leek are occasionally damaged. Bulb mites also have been known to feed on beet, celery, and potato, and perhaps other vegetables with large roots or tubers, but they are not considered a pest of these crops except in storage. Life History and Description. Mites of the genus Rhizoglyphus are relatively large, usually measuring 0.5– 0.8 mm and sometimes up to 1.0 mm in length, and about 0.3 mm in width. They are oval, smooth in general appearance, and with a whitish or colorless body. Long hairs are sometimes found on the body, especially at the posterior end of the mite. Normally there are four pairs of thickened, short, light-brown legs. The legs bear hairs or spines, though they are less dense than in the spider mites. The first larval stage has only three pairs of legs. The eggs are oval, translucent and whitish, and are about one-half the size of the adult female. There are four immature stages. The life cycle is completed in 17–27 days at 18–24°C, and 9–13 days at 20–27°C. Under cooler conditions, however, the life cycle may require over 100  days. Optimal growth and development occurs at 22–26°C. The developmental threshold is about 9.7°C. Longevity is about 40–70  days. The female

Other Invertebrate Pests Chapter | 13  605

deposits eggs singly, producing up to 700. They hatch in 8–15 days. The mites are abundant in soil, and often associated with decaying plant material. The hypopal stage sometimes appears in mite colonies; this small (0.3 mm long), dark-colored, and sluggish form will attach to insects and thereby be transported to new locations. The ecology of R. robini was studied by Gerson et al. (1985) in Israel. They have reported that both clay and loam soils were suitable for the mites, but sandy soils were not. Summer soil temperatures were too high for the mites unless the soil was irrigated. Temperatures of about 35°C caused sterility. Thus, mite populations tended to increase during the autumn and winter, declining in the spring. Virtually any organic material was suitable for mite maintenance, but manure was stimulatory.

FIG. 13.3  Bulb mite. (Drawing by J. Capinera.)

Description and biology of bulb mites were provided by Hodson (1928), Hughes (1961), Gerson et  al. (1983) and Fan and Zhang (2003). Manson (1972) has provided keys to some species and clarified some synonomies. Bulb mites are also included in a key to vegetable-feeding mites in Appendix A. Mite culture techniques were given by Gerson et al. (1991). Díaz et al. (2000) have provided a review of the economically important Rhizoglyphus spp.

Damage Most astigmatid mites feed on fungi or decaying plant tissue, and are not considered to be primary plant pests. However, bulb mites may invade and enlarge bulb tissue previously invaded by fungi. Mechanical injury, which may result from tillage or injury by other pests, also may create an environment suitable for bulb mite population growth, but this form of damage is not as conducive for mite population growth as is plant disease (Okabe and Amano, 1991). Bulb mites apparently inhibit the recovery of bulbs from injury, and aid in the transport of fungi. Rawlins (1955), working in New York, has reported that Rhizoglyphus robini

(reported as R. solani) fed on roots, destroying field-grown onions at both the seedling and mature bulb stages. Ofek et al. (2014) similarly have documented the negative effects of R. robini on seedling growth, and the attractiveness and suitability of fungi to this mite species. Latta (1939) has also reported root damage, but by R. echinopus (reported as R. hyacinthi) on lily. R. echinopus usually is considered to be a pest of stored products, particularly those that have not been dried properly. They are often found in association with rot in stored potatoes. However, there are enough reports of plant damage associated with this species to be very concerned about its abundance.

Management Insecticides. Bulb mites are quite tolerant to pesticides (Knowles et  al., 1988). Soil fumigants may be applied to reduce the incidence of fungi and mites (Jefferson et  al., 1956). However, chemical options for management are increasingly limited (Díaz et al., 2000). In some studies, fungicide application reduces disease, but not mite abundance (Ascerno et al., 1981). Suppression of disease alone often increases crop yield, supporting the belief that bulb mites are not primary pests. However, Ascerno et al. (1983) have suggested that if mites attained a threshold of abundance, it was necessary to suppress them if fungicidal suppression of disease was to be effective. Cultural Practices. Bulbs that are lightly infested can be purged of mites before planting; a hot water bath at 43°C for 1 h is adequate (Weigel and Nelson, 1936). Mulching with transparent polyethylene raises the temperature of soil, and can be used to reduce the abundance of both fungi and mites (Gerson et al., 1981). In fields where bulb mites have been observed, prolonged fallow periods are recommended to eliminate the food supply of the mites. It is important to eliminate organic residues under such circumstances, both crop remains and manure, and to avoid following with a susceptible crop.

Twospotted Spider Mite

Tetranychus urticae Koch

Strawberry Spider Mite

Tetranychus turkestani Ugarov and Nikolski

Tumid Spider Mite

Tetranychus tumidus Banks (Acari: Tetranychidae)

Natural History Distribution. The wide distribution of spider mite species is largely due to their small size and the ease with which they are inadvertently transported in commerce. Their wide host range also allows them to establish ­readily

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in new locations. Recently, their growing tolerance to many insecticides (acaricides) has exacerbated their tendency to survive quarantine treatments, further enhancing spread. The most widespread is twospotted spider mite, Tetranychus urticae Koch, which is found throughout the world in temperate and subtropical locations. Strawberry spider mite, Tetranychus turkestani Ugarov and Nikolski, is found in Europe and Asia in addition to North America. Tumid spider mite, Tetranychus tumidus Banks, is commonly found from the southern United States south through Central America and the Caribbean to Brazil. These mites are most damaging in the warmer, southern areas of North America, though twospotted spider mite survives temperate climates better than the other species. Within greenhouses, these species can be very abundant anywhere in North America, though twospotted mite is the most troublesome of the group. Other tetranychid mites which are known from vegetables, but are less common as pests, include desert spider mite, Tetranychus desertorum Banks in the southern United States, and Pacific spider mite, Tetranychus pacificus McGregor, in the western states. Carmine spider mite, Tetranychus cinnabarinus (Boisduval) is reported to be troublesome on occasion, especially in Hawaii, but it is uncertain whether this is a separate species or a form of T. urticae. The taxonomy of spider mites is uncertain, and name changes are frequent Host Plants. Twospotted spider mite is said to have a host range of over 1000 species (Dermauw et al., 2013), but not all of these “hosts” are accepted equally. For example, although plants in the family Fabaceae are readily, accepted, plants in the family Solanacae are quite variable is their suitability (van den Boom et  al., 2003). These mites affect numerous vegetable crops, including bean, beet, cantaloupe and other melons, carrot, celery, corn, cucumber, eggplant, parsley, pea, pepper, squash, sweet potato, tomato, and likely others. Tomato, bean, and cucurbit crops are affected most often, and twospotted spider mite is the most likely species to cause injury. Most of these mites are also known from cotton, soybean, strawberry, tree fruit crops, and ornamental plants. Of these three species, however, twospotted spider mite is likely to affect tree fruits and ornamentals the most, with the other species normally associated with low-growing crops. Natural Enemies. The natural enemies of spider mites are numerous and diverse. Among the most important are predatory mites, particularly Phytoseiulus persimilis AthiasHenriot (Acari: Phytoseiidae); lady beetles, particularly Stethorus spp. (Coleoptera: Coccinellidae); dusty-wings (Neuroptera: Coniopterygidae); pirate bugs, particularly Orius spp. (Hemiptera: Anthocoridae); some plant bugs (Hemiptera: Miridae); and thrips, particularly Scolothrips sexmaculatus (Pergande) (Thysanoptera: Thripidae) (Huffaker et  al., 1969), and some tomato bugs (Hemitera: Miridae). Also, fungi sometimes cause ­epizootics (Dick and Buschman, 1995). The natural enemies of tetranychids were reviewed by McMurtry et al. (1970). van de Vrie et al. (1972)

have discussed population ecology, including population regulation. Life Cycle and Description. With a life cycle of only 8–12 days at 30°C and about 17 days at 20°C, over 20 generations may develop annually, though conditions rarely allow this rate of population cycling. Overwintering may occur on many hosts in warm-winter climates, and in cold winter areas forage legumes and greenhouses often shelter these pests, but adult twospotted spider mite females also pass the winter under leaves or other organic debris in a state of diapause. The development time of the immature stages is 4–5 days at 30–32°C, but is extended to about 16–17 days when the temperature is 15°C at night and 28°C during the day. Egg. The eggs are whitish and spherical in form. They measure about 0.10–0.15 mm in diameter. They are often deposited singly on the lower surface of foliage, but sometimes the upper, and the leaf surfaces are covered with strands of silk. Females oviposit at 5–6 eggs per day, for a total of 60–120 eggs. Duration of the egg stage is about 3 days at 30°C and 6–7 days at 20°C. Larva. The first instar is called the larva, and is colorless initially but yellowish or pinkish after feeding. The body is nearly spherical in shape, and bears three pairs of legs. The terminal portion of the larval stage is a nonfeeding period called the nymphochrysalis or protochrysalis. Duration of the first instar is 1–2 days at 30°C and 2–3 days at 20°C. Nymph. There are two nymphal instars—the protonymph and the deutonymph. These stages are easily separated from the larva because they bear four pair of legs. They tend to be green or red. As in the larval stage, the terminal portion of each nymphal period is a nonfeeding period called the deutochrysalis and teliochrysalis, respectively. The duration of each instar is 1–2 days at 30°C and about 3 days at 20°C.

FIG. 13.4  Spider mite. (Drawing by J. Capinera.)

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Adult. Adults are 0.4–0.5 mm long, males averaging slightly less than females and are usually less abundant than females. Like the nymphs, adults bear four pairs of relatively long legs. They also have numerous long hairs on their legs, and long but sparse hairs on their body. Females tend to be oval in body shape, males have elongate-oval or diamondshaped body. In twospotted spider mite and strawberry spider mite, the actively feeding female is usually greenish, with dorsolateral dark spots. The two species sometimes may be distinguished by the number of dark spots, which is two in twospotted but four in strawberry spider mite. In tumid spider mite, the female body color is reddish with dark lateral markings. Overwintering females of twospotted spider mite and strawberry spider mite become orangishred. Color is not a very reliable character with tetranychid mites; accurate determination depends on the examination of tarsal characteristics and genitalia of males. Males are attracted to immature females by a sex pheromone, perform extensive mating rituals (Cone et al., 1971a,b) and may mate repeatedly. The preoviposition period of females is 1–2 days. Fertilized females produce both male and female offspring; unfertilized females produce only males. Duration of the adult stage is normally about 30 days except when overwintering. The preoviposition period of adults is less than a day at 30°C, and 1–2 days at 20°C. Adults disperse by crawling and are dispersed by wind. The biology of tumid spider mite was described by Saba (1974). Laing (1969) and Carey and Bradley (1982) gave development and temperature relations for T. urticae, T. turkestani, and T. pacificus. Behavior and ecology of Tetranychus spp. were also described by Huffaker et al. (1969) and van de Vrie et  al. (1972). Comprehensive accounts of twospotted spider mite were given by Ewing (1914) and of strawberry spider mite by Cagle (1956); biology of carmine spider mite was described by Hazan et al. (1974). Descriptions and keys are provided by Jeppson et al. (1975) and Kono and Papp (1977), Baker and Tuttle (1994), and Bolland et  al. (1998). A simple key to mites found on vegetables is included in Appendix A.

Damage Spider mites generally feed on the lower leaf surface, though twospotted spider mite affects the upper surface of some host plants. They pierce individual cells with their stylets, withdrawing the cell contents. Twospotted spider mite can feed on 18–22 cells per minute, resulting in many dead cells, and often a speckled appearance. Leaf transpiration is accelerated, and affected leaves may dry and drop from the plant. Yellowing and speckling are the most common early plant responses to feeding, though reddening may also occur. Injection of plant growth regulators or interference with growth regulators during feeding is also reported. Wilting, tissue death, leaf deformity, and abcission are characteristics of prolonged and high-density infestations. Disruption of photosynthesis results in stunting of plant growth and reduced-fruit yields.

Mite products such as webbing, eggs, cast skins, and fecal material also detract the cosmetic quality of plants. Before the 1940s, spider mites were infrequently considered to be serious pests, but since then they have assumed major pest status in some crops. Apparently, mite problems are induced by crop management practices, particularly the use of broad-spectrum insecticides (see section on “insecticides”). Also, the suitability of crops for mites is greatly enhanced when mites develop on plants which receive excessive nitrogen fertilization, grow in a dusty environment, or are stressed by inadequate moisture and high temperature. These environmental factors can convert plants which might be only poor hosts into very good hosts, resulting in mite population increase and crop damage.

Management Sampling. Visual examination of foliage for leaf stippling on the upper surface, and mites and webbing on the lower surface, is the usual method of sampling. Older, or lower, leaves are usually examined. Infestations tend to be clumped initially, with clumping decreasing as the crop matures and females disperse to younger foliage (Perring et al., 1987). Insecticides. Chemical insecticides (acaricides) are commonly used in greenhouses to prevent injury by mites, though natural enemies usually are capable of maintaining spider mite densities at low levels on crops grown under field conditions. In the field, insecticides directed at other pests can induce mortality among natural enemies of mites, causing increase in spider mite populations. Therefore, considerable effort is now directed at managing pests without disrupting natural control of mites, often by the use of selective insecticides. Avoidance of early season applications of insecticides is also encouraged. Certain classes of insecticides, particularly pyrethroids, are especially disruptive (e.g., Rock, 1979). If nonsystemic chemicals are applied for mite suppression, thorough coverage of the plants is essential (Hagel and Landis, 1972). Frequent application of insecticides has led to many cases of resistance among spider mites. Resistance has not developed to oils, but oils are effective mostly against eggs, and frequent application leads to phytotoxicity in some vegetable crops. Insecticidal soaps are similarly useful, though eggs are not entirely susceptible (Osborne and Petitt, 1985). Neither oil nor soap should be used at temperatures above 32°C. Sulfur is applied to some crops, but not to cucurbits, and is not used if temperatures exceed 32°C. Polyphagous plant feeders such as spider mites must contend with a large number of chemical defenses that evolved in plants to defend against herbivory. They may avoid, sequester, or detoxify natural defense compounds. Thus, they are predisposed to overcome plant protection pesticides as well. This can lead to the rapid evolution of pesticide resistance, and T. urticae is especially adept at developing insecticide resistance (Dermauw et al., 2013). Under field conditions, predators can often provide effective suppression of mites. However, the crops may

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r­equire chemical intervention due to weather that is especially favorable to the mites, or due to the presence of other insects that are not under biological suppression. Thus, insecticides (including microbial insecticides) may be a necessary part of the crop management program. Mites and their predators usually differ in their susceptibility to insecticides, with the predators often being more susceptible to the insecticides. However, predators differ in susceptibility, so research is needed to determine how insecticides can be integrated into the crop management program (Castagnoli et al., 2005; Poletti et al., 2007). Cultural Practices. Cultural practices have considerable influence on mite damage. Water stress disrupts the physiology of the host plant, making it more suitable for mite survival and population increase. Dry, dusty conditions also favor mite survival, because blowing dust interferes with the predators of mites more than it does with the mites; the latter are partially protected beneath their silk webbing. Thus, overhead sprinkler irrigations may alleviate mite problems. Excessive nitrogen fertilization of crops also favors mite population increase. Weeds and senescent crops can be important sources of mites, as can winter legume forage crops. Plant cultivars vary considerably in resistance to mites, and in some crops this offers excellent opportunity to manage mites, whereas in others there seems to be little inherent resistance (Childs et al., 1976). Biological Control. The biological suppression of mites has been well developed for greenhouse crops, but is infrequently practiced for annual crops grown outdoors. The predatory mite Phytoseilus persimilis Athia-Henriot (Acari: Phytoseiidae) is commonly used to suppress twospotted spider mite in greenhouses. Effective use of P. persimilis involves maintenance of a low level of pest mites so that the predators do not starve, and the distribution of pests (prey) uniformly in the greenhouse so that the predatory mites also become widely distributed. Supplemental release of predatory mites may be needed to maintain a favorable ratio of predators to prey, often between 1:6 and 1:25. The maintenance of stable predator-prey-host plant relations is not a simple task, and even seemingly benign environmental changes like variation in light intensity within the greenhouse can affect stability (Nihoul, 1993). The successful use of P. persimilis for twospotted spider mite suppression in strawberry fields (e.g., Decou, 1994) demonstrates the potential for use in field-grown vegetable crops.

Tomato Russet Mite

Aculops lycopersici (Massee) (Acari: Eriophyidae)

Natural History Distribution. Tomato russet mite was first observed in Florida in 1892, but was not viewed as a serious pest in North

America until about 1940 when it caused considerable damage in California. Following its appearance in California, it quickly spread eastward across the United States, attaining New York and Georgia in 1953. It is also known from Hawaii. The origin of this mite is uncertain, though not native to North America, and it is now found throughout the world in both temperate and tropical latitudes. It apparently overwinters outdoors successfully at southern latitudes, but in cold areas it survives only in greenhouses or is reintroduced with seedling plants shipped from the south. Host Plants. This species feeds principally on plants in the family Solanaceae. Among vegetables, it is not only a pest of tomato, but also affects eggplant, pepper, potato, and tomatillo. Other solanaceous plants that can serve as hosts include petunia; black nightshade, Solanum nigrum; cape gooseberry, Physalis peruviana; Chinese thornapple, D. ferox; hairy nightshade, S. villosum; jimson weed, Datura stramonium; popolo, S. nodifolium; small-flowered nightshade, S. nodiflorum; and silverleaf nightshade, S. elaeagnifolium. In general, the Lycopersicon spp. are very suitable, whereas the Nicotiana spp. are unsuitable. The only nonsolanaceous plants known to support this mite are Convolvulus species such as bindweed and morning glory species. Natural Enemies. The principal natural enemies of tomato russet mite are predatory mites, but predatory thrips and a cecidomyiid have been noted. Among specific examples of predators found in North America are Typhlodromus spp. (Acari: Phytoseiidae) (Anderson, 1954); Seiulus sp. (Acari: Phytoseiidae), black hunter thrips, Leptothrips mali (Fitch) (Thysanoptera: Phlaeothripidae), an unspecified predatory gall midge (Diptera: Cecidomyiidae) (Bailey and Keifer, 1943); Euseius concordis (Chant) (Acari: Phytoseiidae), Pronematus ubiquitus (McGregor), and Homeopronematus anconai (Baker) (both Tydeidae) (Royalty and Perring, 1987). Momen and Abdel-Khalek (2008) reported that Amblyseius (Typhlodromips) swirskii (Athias-Henriot) (Acari: Phytoseiidae) and Typhlodromus athiasae Porath and Swirski (Acari: Phytoseiidae) developed successfully while feeding on tomato russet mite, with the former developing more quickly and displaying higher fecundity. In the same study, Paraseiulus talbii (Athias-Henriot) (Acari: Phytoseiidae) did not survive to adulthood when fed only tomato rust mite. Park et  al. (2010) noted that A. swirski could consume in excess of 100 tomato russet mites per day. Life Cycle and Description. This mite reproduces whenever weather permits; in California this tends to be May–November. Their persistence is dependent on the availability of green plants, as there is no overwintering form. A generation may be completed in about 7 days (range 6–13 days). Egg. The eggs are deposited among the leaf hairs or crevices, and on the stem tissue. The eggs are spherical in shape and whitish or yellowish in color. They measure about

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0.45–0.60 mm in diameter. The incubation time of eggs can be as little as 2 days, but 3–4 days is more common. Nymph. The nymphs normally do not move far from the point of hatching, but tend to aggregate along the edges of leaves. Nymphs are elongate, broad anteriorly and tapering to a blunt point posteriorly. They are fairly featureless in shape, but bear two pairs of cephalothoracic legs. Long hairs are found at the tip of the abdomen. Careful examination reveals that the abdomen bears numerous minute rings. There are two instars which measure 0.09–0.10 and 0.14– 0.16 mm in length, respectively. Duration of the instars is 1–2 days for each.

the tomato stalk at the soil surface and spreads upwards on both stalk and leaf tissue. The surface feeding produces a bronzing or russeted appearance on both the stems and leaves. Infested leaves first curl at the leaf edges, then become dry and drop from the plant (Royalty and Perring, 1988). Unlike many mites, eriophyids feed readily on the upper surface of leaves and tolerate direct sunlight. Fruit and blossoms are rarely infested, but fruit may be scalded by the sun following loss of leaves. Injury is usually first observed in small patches of the field, but wind-borne dispersal of the mites soon results in widespread infestation. It is interesting to note that the glandular trichomes found on tomato, which normally protect the plants from feeding by many insects and mites, are ineffective against tomato russet mite because the mites are so small that they can move between the trichomes (van Houten et  al., 2012). Tomato russet mite feeding also triggers a localized degradation and loss of these trichomes, compromising the plant’s defenses.

Management

FIG. 13.5  Tomato russet mite. (Drawing by J. Capinera.)

Adult. The adult form is similarly tapered or wedge-shaped in general appearance. The adults measure 0.15–0.20 mm in length and 0.05 mm in width. They are yellow-orange. The abdomen is covered dorsally by plate-like ridges, and the anterior portion of the body with a cephalothoracic shield. The adults also bear two pairs of cephalothoracic legs and long hairs at the tip of the abdomen. Adult females have a prereproductive period of about 2 days. Reproductive capacity is about 40–50 eggs, and capacity to increase is great. Bailey and Keifer (1943) have estimated that with females producing about two eggs per day, one mated female could result in a population of 350 mites within 21 days and the population would double every 3–4 days. Rice and Strong (1962) have indicated that life processes were favored by temperature of about 27°C and relatively low humidity (about 30%). Adult longevity is about 16–22  days. Eggs from unfertilized females develop into males; those from fertilized females develop into females. The biology of tomato russet mite was described by Bailey and Keifer (1943), Anderson (1954), Rice and Strong (1962), Kim et  al. (2002), and Perring and Farrar (1986); the latter is a comprehensive review. An interesting general discussion of eriophyids was given by Keifer (1946).

Damage Mites injure the plant by puncturing the surface cells with their needle-like chelicerae. Injury usually appears first on

Sampling. These minute mites are barely visible without magnification, making field scouting difficult. Thus, most monitoring is done by watching for damage, with confirmation of mite presence accomplished microscopically. Mite infestations are usually detected when fruits are present but still small. Insecticides. Sulfur in dust or wettable form has long been used for mite suppression, but other insecticides and acaricides also are effective (Perring and Farrar, 1986; Royalty and Perring, 1987). Cultural Practices. Mites are readily dispersed by people and equipment, so care should be taken to minimize traffic within infested fields. Weeds and plants in greenhouses are important in overwintering. In California, early plantings are more likely to be infested than later plantings, but type of irrigation does not affect infestation levels (Zalom et  al., 1986b). There is some indication of differences in susceptibility among commercially available tomato cultivars (Kamau et al., 1992).

CLASS COLLEMBOLA—SPRINGTAILS Garden Springtail

Bourletiella hortensis Fitch (Collembola: Sminthuridae)

Natural History Distribution. The springtails of North America consist of both native and introduced species, though the species most commonly cited as causing injury also occur in Europe and likely are immigrant species. Probably less than 20 species are suspected of causing crop injury in North America. Among the most injurious is garden springtail,

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Bourletiella hortensis Fitch (Collembola: Sminthuridae), which accounts for most reports of injury in eastern North America. However, Onychiurus armatus Tullberg, Onychiurus pseudarmatus Folsom (both Collembola: Onychiuridae), and Ceratophysella armata Nicolet (Collembola: Hypogastruridae) are responsible for most of the injuries reported in western areas. The latter species are poorly studied. Host Plants. Springtails feed mostly on dead plant tissue and fungal hyphae. Some also feed on other animals such as nematodes and other springtails, and a few attack growing plants. Many vegetable crops are reported to be injured, including bean, beet, broccoli, cabbage, cantaloupe, carrot, cauliflower, celery, cucumber, lettuce, lima bean, onion, parsnip, pea, potato, pumpkin, radish, spinach, squash, tomato, turnip, and watermelon. Other crops such as sugarbeet, rye, tobacco, vetch, and wheat, as well as turf grass and ornamental annual plants are occasionally attacked. Natural Enemies. Many arthropods consume springtails, including spiders (Araneae), centipedes (Chilopoda), mites (Acari), and beetles (Coleoptera: Carabidae and Staphylinidae). Vertebrates such as toads, lizards, and birds also consume springtails. Parasitism by nematodes and fungi (and microsporidan fungi, formerly considered to be protozoa) is well documented, but seems to be of little importance as mortality factors. Life Cycle and Description. Springtails are primitive arthropods, considered by some to be insects and by others to be their own class of organisms. They are widespread and numerous. They may be the most abundant soil-dwelling animals, but can be outnumbered by soil-dwelling mites. Springtails bear a ventral tube (collophore) on the first abdominal segment, and many bear a jumping organ (furcula) on the third and fourth abdominal segment. It is the jumping organ and the leaping behavior that it imparts, of course, that is the basis of its common name “springtail.” Egg. The eggs may be laid singly or in clusters, and are often deposited in crevices or other sheltered locations. They are spherical, measuring about 0.2 mm in diameter. Duration of the egg stage is about 10 days. Juvenile. Juvenile springtails closely resemble the adults, differing principally in size and the lack of sexual characters. They normally pass through five to eight instars before attaining adulthood, with instar durations of about 4 days for each. Adult. There are two basic body types present among springtails. One body type is represented by Bourletiella hortensis Fitch and other members of the family Sminthuridae. In sminthurid springtails, the first four abdominal segments are fused to form a globular mass, and overall the body is short and plump. The other basic body

form is represented by the Onychiurus spp. and members of many common families. In these latter springtails, the abdominal segments are clearly differentiated, and the body is relatively long and thin. The presence of antennae, six abdominal segments, a collophore, a furcula, and three pairs of legs serve to distinguish springtails from the numerous other small animals, including immature insects, inhabiting soil and leaf litter. They differ greatly in color, with some species brightly colored or ornately decorated. B. hortensis, for example, is mottled blue to purple brown in body color, with purplish antennae, and sometimes with yellow areas. In contrast, O. armatus is entirely whitish. Springtails range in length from 0.5 to 8.0 mm, though generally 1–3 mm, and are wingless. The period of activity is variable among springtails, with some active during the day but most active principally at night. The adult stage continues to molt, sometimes up to 40 times, but they do not change in appearance. The duration of each instar is often about 6 days for young adults, and lengthen to about 10 days after instar 15. Many springtails live for more than a year, and mean fecundity is often about 400 eggs per female. Sexual dimorphism is rare, and though parthenogenesis occurs in some species, many springtails engage actively in mating. Males normally deposit their sperm externally, in small packets (spermatophores) suspended on stalks. Females either take these spermatophores up willingly, or are guided to them by males. In some species, the males have grasping organs on the antennae, which are used to grasp the female’s antennae and to back her into a spermatophore. Several pheromones are documented in springtails, including sex, alarm, and aggregation pheromones, though they are poorly known. The aggregation pheromone is probably quite important in springtail biology, as aggregation is a regular and widespread feature of their behavior. Interestingly, springtails can respond to the pheromone produced by other springtail species, but respond most strongly to pheromone produced by their own species. Reviews of springtail biology were published by Christiansen (1964) and Hopkin (1997). A comprehensive key to American springtails was provided by Christiansen and Bellinger (1998). Other useful keys included a key to genera (Christiansen, 1990), a pictoral key to genera (Scott, 1961), and regional keys for Iowa (Mills, 1934) and New York (Maynard, 1951).

Damage Damage by springtails is usually observed above-ground, often in the form of small holes in the young leaves, and injury that greatly resembles feeding by flea beetles (Coleoptera: Chrysomelidae). However, they also feed on stems and roots. Garden springtail tends to feed readily above-ground, whereas Onychiurus spp. feed mostly belowground. Although injury is often limited or localized within

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regions of fields, sometimes entire crops are destroyed by these small but numerous arthropods (Folsom, 1933; Scott, 1964; Getzin, 1985; Ireson, 1993). Some of the damage caused by springtails is offset by their beneficial fungus-feeding activities. They feed on plant pathogenic fungi and some have suggested that they could be used to suppress fungi affecting plant roots of container-grown crops (Hopkin, 1997). In natural environments, they are also quite important in the decomposition of organic matter, increasing the rates at which nutrients are released from decomposing vegetation.

Management Sampling. Springtail populations can be assessed by visual observation, pitfall traps, or by soil sampling. In the latter case, springtails are usually separated from the soil by driving them from the soil with heat. Insecticides. Insecticide treatments normally are recommended only in fields that have a history of problems or where cultural practices increase the possibility of injury. Soil treatments, usually applied as granular or liquid formulation in the furrow or in a band over the row, are common practices. Seed treatments also can be effective, but only with certain insecticides (Getzin, 1985). Some systemic materials can protect both the seed and seedling from damage. Foliar applications can be effective for some species, but are not often used unless prolonged cool weather is hampering seedling growth. Cultural Practices. Soil high in organic matter is favorable to springtail population growth, and such fields are more likely to experience injury. Injury also follows application of organic matter, minimum tillage practices favoring retention or crop residue, and conversion of uncultivated fields to crop fields. Also favoring injury are furrow irrigation and soils that tend to produce deep soil cracks in response to drying, because cracks afford good shelter during periods of drought (Scott, 1964).

CLASS DIPLOPODA—MILLIPEDES Garden Millipede

Oxidus gracilis Koch (Diplopoda: Paradoxosomatidae)

Natural History Distribution. Several species of millipedes are reported to be injurious, or present in high enough numbers to be considered a nuisance. Probably, the most important is garden millipede, Oxidus gracilis Koch (Diplopoda: Paradoxosomatidae), which is abundant in southern and western states, and in greenhouses elsewhere. It apparently was accidentally introduced from Asia, though it may have arrived circuitously via Europe, where it has also become a greenhouse pest.

Among other common millipedes that sometimes may be pests are Cylindroiulus caeruleocinctus (Wood), and Diploiulus latistriatus (Curtis) (both Diplopoda: Julidae), which occur in the eastern and Midwestern United States and adjacent areas of Canada, though the distribution of D. latistriatus also extends west to Washington and British Columbia. Also, Blaniulus guttulatus (Fabricius) (Julidae), known as the spotted snake millipede, is reportedly widespread in North America. In California, it has become troublesome in the San Luis Obispo area where it damages vegetables such as squash, bean, potato, and turnip by feeding on the roots, especially during dry soil conditions. All three of these juliids are immigrants from Europe. Another European immigrant species is Polydesmus inconstans Latzel (Diplopoda: Polydesmidae), which is similarly found across the North American continent along the United States-Canada border. North America also has numerous native species that may become abundant locally, but they rarely are damaging. Millipedes are poorly known animals, with almost all of our information based on studies in northern Europe. Host Plants. Millipedes normally eat dead plant material, usually in the form of leaf litter. However, they occasionally graze on roots and shoots of seedlings, algae, and dead arthropods and molluscs. They are selective in their consumption of leaf litter, preferring some leaves over others. They also tend to wait until leaves have aged, and are partially degraded by bacteria and fungi. Thus, they function principally as decomposers, hastening the breakup of leaf material into smaller pieces, and incorporating the organic matter into the soil. Whether they derive most of their nutritional requirements from the organic substrate or the microorganisms developing on the substrate is uncertain. Millipedes also tend to consume their own feces, and many species fare poorly if deprived of this food source. Many vegetable crops have been reported to be injured by millipedes, including bean, cabbage, carrot, cauliflower, corn, cucumber, lettuce, parsnip, pea, potato, radish, tomato, and turnip. In addition, annual flower crops such as coleus, geranium, sweet pea, as well as other greenhousegrown plants may be damaged. Natural Enemies. Despite the formidable chemical defenses of millipedes, several natural enemies are known. Small vertebrate predators such as shrews, frogs, and lizards eat millipedes. Invertebrate predators such as scorpions (Arachnida), ground beetles (Coleoptera: Carabidae), and rove beetles (Coleoptera: Staphylinidae) also consume millipedes, though ants are usually deterred. Various ­disease-causing agents such as fungi, iridoviruses, rickettsia, and protozoa are documented from millipedes, though they seem to be only sporadically effective. Although flies have been reared from millipedes in Europe, they seem not to be important parasitoids in North America.

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Life Cycle and Description. The name “millipede” suggests that these animals have 1000 legs, and though the presence of numerous legs is a characteristic of this group, none have more than 375 pairs and most have considerably fewer. In differentiating this group from the similar-­appearing centipedes and symphylans, the presence of two pairs of legs per body segment is the key character used to identify millipedes. Symphylans also can be pests (see section on garden symphylan, Scutigerella immaculata (Newport)), but centipedes are carnivorous and therefore beneficial. Millipedes are morphologically quite diverse, though they all consist of a long chain of rather uniform body segments, and lack wings. Some are rather short, and may be covered with feather or scale-like adornment. Others look greatly like woodlice, and even roll into a ball in the manner of pillbugs. Most, however, are elongate and thin in general body form. There are three basic body regions: the head, which bears a pair of moderately long antennae; the body, consisting of numerous leg-bearing segments, and which normally are rather cylindrical but sometimes bears prominent lateral projections; and the telson, or posterior body segments bearing the anus. The integument is very hard. The life cycle of millipedes is often long. Many live for a year, but some persist for 2–4 years before attaining maturity. In a Julus sp. studied in a temperate environment, oviposition took place in April, with instar 6–7 attained by winter and instar 8–9 by the second winter. They overwintered as instar 9–11, then mated and oviposited the following spring, their third year of life, before they perished. Following is a description that is based, as noted, mostly on garden millipede, a species with a 1-year development time. Egg. The creamy white, yellow or brownish eggs are deposited in the soil, usually in clusters. A glutinous material causes them to adhere to one another. They are nearly spherical in shape, measuring 0.35–0.4 mm in width and about 0.40 mm in length. The female may create a chamber or cell for her eggs. Garden millipede deposits about 50–300 eggs in a cluster, and they can be found throughout the summer months. Duration of the egg stage is 9–10 days. Juvenile. The first instar juvenile millipedes bear only a few segments and three pairs of legs, but body segments, legs, and ocelli are added with each molt. By counting the number of rows of ocelli and adding one, the instar can be estimated for many millipedes. Thus, fourth instar millipedes have three rows of ocelli, fifth instars have four rows, etc. (Hopkin and Read, 1992). Maturity is often attained after about 10 instars, but some species continue to molt as adults. In garden millipede, most individuals develop through eight instars before they attain the adult stage. The number of pairs of legs present in the juveniles is about 3, 6, 11, 16, 22, 26, 28, and 30 for instars 1–8, respectively. Body length

is about 0.5, 1.5, 3.6, 4.1, 4.8, 7.4, 12.4, and 20 mm during the corresponding instars. Development time is 1, 11–18, 13–18, 16–30, 20–38, 28–46, 42–60  days during instars 1–7, with the final (eighth) instar generally overwintering, though in some cases instar 7 overwinters.

FIG. 13.6  Millipede. (Drawing by J. Capinera.)

Adult. The adult millipedes vary considerably in size, often measuring from 10 to 30 mm in length, but in some species exceeding 100 mm. Their color ranges from whitish to brown and black. Their sexes are separate. The external genitalia of adult millipedes are located between the second and third pairs of legs. Some adult millipedes have the ability to molt from a sexually active adult to an intermediate stage which is not functional sexually. Parthenogenesis occurs in some species and some populations, but this is not usual. Millipedes lack a waxy cuticle and are susceptible to desiccation. They have glands, with openings usually located laterally, which secrete chemicals that are toxic and may immobilize predatory arthropods like spiders and ants. The body segments of garden millipede appear to be flattened dorsoventrally due to the presence of lateral extensions called paranota, but upon close examination this millipede is quite circular in cross section, but the extensions result in the presence of lateral “keels,” giving these animals a flattened look. Adults typically bear 30 or 31 pairs of legs in males and females, respectively. They generally measure 18.5–22.2 mm in length and 2.0–2.5 mm in width. Initially, they are light brown but gradually attain a chestnut-brown coloration, and sometimes are bordered with yellow. Adults seem to live for about 2 months in the spring or summer, and like all millipedes, they are intolerant to dry conditions. Garden millipede is nocturnal. The biology on millipedes was reviewed by Hopkin and Read (1992) and Blower (1974), and a good synopsis including keys to North American families provided by Hoffman (1990). Chamberlin and Hoffman (1958) gave an annotated list of North American millipede species. Causey (1943) presented the life history of garden millipede.

Damage Millipedes occasionally damage seedlings of vegetable crops, but generally they are not primary pests. The destruction of seedling carrot, lettuce, and tomato was observed

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in Pennsylvania, for example, in both spring and autumn crops grown under shadecloth (Horsfall and Eyer, 1921). Reports of injury to some crops, such as damage to potato tubers in New York (MacLean and Butcher, 1934) were subsequently shown to be related to the presence of potato scab fungus (Butcher, 1936), with millipedes feeding preferentially on scab-infested tissue but not on healthy potato tubers. Once they commence feeding on the tuber, however, damage can be substantial. Indeed, in most cases of substantial injury, feeding by millipedes has followed damage by another insect or infection with a plant disease. Damage usually occurs within the context of cool weather and soil rich in organic matter. Millipede damage is not distinctive, so sometimes they are undeservedly blamed for damage. In Ontario, Canada, for example, Brunke et al. (2012) reported that damage to carrots and sweet potatoes was attributed to the millipede Cylindroiulus caeruleocinctus (Wood) by vegetable growers. However, research on the association of millipedes with damage to carrots showed that their abundance was inversely related to damage, and that wireworms were linked to the damage. There was some indication of millipede damage to sweet potatoes, however. Millipedes also burrow into fruits of crops such as cucumber, melon, squash, and tomato where the fruit comes in contact with the soil. Similarly, millipedes are sometimes found feeding within the heads of cabbage, cauliflower, and lettuce. Millipedes sometimes are viewed as a severe nuisance as a result of exceptional abundance in an inappropriate location such as in yards, homes, or commercial or food processing facilities. Millipedes can exist in tremendous quantities in the soil and become a problem only when they come to the surface and disperse as a group. This often occurs following abnormally large rainfalls (O’Neill and Reichle, 1970), though hot and dry conditions are also sometimes suspected to be a stimulus for dispersal (Appel, 1988).

Management Insecticides. Liquid, dust, and granule formulations of residual insecticides have been used with success to kill millipedes by application to soil, though not all products work equally well. On occasion, bait formulation consisting of wheat bran, molasses, and toxicant has been recommended, but this does not consistently meet with success. Barrier treatments of insecticides are sometimes recommended to disrupt the movement of millipedes from breeding areas (Rust and Reierson, 1977). Cultural Practices. Damage by millipedes often occurs when seedlings are grown under cool, early spring conditions or within cold frames or greenhouses where growth is not optimal. Delay in planting until weather is more favorable for plant growth is often recommended as a means to minimize injury by this, and other, seedling pests. Heavy

application of manure or other organic matter also predisposes seedlings to injury by millipedes and several other pests because they are attracted to organic matter, their principal food source. Conditions that favor high ambient humidity, including coarse mulch, vegetative groundcover, and thick weeds favor survival and activity of millipedes, so these should be removed (Appel, 1988).

CLASS ISOPODA—WOODLICE (PILLBUGS AND SOWBUGS) Common Pillbug

Armadillidium vulgare (Latreille) (Isopoda: Armadillidiidae)

Dooryard Sowbug

Porcellio scaber Latreille (Isopoda: Porcellionidae)

Natural History Distribution. Common pillbug, Armadillidium vulgare (Latreille), and dooryard sowbug, Porcellio scaber Latreille, are found throughout North America. They are cosmopolitan species, apparently originating in Europe, but now found throughout the world. They have adapted well to humans and human habitations and are often considered to be anthropophilic, but they also survive well in forests and grasslands, particularly if they can find shelter beneath logs and rocks. The pillbugs and sowbugs are collectively known as woodlice in Europe. This term conveniently depicts their relatedness and preferred habitat, and deserves wider recognition and use in North America. There are numerous species of woodlice in North America, though many seem to be immigrants from Europe. For example, Hatchett (1947) has reported 10 species of woodlice in Michigan, Drummond (1965) reported 28 species from Florida, and Hatch (1949) found 13 species dwelling within greenhouses in the northwest. In addition to common pillbug, A. vulgare (Latreille), and dooryard sowbug, P. scaber Latreille, other common and potentially important woodlice include A. nasatum Budde-Lund (Isopoda: Armadillidiidae); Oniscus asellus Linneaus (Isopoda: Oniscidae); P. laevis Latreille, P. dilatatus Brandt, Metoponorthus pruinosus (Brandt), and Trachelipus rathkei (Brandt) (all Isopoda: Porcellionidae). Host Plants. Woodlice generally feed on dead plant material, though they also accept dead animal remains and dung, and occasionally ingest bacteria, fungi, and living plants. They are best viewed as decomposers, similar to earthworms, breaking down plant material and mixing it with mineral particles to produce soil. However, they are not the first organisms to attack leaf litter, waiting until microorganisms have begun the degradation process. Also, they

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sometimes have the unfortunate habit of grazing on plants, particularly seedlings. Pierce (1907) has reported damage to seedlings of bean, cowpea, lettuce, mustard, pea, radish, tomato, and other crops in Texas by A. vulgare. In Australia, chickpea, faba bean, and lentil were not much affected by A. vulgare (Douglas et al., 2017), whereas alfalfa, canola, and oat were damaged. In a study of A. vulgare in natural California grasslands, pillbugs consumed green leaves of milk thistle, Silybum marianum, and ox tongue, Picris echioides, though dead leaves of both plants were greatly preferred (Paris, 1963). In Britain, the leaves, stems, and fruit of cucumber were reportedly damaged by A. nasatum (Goats, 1985). Natural Enemies. Woodlice in North America and elsewhere are parasitized by Melanophora roralis (Linnaeus) (Diptera: Tachinidae). Porcellio scaber is especially often attacked, with levels of up to 10% parasitism observed. Several other species of tachinids are known from Europe, with most being fairly specific in host range. Predation and cannibalism are known to occur, but it is uncertain whether these are important mortality factors in nature. Lizards, salamanders, shrews, spiders, centipedes, and ground beetles (Coleoptera: Carabidae) eat woodlice. An iridovirus has been found to occur in woodlice populations in California. In addition to causing a slight blue to purple discoloration in infected woodlice, the longevity of infected hosts is greatly reduced when woodlice are infected with iridovirus (Federici, 1980). Fungus, nematode, and protozoan parasites seem to be of little importance (Federici, 1984). Life Cycle and Description. Woodlice are found in the class Crustacea, whereas insects are in the class Insecta. Though superficially similar to insects because they have a rigid exoskeleton and jointed appendages, there are some important differences. As in insects, the body of woodlice is divided into three major regions—the head, which bears the antennae and mouthparts; the thorax or pereion which bears the legs but never wings; and the abdomen or pleon. The head bears two pairs of antennae instead of only one, as is found in insects, but one pair of the antennae in woodlice is greatly reduced in size and therefore is not often observed. The pereion (thorax) consists of seven segments instead of three found in insects, with each segment bearing a pair of legs ventrally. The pleon (abdomen) consists of six segments, but invariably it is much smaller than the pereion. The ventral surface of the pleon bears plate-like structures, and is an important site for gas exchange. A terminal pair of tail-like appendages, called uropods, may be located at the tip of the pleon. Uropods are present in sowbugs but are absent in pillbugs. Sowbugs cannot completely roll into a ball, though pillbugs are capable of this behavior. As the pillbugs can roll into a ball, they are sometimes called “rolly-pollys.” Woodlice can survive for longer than a year. However, Sorensen and Burkett (1977), working in Texas, noted that

individuals 2 and 3 years of age are not common, and older individuals are rare. Females tend to outnumber males during the breeding season, which extends from March until August. Males tend to outnumber females for the remainder of the year. Egg. The female woodlouse carries her eggs and young ones with her in a special compartment, called the marsupium, on the underside of her body. Fertilized eggs are inserted into the marsupium where the embryos (and later the young) obtain water, oxygen, and nutrients from a nutritive fluid called, appropriately, marsupial fluid. They may be up to 0.7 mm in diameter, and in some species over 100 eggs may be produced. The eggs persist for 3–4 weeks, then hatch, but the young remain in the marsupium for another 1–2  weeks before crawling out. They are only 2 mm long at this stage of development. Woodlice commonly produce offspring one to three times per year, with spring and autumn broods most common. Larger females produce more offspring. Juvenile. Hatchett (1947), working in Michigan, found that A. vulgare and P. scaber normally produced 20–40 young and one to three broods per season. Brood size was positively correlated to female size. The young are highly gregarious, and sometimes cannibalistic. Once they have left the female they molt, usually within 24 h, acquiring a seventh pereion segment. After an additional 14 days a second molt occurs, and a seventh pair of legs is produced. Thereafter, they do not change in morphology, other than an increase in size. The interval between molts is 1–2 weeks until the age of about 20 weeks, and molting continues irregularly for the remainder of their life, including adulthood. Molting occurs in two stages, with the posterior portion of the body shedding its old skin first, followed about 3 days later by the anterior portion.

FIG. 13.7  Pillbug. (Drawing by J. Capinera.)

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Adult. A length of 8.5–18.0 mm is eventually attained as woodlice reach adulthood. The width of the body is about half of its length. Woodlice are somewhat flattened and elongate-oval, seven pairs of legs and 13 body segments are apparent, and they have long, jointed antennae. Eyes are evident on the side of the head. They are brownish or grayish in general body color, though often marked with areas of black, red, orange, or yellow. Hatchett (1947) has reported that woodlice attained a larger size during summers when precipitation is above normal. Woodlice are nocturnal. A key to the genera of North American woodlice was published by Muchmore (1990). The key to British woodlice published by Sutton (1972) generally applies to North American woodlice found in crop, yard, and greenhouse situations because our damaging woodlice are of European origin. Good treatments of North American woodlice biology were given by Hatchett (1947) and Paris (1963). Sutton et al. (1984) have provided important information on biology of common pillbug, with Nair (1984) treating dooryard sowbug. Edney (1954), Sutton (1972), and Warburg (1993) also provided good overviews of woodlice ecology and physiology. Van Name (1936) described and pictured each of the aforementioned species, and in conjunction with supplements (Van Name, 1940, 1942) described most other woodlice in North America. Sutton (1972) provided culture methods. Nutritional ecology is reviewed by Zimmer (2002).

Damage Woodlice occasionally attack seedlings aboveground, feeding especially on stems and young leaves, and belowground, feeding on roots. Older plants may also be attacked, but this is generally of little consequence. Cool wet weather favors the activity of woodlice but inhibits the growth of seedlings, resulting in greater likelihood of seedling damage. Douglas et al. (2017) have noted that retention of organic matter associated with minimum tillage or stubble retention practices can lead to increased abundance of A. vulgare and higher incidence of seedling damage.

Management Woodlice can be sampled using objects that provide shelter, such as overturned flower pots, or items of food, such as bread dough. Woodlice are most common in soils with neutral or alkaline pH, good crumb structure, high organic matter content, and where soil bacteria and other macrodecomposers such as earthworms and millipedes flourish. They tend to be absent from acid and waterlogged soil. Woodlice are virtually absent from thoroughly tilled land due to either disturbance or lack of shelter. On the other

hand, straw or other coarse mulch provides good habitat for woodlice and can lead to crop damage. Though infrequently damaging, woodlice can be suppressed with liquid, granular, dust, and bait applications of insecticide to the soil around seedlings, or to protected habitats where woodlice tend to aggregate. Bait formulations developed for slugs and snails are sometimes recommended for woodlice.

CLASS GASTROPODA—SLUGS AND SNAILS Slugs

Deroceras, Leidyula, Limax, Milax spp. and others (Mollusca: Gastropoda: various families)

Natural History Distribution. Although North America has numerous native slugs, few of the native species are serious crop pests. Most of the slugs that damage crops have been introduced accidentally, mostly from Europe, but a few from the tropics. The major seaports on the east coast of North America were known to be infested with European slugs by the mid-1800s. Slugs are easily and routinely transported long distances in commerce, then spread relatively slowly by their own means. Thus, their distribution is sometimes discontinuous, often being limited to seaports and metropolitan areas. Of the troublesome slugs, only the veronicellid slugs, which are of tropical origin, have not been introduced from Europe. Getz and Chichester (1971) gave an account of introduced species. The slugs in the family Agriolimacidae are the most widespread, and in some areas most damaging (Douglas and Tooker, 2012). Foremost is gray garden slug, Deroceras reticulatum (Müller). It is found throughout southern Canada and in much of the United States, though it is uncommon in the Gulf Coast region. It was likely introduced from Europe. Marsh slug, Deroceras laeve (Müller) is also found widely, including the Gulf Coast area. Its native range is considered to be northern Asia and Europe. The western hemisphere is thought to be an invaded area, probably occurring in colonial times. It now occurs on all continents except Antarctica. A very similar species, Deroceras invadens Reise, Hutchinson, Schunack & Schlitt (formerly D. panormitanum) from southern Europe has spread widely and rapidly around the world, but until recently was overlooked due to its similarity to D. laeve. Its economic impact in North America is uncertain, but it is highly likely that some of the damage to gardens attributed to D. laeve or even D. reticulatum (O. F. Müller) is due to D. invadens.

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FIG. 13.8  Marsh slug. (Photo by L. Buss.)

FIG. 13.9  Gray field slug. (Photo by L. Buss.)

The family Limacidae also contains important species. Spotted garden slug, Limax maximus Linnaeus, occurs in the northern states south to Virginia and California, and in portions of southern Canada. Tawny garden slug, Limax flavus (Linnaeus), is widespread. Greenhouse slug, Milax gagates (Draparnaud), is the only slug in the family Milacidae that is important in North America. It is quite destructive to crops in California, and elsewhere in greenhouses, but is absent from the southeast. However, the subterranean slug, Tandonia budapestensis (Hazay), is a relatively recent introduction to North America from Europe that may assume serious status at some time in the future. The family Arionidae contains several slugs of importance. Black slug, Arion ater rufus (Linnaeus), is common in southern Canada, the northernmost areas of the United States, and in California. Banded slug, Arion circumscriptus Johnston, and garden slug, Arion hortensis Férussac, are pests in gardens and greenhouses, particularly in California. Among the Arionidae, Arion subfuscus (Draparnaud) is most widespread in the United States, and can cause damage in many habitats.

FIG. 13.10  Dusky arion. (Photo by L. Buss.)

Slugs in the family Veronicellidae are of little importance to most of North America, though quite damaging in more tropical locations. In Hawaii, Laevicaulis alte (Férussac), introduced from Australia or Southeast Asia but originating in Africa, is a pest of gardens. Other veronicellids found in Hawaii include Cuban slug, Veronicella cubensis (L. Pfeiffer), and bean slug, Sarasinula plebeia (P. Fischer). Veronicella cubensis has proven to be quite destructive in Hawaii, and also on several islands in the Caribbean and Pacific regions, and has been detected in California. Sarasinula plebeia is damaging in Central America and some Pacific Islands, but has not yet attained pest status in Hawaii (Kim et al., 2016). The Florida leatherleaf slug, Leidyula floridana (Leidy), was described from Florida but probably originated in Cuba. It affects gardens in Gulf Coast states, Mexico, and the Caribbean.

FIG. 13.11  Florida leatherleaf slug. (Photo by L. Buss.)

A slug native to eastern North America, Philomycus carolinianus (Bosc) (Philomycidae), is known as Carolina mantleslug. It can feed, grow, and produce eggs on diets of lettuce or carrots (White-McLean and Capinera, 2014). This species also feeds readily on several species of mushrooms. However, it is not known to be a garden pest.

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Host Plants. Slugs eat plant tissue, decaying organic matter, and occasionally animal tissue. Although they seemingly feed on a wide range of plants, their pattern of preferences is much like that of insects, and often is related to the allelochemicals found in the plants. Among the vegetable crops injured frequently are bean, beet, Brussels sprouts, cabbage, carrot, cauliflower, celery, corn, cucumber, eggplant, lettuce, lima bean, melon, pea, potato, radish, tomato, and turnip. Fruits such as strawberry, and to a lesser extent currant and gooseberry, may be injured. Slugs also consume barley, clover, oat, wheat and other field crops, but unlike the situation in Europe, they are not normally considered to be serious pests of these crops in North America. Slugs are opportunistic feeders. In addition to crops, slugs eat flowers, weeds, grasses, and other materials. Pallant (1972) has studied food preferences of Deroceras reticulatum in a British grassland, and found it to feed principally on the dominant grass species. However, it was discriminatory, not feeding on plants solely on the basis of availability. Also, it fed on mites, collembolans, and earthworms. Fox and Landis (1973) have reported predation by Deroceras laeve on aphids and moth eggs. Natural Enemies. Several invertebrates are potentially important natural enemies of slugs (Stephenson and Knutson, 1966; Godan, 1983). Tetanocera spp. marsh flies (Diptera: Sciomyzidae) paralyze and parasitize slugs. Several beetles, including ground beetles (Coleoptera: Carabidae), rove beetles (Coleoptera: Staphylinidae), and larval fireflies (Coleoptera: Lampyridae) are thought to be important. Protozoan pathogens of slugs, particularly Tetrahymena spp., reduce longevity and fecundity, and may cause death. Field studies frequently demonstrate about 10%–50% infection, and sometimes field colonies are eliminated by these pathogens. Fungi affect slugs, especially their eggs, but this is believed to be mostly an artifact of laboratory culture, and not very important under field conditions. Numerous helminths are associated with slugs, including trematodes, cestodes, and nematodes; some helminths are parasitic on slugs, but many use slugs as intermediate hosts and eventually infect livestock and humans. The nematode Phasmarhabditis hermaphrodita (A. Schneider) (Rhabditidae) has become commercially available as a biological control agent of certain slugs, and has been available in Europe for many years. Because it did not occur naturally in North America, sale of the nematode was prohibited in the United States. However, it has been found to be present in some slugs in California, as has a native Phasmarhabditis species heretofore unknown (Tandingan De Ley et  al., 2016), so these likely will become available for use in North American agriculture. Phasmarhabditis hermaphrodita infects some slugs, including Deroceras laeve, Deroceras reticulatum, and Leidyula floridana, but some of the important invasive species from Europe (Arion hortensis, Arion

subfuscus, Limax maximus) are not susceptible to infection (Speiser et al., 2001; Grewal et al., 2003). Vertebrates also consume slugs, particularly birds and toads. Domestic poultry is sometimes suggested as a means of reducing slug abundance. Life Cycle and Description. Slugs are molluscs, but unlike the closely related snails, their shell is vestigial and internal. Their eyes are rounded knobs borne on filamentous stalks that may be retracted into the head. Slugs secrete from pores in the body, and especially from the anterior ventral surface of the body, a slimy secretion called mucus. Located behind the head is a swelling, or mantle, which contains the thin shell and a respiratory opening or breathing pore. Slugs have phases of growth that are marked by changes in their reproductive status, but often are not indicated outwardly other than by a gradual increase in size. The first phase, or infantile period, marks the time from egg hatch to differentiation of the sexual organs. In the second phase, or juvenile period, the male genital tract develops to sexual maturity. In the third phase, or adult period, sperm are produced and the slug is ready for copulation. Later in this period the female genital tract develops and eggs are produced. In the final phase, or senile period, the slug senesces and the genital tracts deteriorate. Unlike insects, slugs are hermaphrodites, each individual possessing both male and female sexual organs. Nevertheless, self-fertilization is rare, and slugs normally pair for copulation. The mating process normally requires only a few minutes, followed by a period of several days to weeks before oviposition commences. Egg. Individuals slugs produce about 100–200 eggs during their life span. Eggs are deposited in small cavities in the soil, usually in small clusters. They are spherical or elliptical. In the case of Deroceras reticulatum, they are translucent, bluish white, and slightly iridescent. They measure 2.0–3.0 mm in length and 1.7–2.0 mm in width, and sometimes bear a nipple-like extension. The clusters of eggs are held together with a transparent secretion. Juvenile. Young slugs initially are transparent or nearly so, but soon take on a pinkish color, and then become darker as they feed. Slugs have a protracted period of development, and development ceases in the absence of favorable environmental conditions. Juveniles do not differ in appearance from adults, except for being smaller in size. Adult. Slugs attain reproductive maturity within a few months, but they continue to grow in size and continue to reproduce, sometimes for a year or more. Longevity varies greatly among slugs, with D. reticulatum living 9–13 months, Arion spp. living 7.5–12 months, and Limax spp. living 24–36 months.

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Discrete generations generally are not present, with all stages present during much of the year. Up to three overlapping generations may be present simultaneously. In England, Hunter (1968) has reported that D. reticulatum had two generations per year, A. hortensis had one generation annually, and Tandonia budapestensis required 2 years for a single generation. Others have observed different patterns of abundance. In Ontario, for example, Rollo and Ellis (1974) have observed only one generation of D. reticulatum annually in corn fields, with overwintering in the egg stage, and egg production only in the autumn. During the summer months, most slug populations appear to be low, but adults emerge following heavy rains, suggesting that they disperse deep into the soil in response to dry conditions. In the same region but in uncultivated areas where vegetation holds an insulating layer of snow, all stages overwinter. Slugs are not well adapted to life on land, surviving only where water is fairly abundant. Gray garden slug, for example, has a body water content of over 80%, and its eggs of over 85%. Loss in body water content of 20% results in death. Water is replenished both by drinking and by direct absorption of water through the skin. Thus, cool and wet environments favor the existence of most slugs. Feeding and dispersal are limited principally to periods of darkness, though the period of peak activity varies among species, and some remain active into the morning hours. Slugs may also be active during daylight hours, if it is cloudy or foggy. Some slugs avoid the soil surface, and remain belowground. Although slugs can dig through the soil, their activity and damage potential are greatly favored by natural spaces in the soil, as results when heavy soil is plowed and left in large aggregates. The lack of a shell likely facilitates tunneling in the soil, relative to snails, and some species remain buried in soil when not feeding, even in the presence of a saturated environment. Comprehensive treatments of pest slugs and snails, including culture, were presented by Godan (1983) and Port and Port (1986). Barker (2002) edited a useful treatment of mollusk pests of crops. A key to common slugs is given in Appendix A. More complete keys to pest molluscs were given by Kono and Papp (1977) and McDonnell et al. (2009) for California; Stange (1978), Capinera (2017), and Capinera and White (2017) for Florida; Thomas et al. (2010) for Kentucky; Godan (1983) for Europe; Forsyth (2004) for British Columbia; and Grimm et al. (2009) for Canada. Burch (1960) has described molluscs of quarantine significance to the United States. Burch and Pearce (1990) have published a key to North American genera of slugs and snails. Barnes and Weil (1944) have provided brief description of, and a key to, British slugs. Useful accounts of D. reticulatum and L. maximus were given by Hawley (1922a) and Barker and McGhie (1984), respectively. Lovett and Black (1920) have described slugs in Oregon. Other good sources of information on slugs were provided

by Henderson (1989, 1996). Note that the names of slugs and snails have been changed frequently, making it difficult to rely on identities of organisms when using publications more than a few years old.

Damage Slugs cause direct injury by feeding on plant tissue, causing serious defoliation injury to leafy vegetables such as lettuce, and to seedlings of many vegetables. Seedlings are often cut off at the soil surface by slugs. They also feed below-ground on roots and tubers of beet, carrot, potato, radish, and turnip, with potato most frequently injured. Slugs usually feed on the surface of potato tubers and tomato and eggplant fruits, but sometimes dig deeply into the flesh of these vegetables. Contamination of vegetables with slugs, or with their slime trails, can be as damaging as defoliation in commercial vegetable production. Slugs can also serve as vectors of bacterial and fungal diseases of plants (Dawkins et al., 1985, 1986). Although there is a direct relationship between slug abundance and crop damage, crops vary greatly in their susceptibility to damage (Barratt et al., 1994). Slugs can also cause damage indirectly. Molluscs (including slugs) are intermediate hosts of rat lungworm, Angiostrongylus cantonensis. This is a nematode parasite of rats, but it can also affect primates, including humans. The situation is exactly the same as with snails (see section on snails).

Management Sampling. Direct assessment of population density is difficult because slugs are nocturnal, and usually hide below-ground during daylight hours. Damage assessment is a useful approach for plants that have already emerged from the soil, but is useless for assessment of slug populations before plant emergence. Soil-washing techniques can be used to separate slugs from soil (Rollo and Ellis, 1974), but this approach is very labor intensive. Therefore, traps are usually used to census populations. The most common type of trap uses a board or inverted flower pot bottom as an attractive shelter under which slugs can aggregate during the day. Catches can be increased by baiting the trap with grain or such grain products as bran. The bait can also be poisoned to ensure that the slugs will not disperse. Beer is highly attractive to slugs, and they often enter containers with beer and drown (Smith and Boswell, 1970). The efficiency of traps suffers from dependency on slug mobility, which is influenced by weather. For maximum slug activity, soil temperatures at a depth of 30 cm should exceed 7°C, air temperature should attain at least 9°C, and soil moisture content should be at least 26% (Young et  al., 1993). The accuracy of traps for estimating slug density has been questioned (Young, 1990a), but it remains the most convenient and widely used method.

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Pesticides. Few pesticides cause consistent and high levels of slug mortality. Metaldehyde and methiocarb (mesurol), and to a lesser extent other carbamate insecticides, traditionally have been recommended and formulated as toxic baits (Airey, 1986). Metaldehyde is much less available now due to problems with nontarget toxicity to wildlife and pets. Also, after temporary immobilization slugs may recover from low or moderate levels of exposure to metaldehyde, especially in the presence of moisture (Cragg and Vincent, 1952). The more recent availability of iron-based molluscicide bait formulations (iron phosphate and ferric sodium EDTA) provide moderately effective, less toxic (to nontarget animals) alternatives for slug control. They stop the slugs from feeding as soon as they are ingested, preventing further plant damage, though it takes days for the slugs to actually perish. Sulfur is also a relatively safe (to nontarget animals) ingredient for mollusk baits, though it is less effective than the iron-based products. Boric acidbased bait has only limited toxicity to molluscs. Recent research on molluscides includes work by Hollingsworth and Armstrong (2003), Capinera (2013, 2018b), and Capinera and Guedes Rodrigues (2015). Although pesticides may be applied as sprays and dusts (Howitt and Cole, 1962), they are usually formulated as baits (Prystupa et  al., 1987; Hammond et  al., 1996). The dispersal of pesticide on bait is more effective than dispersal without bait (Barnes and Weil, 1942). Slug species vary in their propensity to accept bait (Airey, 1986), though this variable is usually ignored. No pesticide is completely satisfactory because effectiveness varies with slug species, weather it is the need for the slug to be feeding on the soil surface, attractiveness of the bait, and ability of the mollusk to detoxify the pesticide. Slugs are not very mobile, so broadcast applications are most effective. The application of molluscicides to seed to protect seed and seedlings from slugs has been evaluated and can be effective, but the hazard to birds is high. Light irrigation is usually recommended prior to application of bait formulations because the moisture encourages foraging by the molluscs. Heavy irrigation after bait application is discouraged as it encourages deterioration of the bait. Cultural Practices. Slugs are a greater problem on heavy soil, though in the presence of irrigation slug populations can increase on light soils also. The use of drip irrigation rather than overhead sprinkling systems lowers the ambient humidity and makes an area less suitable for slugs. Slug problems are greatest if soil remains in large aggregates, which provide slugs with shelter. If soil is finely tilled or compacted, it reduces the amount of shelter available to slugs, and plant seedling survival is higher (Hunter, 1967; Stephenson, 1975). Sanitation also is important. In the home garden, it is important to eliminate boards, stones, debris, and weedy areas that may provide shelter for slugs. In commercial

agriculture, residual organic matter contributes greatly to slug problems by providing both food and shelter. Thus, mulching and minimum tillage practices exacerbate slug problems (Tonhasca and Stinner, 1991; Barratt et al., 1994; Hammond et  al., 1996, 1999). Planting crops into former pastureland can also be hazardous. For planting beds or particularly prized specimen plants, copper screen or metal flashing can be used as a barrier. Copper reacts with the mucus produced by slugs, producing an electric charge. The use of row covers to protect crops from insects have received considerable study, and in some cases may protect plants against slugs. However, slugs can burrow through soil and penetrate such barriers, perhaps explaining the failure of covers to protect vegetables in New Zealand (Evans et al., 1997). Slugs are said to be reluctant to cross barriers of diatomaceous earth, but there are no scientific data to support this claim, and some to refute it. On the other hand, hydrated lime or sulfur affect slug behavior and can be used to create barriers, though once the barrier material is wetted by rainfall, irrigation, or soil moisture the repellent properties are compromised, so they are not very practical (Capinera, 2018b). Copper hydroxide fungicide, Bordeaux mixture (a copper sulfate and lime mixture used mostly as a fungicide), and also copper sulfate alone, can deter slugs, but plant toxicity may occur with copper sulfate. Essential oils are available commercially as slug and snail repellents, but the limited research conducted so far shows that they vary from inactive to very active as plant protectants. Capinera and Dickens (2016) and Capinera (2018a) have provided a comparative data on copper-based fungicides that have repellent and feeding deterrent properties. Biological Control. Although there are numerous microbial, invertebrate, and vertebrate natural enemies of slugs, few have shown much promise for manipulation to achieve biological suppression. One exception seems to be the nematode Phasmarhabditis hermaphrodita (Nematoda: Rhabditidae). This nematode kills slugs within 1–3  weeks of application, kills a wide range of species, and can be cultured on artificial medium (Wilson et al., 1993a,b, 1995). A ground beetle, Abax parallelepidedus Piller and Mitterpacher (Coleoptera: Carabidae), feeds readily on slugs and is under investigation in Europe for the suppression of slugs in the field and in enclosures; it also seems amenable to mass culture (Asteraki, 1993; Symondson, 1994).

Snails

Bradybaena, Cepaea, Cornu, Rumina, Zachrysia spp. and others (Mollusca: Gastropoda: various families)

Natural History Distribution. Native snails are widespread in North America, but usually only imported snails are of consequence

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as plant pests. Many of the snails are of European origin, but some originated in tropical countries. Unlike the situation with the closely related slugs, which were accidental introductions, the snails were in many cases (at least the larger species) deliberately introduced to North America to serve as a source of food. Unfortunately, such introductions were made without legal and environmental considerations, and have proved to be costly. Within North America, the redistribution of snails results mostly from the movement of nursery stock. The snails in the family Helicidae have proven to be most adaptable to North America, and most damaging to plants. The brown garden snail, Cornu aspersum O. F. Müller (also known as Helix aspersa, Cantareus aspersus (or Cryptomphalus aspersus)) was introduced to California in the 1850s to serve as a source of escargot. It has adapted well to California and is very troublesome as a pest of crops and ornamentals. It also occurs now along the west coast north to British Columbia, and occasionally is reported from southeastern states and along the east coast north to New Jersey (though these may be interceptions or transient introductions, not permanent establishments), but has not developed the serious pest status found in California. Similarly, the singing snail, Helix aperta Born, apparently was introduced deliberately into southern California and Louisiana, where it damages vegetable and flower crops. The milk snail, Otala lactea (Müller), another popular food snail, is established in most of the southern states, but has been seriously damaging only in California. The brown-lipped snail, Cepaea nemoralis (Linnaeus) and the white-lipped snail, Cepaea hortensis (Müller), though edible, were apparently introduced to northeastern North America for their ornamental value. However, some have argued that C. hortensis is an endemic species. The Cepaea spp. have proven to be more of a nuisance than an actual pest. The white garden snail, Theba pisana (Müller), was introduced to southern California some time before 1918, and proved to be very damaging. Though apparently eradicated, it occasionally reappears. Many other species have been introduced but have not developed into serious pests, or have been successfully eradicated. A history of helicid introductions to North America was given by Mead (1971a). Family Achatinidae contains at least one snail of note. The giant African snail, Lissachatina (Achatina) fulica Bowdich, which is native to Africa and has become established throughout Southeast Asia, is well established in Hawaii, where it was introduced in the 1930s. Also, it was introduced into south Florida in 1966, and successfully eradicated. It was rediscovered in southern Florida 2011 after a probable reintroduction from the Caribbean, where it occurs on several islands. There are several features about this snail that make its presence threatening. This snail has a very high reproductive capacity (averaging

5000 eggs per snail), and a small percentage of snails can reproduce without mating (Dickens et al., 2018). Thus, the transport of even a single snail to a new region could result in development of an infestation. Based on reports from around the world, it has a wide host range. Perhaps most alarming for the United States, these snails not only thrive in warm climates, but can burrow into the soil and survive cold weather. This includes weather found at a latitude of about 40 degrees, which may include states such as New Jersey and Colorado.

FIG. 13.12  Giant African land snail. (Photo by L. Buss.)

The decollate snail, Rumina decollata (Linnaeus), originated in the Mediterranean region, but has been introduced to many areas of the southern United States, and now occurs from California to Florida. Small colonies occur as far north as Pennsylvania. It is perhaps the most interesting of the “pest” snails, because it is omnivorous, and quite effective at killing small brown garden snails. It is considered to be beneficial in areas suffering from brown garden snail infestations, but is a minor pest of plants in other areas. Currently, Rumina is placed in the family Subulinidae. The family Bradybaenidae contains one species of concern in North America, Bradybaena similaris (Rang), which is also known as Asian trampsnail. Originally found in Asia, it has spread to the Caribbean and South America, and now occurs in Hawaii and the Gulf Coast region. However, though it is only a minor pest of vegetable, ornamental, and citrus plants, it is of regulatory concern. Among the Camaenidae (Pleurodontidae), Zachrysia provisoria (Pfeiffer), brown Cuban snail, is an important plant pest in southern Florida (Capinera, 2013). Like many other snails, it was introduced deliberately as a potential source of food. Because the source was Cuba, it is also known as brown Cuban snail. It occurs widely in the Caribbean region. It occurs mostly on ornamental plants but can feed on vegetables.

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FIG. 13.13  Cuban brown snail. (Photo by L. Buss.)

Host Plants. Snails tend to be omnivorous, consuming decaying organic matter, foliage, and fruits. Not all vegetation is consumed equally, and preferences vary among snail species. Among the vegetables known to be severely injured by snails are bean, beet, cauliflower, chicory, cucumber, endive, lettuce, pea, pumpkin, and spinach. Flowers and other ornamental plants are frequent hosts. Acceptability of different plant hosts for Zachrysia provisoria is given by Capinera (2013) and suitability of plants for Lissachtina fulica by Lange (1950) and Dickens et al. (2017). Natural Enemies. The natural enemies of snails are substantially the same as those affecting slugs (see section on slugs). However, predatory snails can also be important mortality factors affecting phytophagous snails. The rosy wolf snail, Euglandina rosea (Férussac) (Spiraxidae), occurs in the southern United States and Hawaii, as well as in other countries where giant African snail occurs. Its origin is Central America, but it was redistributed widely due to its appetite for giant African snail. It feeds on numerous species of snails, and small slugs, and threatens rare and endangered species in some locations. The decollate snail, Rumina decollata (Linnaeus) (Subulinidae), also occurs widely, and as noted previously, can feed on plants and native snail species. Life Cycle and Description. Basinger (1931) has provided an account of brown garden snail that illustrates the basics of snail biology, and the following description is based on this. Terrestrial snails are typical molluscs, dwelling in a shell that consists principally of calcium carbonate. The shell grows in size as the snail matures by the deposition of more shell at the mouth of the shell opening. Snails secrete an acidic substance from their foot which dissolves calciferous materials and allows them to extract calcium from the soil, though they also obtain calcium by direct ingestion. Lack of calcium in the environment limits the health and growth of snails. The fleshy body tissue may be withdrawn into the shell for protection, but part is extruded during normal bodily functions like eating and moving.

The eyes are at the tips of filamentous tentacles which can be extruded or retracted back into the body. Snails are better able to withstand long periods of dryness than slugs because they can withdraw into their shell. They may withdraw and become dormant for months during periods of dryness. They may secrete several layers of dried mucus over the opening of the shell to aid water retention during periods of inactivity. The body also secretes slimy mucus which aids the snail in movement. Snails are active mostly at night, and may hide during daylight hours, often beneath rocks, refuse, or dense vegetation. These sites also are preferred for overwintering. Some species, however, remain in the open and attached to their food plant or another substrate even during the day, secure because they are protected by their hard shell. Most snails, like slugs, are hermaphroditic, containing the sexual organs of both sexes within a single body. However, they normally mate with another snail, and self-fertilization is rare. Mating requires 4–12 h. Though copulation is initiated at night, snails are often observed copulating in the morning due to the duration of fertilization. Both individuals deposit eggs, beginning 3–6 days after mating. Damp, loose soil is preferred for oviposition. Egg. Snails deposit eggs in nest holes that they dig in loose soil. The snail uses its foot to shovel soil upwards, digging a hole about 2–3 cm deep and 1–2 cm in diameter. The eggs are deposited singly but adhere to one another, forming a loose mass. The number of eggs deposited at one time varies from about 30 to 120, averaging 86. The snail fills the nest hole with soil after oviposition and deposits excrement atop the hole. The entire oviposition process may require 24 h for completion. Eggs may be deposited as often as monthly under ideal environmental conditions, but no eggs are deposited during the winter months even in California. Thus, total fecundity is estimated at 400–450 eggs annually. The eggs hatch in about 2 weeks. Juvenile. The young snails are miniatures of the adult form, but measure only about 5 mm in diameter. The snail may require 2 years to reach maturity. Snails add calcareous material to the opening of the shell continuously; as the shell is enlarged, it spirals around the old shell tissue. Brown garden snails attain a diameter of 16–20 mm within 1 year, but 26–33 mm by the second year. In addition to becoming reproductively dormant during cold weather, they also are inactive during dry weather. Adult. The snails are a mixture of grayish yellow and brown, with the brown concentrated into spiral bands. The adult typically possesses 4–5 whorls, and may attain a maximum diameter of 38 mm. Brown garden snails are nocturnal. They feed on organic matter in the soil, bark from trees, and especially on vegetation. Nearly anything growing in

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a vegetable or flower garden can be consumed. They normally feed only within the temperature range of 5–21°C. Comprehensive treatment of pest snails, including their culture, was presented by Godan (1983). A key to common snails can be found in Appendix A; more complete keys were provided by Burch (1960), Kono and Papp (1977), and Godan (1983). Burch and Pearce (1990) have published a key to North American genera of slugs and snails. Useful discussion of helicid snails in California was given by Gammon (1943); the helicids Achatina and Rumina were treated by Mead (1971a,b). An interesting treatment of white garden snail and the eradication campaign developed for was provided by Basinger (1927). Common predatory snails in Florida were described by Auffenberg and Stange (1986). Rumina decollata was discussed by Fisher and Orth (1985).

Damage The direct damage caused by snails is very similar to that inflicted by slugs, though in most of North America snails are much less damaging. Snails may feed on plant tissue, chewing irregular holes in leaves and causing serious injury to leafy vegetables such as lettuce and seedlings of many vegetables. Contamination of vegetables with snails, or with their slime trails, is often considered to represent damage. Commercial vegetables normally are not subject to attack by snails. Snails can also cause damage indirectly. Snails are intermediate hosts of rat lungworm, Angiostrongylus cantonensis, a nematode parasite of rats. Nematodes can be released in the feces of rats, which may be consumed by molluscs, and in turn can contaminate vegetables. Humans who consume vegetables that are not cleaned adequately or cooked can be infected with these nematodes. Primates, including humans, are not natural hosts of this nematode, but the nematodes can nevertheless cause injury. When large numbers of nematodes are ingested by humans, damage to the human brain and spinal cord can occur, producing meningitis-like symptoms. This nematode, long known from Asia and Central America, now occurs in the United States (Capinera and Walden, 2017; Stockdale-Walden et  al., 2015, 2017) and is found in numerous species of terrestrial molluscs. Thus, where vegetables might be infested by molluscs, it is important to wash the vegetables thoroughly if they will be eaten uncooked. So far, this is a problem for humans only in tropical climates, including Hawaii, but the potential exists for it to occur elsewhere as the nematode infests molluscs in the southeastern United States and possibly elsewhere.

Management The approaches to management of snails is not significantly different from that used for slugs (see section on slugs).

CLASS SYMPHYLA—SYMPHYLANS Garden Symphylan

Scutigerella immaculata (Newport) (Symphyla: Scutigerellidae)

Natural History Distribution. Garden symphylan has a wide distribution in North America and can be a pest under both greenhouse and field conditions. It is most common and damaging along the west coast, from southern British Columbia to southern California. However, at times it has been damaging in other western states such as Idaho, Utah, and Colorado, as well as most of the midwestern states and New England and the Great Lakes region of Canada. Garden symphylan is infrequent in arid regions such as the Desert southwest and Great Plains. It also is rare in the southeast. This species is not the only symphylan to cause plant injury, but most symphylan damage in crops is justifiably attributed to this species (Waterhouse, 1970). Scuigerella immaculata also occurs in Europe and North Africa, and has been introduced into Argentina. It likely is a native of Europe. Host Plants. Garden symphylan has a wide host range. Among vegetables fed upon are asparagus, beet, broccoli, carrot, cauliflower, celery, corn, cowpea, cucumber, eggplant, garlic, lettuce, lima bean, onion, parsley, pea, pepper, potato, pumpkin, radish, rhubarb, snap bean, spinach, and tomato. Not all vegetable crops are equally susceptible, with asparagus, snap bean, and lima bean commonly mentioned as damaged. Umble and Fisher (2003a) have compared the suitability for several vegetable crops in pot tests and reported that population growth was greater on spinach than on tomato, sweet corn, or potato. Potato was significantly less suitable than the other crop plants. Field crops injured include clover, dry beans, field corn, hop, and sugar beet. Flowers and other ornamentals also are affected. Garden symphylan also feeds on yeast, algae, dead insects, and decaying vegetation. Natural Enemies. Various centipedes (Chilopoda) and mites have been shown to devour garden symphylan (Wymore, 1931; Waterhouse, 1969; Berry, 1973; Peachey et  al., 2002). Predatory beetles (Coleoptera: Carabidae) have been suggested to be mortality factors, but there are no data related to their effect on symphylans. Fungi have been reported to decimate symphylan populations under laboratory conditions (Getzin and Shanks, 1964), but have not been observed to cause mortality in the field. Life Cycle and Description. Symphylans are not true insects, belonging instead to the class Symphyla in the phylum Arthropoda. They resemble centipedes, and the garden symphylan has often been called the “garden centipede.” Nevertheless, they are not centipedes, which belong to the

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class Chilopoda, and are best thought of as insect relatives. In most respects their biology is like that of insects. Garden symphylan prefer cool soil temperature, about 18°C, but the temperature range of 12–20°C is suitable. They may be found near the soil surface, and to depths of over 1 m. Symphylans retreat to greater depths in the soil as the soil warms in the summer, but migrate upwards again in the autumn. They are more likely to be found near the surface if growing plants are present, though if the environmental conditions are unfavorable, they may disperse for episodes of molting and egg laying, and return only to feed (Edwards, 1959, 1961). Symphylans apparently line the runways or channels through which they travel in the soil with a layer of fine silk. They seem unable to dig their own passageways through the soil. Rather, symphylans inhabit earthworm burrows, natural crevices, and openings in the soil left by the decay of roots. Packed soil is not favorable. Soil moisture levels affect symphylans, with optimal levels for both plants and symphylans about the same. Relative humidities of less than 95% are deleterious to survival (Waterhouse, 1968). Garden symphylans generally require about 90 days for a complete generation, but temperature may affect development rate considerably. Egg. The eggs are deposited in tunnels in the soil. They are deposited in small clusters of 4–25, but usually 9–12 eggs. They are spherical and measure about 0.5 mm in diameter. They bear minute ridges on their surface. Initially, the eggs are white, but darken to brownish or greenish as they mature. Eggs can be found through most of the year in California, but are most abundant in the spring. At this time they hatch in 7–10 days. They were reported to hatch in 39.8, 24.9, and 12.8 days when held at 10°C, 20°C, and 25°C, respectively (Berry, 1972). Immature. Young symphylans measure about 0.75 mm in length, and have 10–11 dorsal body plates, six pairs of legs, and six antennal segments. The posterior segment also bears a pair of cerci, and the body is sparsely clothed with long hairs. Young symphylans remain near the site of hatching, and feed little or perhaps not at all, until the first molt. Additional pairs of legs, body segments, and antennal segments are added as the symphylans molt and grow. The immatures have seven instars. The number of antennal segments is typically about 6, 13, 15, 17, 19, 22, and 25 for instars 1–7, but the numbers are somewhat variable (Waterhouse, 1968). The duration of the first instar is only 2–3  days, but the other instars persist from 10 to 14 days. Development rates are highly variable, but Berry (1972) has reported total instar development time are about 120.0, 65.8, and 40.4 days at 10°C, 20°C, and 25°C, respectively.

FIG. 13.14  Garden symphylan. (Drawing by J. Capinera.)

Adult. The adults vary in length from 5 to 8 mm. They have 12 pairs of legs and 14 body segments, and though they continue to molt after becoming adults they do not acquire additional body segments or appendages. They continue to add antennal segments up to at least 50–59 in instar 24 (Waterhouse, 1968). Young adults, however, usually bear about 25–40 bead-like antennal segments. Compound eyes are absent, and these insects depend heavily on their long antennae for orientation. The anal body segment bears a pair of pointed cerci, each producing silk. The color of garden symphylan is normally white, but is affected by the food plant on which it feeds. Thus, it acquires a pink tint when feeding on radish, red when feeding on garden beet, and brownish when feeding on decaying vegetation. Duration of the adult is probably 9–10 months but there are records of individuals living for more than 4  years. The sexes are difficult to distinguish; Filinger (1931) has provided useful characters. Egg production is reported to cycle, with a 2-month period of high egg production followed by a 3–4-month period of relatively low egg output, and this is repeated several times over the course of the symphylan life span (Berry, 1972). Good summaries of garden symphylan biology were given by Filinger (1931) and Wymore (1931), but the most complete description was provided by Michelbacher (1938). Michelbacher (1942) provided a key to the Scutigerella spp. Culture methods were given by Shanks (1966) and Ramsey (1971).

Damage Both immatures and adults feed on roots and root hairs. Symphylans chew into the large roots, creating small holes

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or pits, but completely consume fine roots. Warty and corky growths may appear on carrot, beet, and other vegetables that develop below-ground. Heavy infestations result in severe root pruning. Plants are stunted, grow slowly, and may wilt or even perish. When asparagus is blanched by throwing soil over the emerging shoots the asparagus stems may be damaged. Foliage is not normally injured, because symphylans do not leave the soil. However, when foliage comes in contact with soil it may be eaten. In greenhouse studies, Eltoum and Berry (1985) have found that though five symphylans per young bean plant did not affect plant growth, densities of 10 or 20 symphylans disrupted the physiology and weight gain of the seedlings.

Management Sampling. Symphyla move in the soil, but do not tunnel. Instead they travel through soil along insect and earthworm tunnels, and seasonal cracks. The distribution of garden symphyla is patchy, and often only parts of a field are affected. The problem, once established, tends to persist, but to shift slightly, possibly because the symphyla do not move long distances laterally. They readily move vertically, however, moving to the soil surface region (plant root zone) to feed, by deeper into the soil when molting. Sampling garden symphylans traditionally has been accomplished by digging soil from a number of spots in crop fields prior to spring cultivation, then breaking up the soil aggregates and sifting through the soil to make a visual count. A dark background is usually used for the sorting because the white-colored symphylans are easier to see on a dark background. Sampling can be done after tillage as well, but it is advisable to sample deeply in this case, as the tillage may drive the symphylans deep into the soil. Bait sampling methods have also been developed. Bait samples are generally much faster to take than soil samples, but they are also more variable and more sensitive to factors such as soil moisture, temperature, and the presence of vegetation (Umble and Fisher, 2003b). For bait sampling, half of a potato or beet is placed on the soil surface and sheltered with a protective cover (e.g., a white pot). The bait and soil beneath it is examined for symphylans after 1–3 days. The number of baits used per site is usually 35–50. Insecticides. This arthropod has proven difficult to suppress with many chemical insecticides. Fumigants applied before planting can be effective but they are costly and may kill only the individuals near the surface. Insecticides mixed into the soil at planting also can be effective (Howitt, 1959; Howitt et al., 1959; Sechriest, 1972). Broadcast rather than banded treatments are preferred by some, but adequate

suppression is usually attained with the banded treatments (Gesell and Hower, 1973). Dipping of transplants has been proposed as a method of insecticide delivery (Berry and Crowell, 1970). Persistent materials tend to be most effective, and some are repellent, providing another means of protecting roots (Joseph, 2015). Cultural Practices. Soil type affects symphylan abundance and damage to crops. Loam soils are more suitable for symphylans than sandy and clay soils. High organic matter, good moisture holding capacity, and the ability of the soil to form crevices favor the occurrence of symphylans (Edwards, 1958). Sandy soil is generally not suitable, but if organic matter is added to the soil, suitability for symphylans increases. As moist soil favors symphylans, cultivation of the soil to increase surface drying can drive the symphylans deeper into the soil and decrease feeding on plants. Crop management also affects symphylan abundance. Historically, garden symphylans are serious greenhouse pests, but they are largely limited to greenhouses where plants were grown in the ground, or where pots were in direct contact with soil. If plants are grown in sterile soil, or not in direct contact with soil, few problems can develop. Flooding can destroy symphylans, and in California flooding for 1–2 weeks during the summer is recommended before planting the autumn crops. Asparagus is quite tolerant of flooding, and in California good control of symphylans has been attained by submerging fields for 3–4 weeks during the winter (Wymore, 1931). Despite the ability of symphylans to feed on other soil organisms, their populations can decrease dramatically if less suitable crops are planted. Rotation of a field to potatoes is most effective for reducing populations, but grain crops can also be beneficial for decreasing symphylan abundance. Peachey et al. (2002) have concluded that the nature of the cover crop was more important than spring tillage in reducing symphylan populations; oat residue made a better cover crop than barley residue. Young plants are more susceptible to damage than old, especially in the case of less favorable crops such as tomato. Growers can reduce the effect of symphylan feeding by planting tomato transplants rather than direct seeding the crop. However, this is not effective for more favorable hosts such as broccoli and eggplant. There has been only limited work on the identification of plant varieties resistant to garden symphylan. Simigrai and Berry (1974) have observed significant differences among commercially available cultivars of broccoli, and suggested that this approach might hold some promise in other crops as well.

Appendix A   625

Beetle

Head

Forewing (elytron in beetles)

Antenna Pronotum Scutellum

Abdomen

Grasshopper Antenna Front leg Head Compound eye

Tarsus Tibia Femur Coxa

Hind wing

Forewing (tegmen in grasshoppers)

Pronotum Hind wing Middle leg Hind wing Hind leg

Femur of hind leg Abdomen

Bug Antenna Front leg Head Compound eye Pronotum

Tarsus Tibia Femur

Forewing (hemelytron in bugs)

Scutellum Middle leg

Hind leg Hind wing Abdomen

Caterpillar

Thorax

Abdomen

Thoracic legs

Abdominal prolegs

Head Mandible

Beetle, grasshopper, bug, caterpillar.

Anal prolegs

626   Appendix A

Aphid.

Representative head of chewing insect.

Representative head of piercing-sucking insect.

Appendix A

Keys to Selected Groups of Pests KEY TO MAJOR ORDERS OF INSECTS AFFECTING VEGETABLE CROPS 1. Insect bearing wings .................................................. 2 Insect lacking wings ................................................. 15 2. Front wings entirely or partly membranous, with veins evident .............................................................. 3 Example of insect with body covered with scales’ mouthparts long and tubular. (Drawing by J. Capinera.)

Example of insect with membranous wings and wing vines: aphid. (Drawing by J. Capinera.)

Wings lacking scales .................................................. 4 4. Only one pair of wings present, hind wings reduced to small knob-like structures Flies; order Diptera

Front wings hard or thickened, veins often lacking ...................................................................... 11

Insects with only one pair of wings: flies. (Drawing by J. Capinera.) Example of insect with hardened front wings: beetle. (Drawing by J. Capinera.)

3. Wings covered with scales, mouthparts long and tubular but coiled Butterflies and moths; order Lepidoptera

Two pairs of wings present, though hind wings may be smaller ........................................................... 5 5. Wings narrow and bearing long fringe around margin; body 5 mm long ....... 6 6. Chewing mouthparts; tarsi with five segments

Opague, veinless wings: whitefly. (Drawing by J. Capinera.)

Example of insect with chewing mouthparts: grasshopper. (Drawing by J. Capinera.)

Ants, bees, wasps, and sawflies; order Hymenoptera Piercing-sucking mouthparts; tarsi with two to three segments ..................................................................... 7 7. Mouthparts arising at front of head

Whiteflies; order Hemiptera, family Aleyrodidae Front wings not opaque white; veins evident; body size larger ................................................................... 9 9. Front wings appearing transparent; veins few or many; body robust ............................................... 10 Front wings pigmented but veins evident; body relatively long and thin; head often tapering to blunt or acute point Leafhoppers and planthoppers; order Hemiptera,

A thin-bodied insect with pointed head and wing veins: leafhoppers. (Drawing by J. Capinera.)

Piercing-sucking mouthparts arising at front of head. (Drawings by J. Capinera.)

Piercing-sucking mouthparts arising at back of head. (Drawings by J. Capinera.)

families Cicadellidae and Delphacidae 10. Dorsal surface of posterior end of abdomen with short- or long-tubular projections (cornicles) Aphids; order Hemiptera, family Aphididae

Appendix A   629

Abdomen with dorsal projections (cornicles): aphid. (Drawing by J. Capinera.)

Dorsal surface of abdomen lacking apical projections; insect hops when disturbed Psyllids; order Hemiptera, family Psyllidae 11. Tip of abdomen with forceps-like cerci Earwigs; order Dermaptera

Thickened front wings, and legs about equal in size: Coleoptera. (Drawing by J. Capinera.)

14. Antennae short, usually less than half length of body; wings often long Grasshoppers and mole crickets; order Orthoptera, families Acrididae and Gryllotalpidae Antennae long, usually extending beyond tip of abdomen; in vegetable-feeding species, front wings often are abbreviated Crickets; order Orthoptera, families Gryllidae and Tettigoniidae 15. Body constricted at juncture of thorax and abdomen to form narrow “waist,” antennae elbowed Wingless ants; order Hymenoptera

Abdomen constricted and antennae elbowed: ant. (Drawing by J. Capinera.)

Abdomen with forceps-like cerci: earwig. (Drawing by J. Capinera.)

Tip of abdomen lacking cerci or cerci not forceps-like ............................................................... 12 12. Mouthparts adapted for chewing .............................. 13 Mouthparts adapted for piercingsucking ......... go back to ............................................ 7 13. Front wings slightly thickened but bearing branched veins; hind legs appreciably larger than other legs or front legs adapted for digging .............................. 14 Front wings thickened and often with ridges or grooves but lacking branched veins; hind legs about same size as other legs Beetles; order Coleoptera

Body not constricted at abdomen; antennae not elbowed .................................................................... 16 16. Antennae apparent; legs usually normal in length.................................................................... 17 Antennae absent or not apparent; legs usually short or absent .......................................................... 24 17. Abdomen with a spring-like structure (furcula) beneath abdomen Springtails; class Collembola

Spring-like structure beneath abdomen: springtail. (Drawing by J. Capinera.)

630   Appendix A

Abdomen lacking furcula ......................................... 18 18. Body with about 10 segments and at least six pairs of legs; dwelling in soil Symphylans; class Symphyla Body not as described above; if similar, antennae not apparent .............................................................. 19 19. Body flattened; apparently legless; clinging to plant without moving ........................................... 20 Body not flattened; legs present or absent but insect mobile ............................................................ 21 20. Long waxy filaments protruding dorsally Immature whiteflies; order Hemiptera, family Aleyrodidae

Abdomen without cornicles ..................................... 22 22. Body narrow, minute in size, about 1 mm long; short piercing-type mouthparts; tarsi without claws Immature thrips; order Thysanoptera

Flattened body with waxy filaments protruding: immature whitefly. (Drawing by J. Capinera.)

Insect very small with narrow body, and tarsi lacking claws: immature thrips. (Drawing by J. Capinera.)

Short waxy filaments confined to margin of body Immature psyllids; order Hemiptera, family Psyllidae

Flattened body with waxy filaments at margin of body: ­immature psyllid. (Drawing by J. Capinera.)

21. Abdomen with a pair of tubular projections (cornicles) at posterior; body robust; often present in groups along with winged adults Wingless or immature aphids; order Hemiptera, family Aphididae

Body larger in size; mouthparts normally long piercing-sucking type or chewing type; tarsi with claws ................................................................. 23 23. Mouthparts piercing-sucking and located at front of head; antennae with four to five segments; cerci absent Immature stink bugs and plant bugs; order Hemiptera Mouthparts for chewing; antennae with seven or more segments; small cerci present Immature crickets, grasshoppers, and katydids; order Orthoptera 24. Legs present ............................................................. 26 Legs absent .............................................................. 25 25. Body lacking distinct head capsule; mouth hooks present; body usually tapers strongly toward head Immature flies (maggots); order Diptera

Tapered, legless body with small head: immature fly. (Drawing by J. Capinera.)

Abdomen robust and with a pair of dorsal projections: ­immature or wingless adult aphid. (Drawing by J. Capinera.)

Body with distinct head capsule; chewing mouthparts present Immature weevils, primitive weevils, and seed beetles; order Coleoptera, families Curculionidae, Brentidae, and family Chrysomelidae, Bruchinae

Appendix A   631

27. Body with two to five pairs of prolegs Immature moths and butterflies (caterpillars); order Lepidoptera

Body legless but with distinct head: immature weevil or seed beetle. (Drawing by J. Capinera.)

26. Body with fleshy prolegs on abdomen ..................... 27 Body lacking prolegs, except perhaps anal appendages resembling prolegs Immature beetles except Buchinae; order Coleoptera

Body with —two to five pairs of prolegs on abdomen: immature moth. (Drawing by J. Capinera.)

Body with at least six pairs of prolegs Immature sawflies; order Hymenoptera

Body with at least six pairs of prolegs on abdomen; immature sawfly. (Drawing by J. Capinera.)

Body lacking prolegs on abdomen, except for fleshy protuberance on terminal segment: immature beeetle. (Drawing by J. Capinera.)

Note: This simple key was modified from Howard et al. (1994). More complete keys were found in Arnett (1985), Stehr (1987, 1991), and many introductory texts on entomology.

632   Appendix A

KEY TO COMMON STINK BUGS AFFECTING VEGETABLES Note: In this key, “stink bug” is broadly defined to include not only Pentatomidae but also the related Cydnidae and Thyreocoridae. Also, be aware that the color of stink bugs changes after they are killed. Within months, they may change from green to yellowish or brownish, so identifications are best made using fresh specimens. Seasonal changes are also observed in some species. 1. Base of mouthparts (beak) not closely pressed to head, and at least the basal segment of the beak thickened Predatory stink bugs (not plant pests)

Base of mouthparts (beak) not closely pressed to head: beak thick: predatory stink bugs. (Drawing by J. Capinera.)

Base of mouthparts pressed close to head; beak slender throughout Plant-feeding stink bugs ............................................. 2

Scutellum not covering most of abdomen; medium to large bugs measuring >5 mm long ............................. 3 3. Color black or reddish-brown; medium-sized bug measuring 5–8 mm long; tibiae with numerous stout spines Burrowing bug, Pangaeus bilineatus (Say)

Body elongate, wing tips transparent: burrowing bug. (Drawing by J. Capinera.)

Color not uniformly dark; generally larger than 8 mm long; lacking numerous spines on tibiae ...................................................................... 4 4. Color black and orange, or black and yellow; usually small to medium size (12 mm) .............................................. 6 5. Medium-sized bug measuring 8.0–11.5 mm long; backgound color orange, with black spots Harlequin bug, Murgantia histrionica (Hahn)

Base of mouthparts (beak) pressed close to head; beak slender: plant-­feeding stink bugs. (Drawing by J. Capinera.)

2. Scutellum large, covering most of abdomen; small bug measuring 2.2–3.5 mm long, and black in color Little ebony bug, Corymelaena pulicaria (Germar)

A medium-sized, black and red or red and yellow bug: harlequin bug. (Drawing by J. Capinera.)

Scutellum large, covering abdomen: ebony bug. (Drawing by J. Capinera.)

Small bug, 5.0–7.0 mm long; background color black, with orange spots Painted bug, Bagrada hilaris (Burmeister)

Appendix A   633

6. Body color predominantly green or greenish yellow ......................................................................... 7 Body color predominantly brown or grayish-brown .......................................................... 13 7. With a very pronounced spine (usually yellow-tipped) laterally on each side of the pronotum, and with numerous small dark spots on the pronotum and scutellum and white or yellow spots on the hemelytra Tomato stink bug, Arvelius albopunctatus (DeGeer) Lacking the large yellow-tipped spine, and black and yellow/white spots ..................................................... 8 8. Red coloration, varying from faint to dark, crossing the dorsum of the bug at the juncture of the pronotum and scutellum ............................................................. 9 Lacking reddish coloration at juncture of pronotum and scutellum ........................................................... 10 9. Long spine present ventrally between base of legs; smaller, typically 8–11 mm long Red-banded stink bug, Piezodorus guildinii (Westwood) Long spine not present between legs; larger, typically 9–13 mm Red-shouldered stink bugs, Thyanta spp. 10. When viewed from below, second abdominal segment without spine extending forward between the base of the hind legs; body green with a yellow or red spot at the posterior tip of the scutellum and often additional spots along the anterior border of the scutellum; body length 13–15 mm; distribution is limited to Western North America ......................................................... 11

11. Distal region (membrane) of front wing with some purple flecks along vein; tip of scutellum and marginal areas often orange Say stink bug, Chlorochroa sayi (Stål) Distal region (membrane) of front wing without purple flecks along vein; tip of scutellum and marginal areas usually not orange Uhler stink bug, Chlorochroa uhleri (Stål) 12. Spine on second abdominal segment extending forward between base of hind legs with sharp point; scent gland opening located near the base of the middle pair of legs elongate, measuring at least three times as long as wide; body size 13–19 mm Green stink bug, Chinavius hilaris (Say) Spine on second abdominal segment with dull point; scent gland opening located near the base of the middle pair of legs not elongate, measuring about twice as long as wide; body size 14–17 mm Southern green stink bug, Nezara viridula (Linnaeus) 13. Mottled gray or brown body, with small patches of blue or copper; white bands present on antennae and tibiae; abdominal segments protrude from beneath wings, and alternating brown and white in color Brown marmorated stink bug, Halyomorpha halys (Stål) Lacking combination of pronounced mottling, and protruding brown and white abdominal segments ................................................................... 14 14. Sides of pronotum generally acutely pointed ........... 15 Sides of pronotum rounded ...................................... 16 15. Abdominal segments, when viewed from below, with slim black markings at lateral angles Brown stink bug, Euschistus servus (Say)

Ventral surface of stink bugs. Spine absent from between base of hind legs (left); spine with sharp point extending forward between base of hindlegs (center) and with adull point. Scent gland located near base of middle legs elongate (middle) and not elongate (right). (Drawing by J. Capinera.)

When viewed from below, second abdominal segment with small spine extending forward between the base of the hind legs; body color uniformly green; body size viable but often exceeding 15 mm; distribution is principally Eastern North America .......................................................................... 12

Ventral surface of abdomen with slim dark spots at lateral angles of segments. (Drawing by J. Capinera.)

634   Appendix A

Abdominal segments, when viewed from below, without black markings at lateral angles One-spotted stink bug, Euschistus variolaris (Palisot de Beauvois) 16. Dark spots at base of front wing and elsewhere discrete and relatively large, encompassing several small and darker punctures; body size moderate, measuring smaller, 11–12 mm long; distribution limited to west of Rocky Mountains

Consperse stink bug, Euschistus conspersus Uhler Dark spots on front wing and elsewhere not large and discrete, usually limited to punctures; body size averaging larger, 11–15 mm long; widespread in North America Brown stink bug, Euschistus servus (Say)

Appendix A   635

KEY TO COMMON ARMYWORMS AND CUTWORMS Note that the armyworms and cutworms consist of a large and diverse number of caterpillars in the family Noctuidae. They are not easily differentiated, and nearly all keys, including this one, are based on the mature larva. Keys usually require high levels of magnification and examination of setal patterns and mouthparts; by avoiding these difficultto-discern characters, this key sacrifices accuracy for easeof-use. A more detailed key to noctuid larvae was provided by Stehr (1987); other detailed keys included by Crumb (1956) and Godfrey (1972). 1. Caterpillars that might be confused with armyworms and cutworms Caterpillar with a single large spine or “horn” at tip of abdomen Hornworms (Lepidoptera: Sphingidae)

Hornworm. (Drawing by J. Capinera.)

Caterpillar covered with dense covering of long hairs (“woollyworms”) Woollybears and saltmarsh caterpillar (Lepidoptera: Erebidae)

Caterpillar with three to four pairs of abdominal prolegs Loopers (some Lepidoptera: Noctuidae or Erebidae)

Looper. (Drawing by J. Capinera.)

Caterpillar with six or more pairs of abdominal prolegs Sawflies (Hymenoptera: Argidae)

Sawfly. (Drawing by J. Capinera.)

Caterpillars with five pairs of prolegs on the abdomen, having a tendency to feed at or just below the soil surface (so-called subterranean cutworms) or on plant foliage (so-called climbing cutworms), and sometimes having a strong tendency to disperse in aggregations when densities are high (so-called armyworms) Armyworms and cutworms (Lepidoptera: Noctuidae) .............................................................. 2

Saltmarsh caterpillar. (Drawing by J. Capinera.)

Saltmarsh and woollybear caterpillars. Caterpillar producing significant amount of silk, or webbing together leaves with strands of silk Leaftiers and webworms (usually Lepidoptera: Crambidae) Caterpillar burrowing into plant stems and roots Borers (Lepidoptera: some Crambidae, Noctuidae, and Sesiidae) Caterpillar on cabbage and related plants. Often pierid larvae (Lepidoptera: Pieridae), loopers, or diamondback moth.

Armyworm or cutworm. (Drawing by J. Capinera.)

2. Caterpillar with a uniform tan or pink abdomen, lacking stripes or bands .............................................. 3 Caterpillar with a dark abdomen, or with stripes or bands .......................................................................... 6 3. Sutures on top of head bordered by distinct, dark brown bars forming submedial arcs ........................... 4

636   Appendix A

Submedial arcs on top of head. (Drawing by J. Capinera.)

Top of head without submedial arcs .......................... 5 4. Submedial arc in the form of a discrete narrow band; body lacking other dark markings Pale western cutworm, Agrotis orthogonia Morrison

Narrow submedial arc on top of, and in front of, head: pale western cutworm. (Drawing by J. Capinera.)

Submedial arc broad or dark band spreading over most of head; body with dark flecks forming weak stripes laterally and dorsally Granulate cutworm, Agrotis subterranea (Fabricius) 5. Prothoracic shield with irregular central dark bands Western bean cutworm, Striacosta albicosta (Smith)

Caterpillar lacking transverse bands, though longitudinal stripes may be present ............................ 7 7. Abdomen dark but with a row of four or more distinct whitish or yellowish spots mid-dorsally Variegated cutworm, Peridroma saucia (Hübner) Abdomen lacking row of light-colored mid-dorsal spots ........................................................................... 8 8. Abdomen marked dorsolaterally with paired black triangular spots ........................................................... 9 Abdomen lacking paired triangular spots ................. 13 9. Largest triangular black spots located posteriorly on abdomen Spotted cutworms, Xestia c-nigrum Franclemont or X. dolosa Franclemont Triangular black spots on abdomen fairly uniform in size ....................................................................... 10 10. Abdominal segments with a prominent yellowish subspiracular line interrupted by a large dark spot on the first abdominal segment Southern armyworm, Spodoptera eridania (Cramer) Adominal segments without prominent yellowish subspiracular line or, if line is present, line extends without interruption through first abdominal segment ..................................................................... 11 11. Dark markings found dorsolaterally on mesothorax are triangular in shape; triangular marks on abdomen are bisected by a thin white line Yellow-striped armyworm, Spodoptera ornithogalli (Guenée) Mesothoracic markings are semicircular or trapezoidal in shape; abdominal dark marks without white line ..................................................... 12 12. Mesothoracic dark dorsolateral markings semicircular Velvet armyworm, Spodoptera latifascia (Walker) Mesothoracic dark dorsolateral markings trapezoidal Sweetpotato armyworm, Spodoptera dolichos (Fabricius)

Prothoracic shield without central dark bands. (Drawing by J. Capinera.)

Glassy cutworm, Apamea devastator (Brace) 6. Caterpillar with yellow and black transverse bands laterally; head reddish Zebra caterpillar, Melanchra picta (Harris)

Mesothoracic makings of sweetpotato armyworm (left) and velvet armyworm (right). (Drawings by J. Capinera.)

Appendix A   637

13. Head with white bands at margins of sutures, forming an inverted “V” or “Y” when viewed from the front .................................................................... 14 Head lacking white bands along sutures .................. 15 14. White bands form inverted “V” when viewed from front; dorsal surface of abdominal segment eight without dark spots Yellow-striped armyworm, Spodoptera ornithogalli (Guenée) White bands form inverted “Y” when viewed from front; dorsal surface of abdominal segment eight with four dark spots Fall armyworm, Spodoptera frugiperda (J.E. Smith) 15. Abdominal spiracles surrounded by black areas which do not unite to form a continuous black line connecting the spiracles; yellow or pink stripe below the spiracles Clover cutworm, Anarta trifolii (Hufnagel) Abdominal spiracles not surrounded by discontinuous black areas ........................................ 16 16. Caterpillar with brick-red stripes dorsally, separated by a pale stripe Red-backed cutworm, Euxoa ochrogaster (Guenée) Caterpillar lacking brick-red stripes dorsally ..................................................................... 17 17. Epicranium (sides of head) joined narrowly when viewed from above ................................................... 18 Epicranium (sides of head) joined broadly when viewed from above ................................................... 23 18. Dark markings on top of head restricted to submedial arc; prothoracic spiracle round or only slightly oval; abdomen generally uniform tan or gray Pale western cutworm, Agrotis orthogonia Morrison Dark markings on top of head widespread; abdomen usually dark or distinctly marked ............................. 19 19. Abdomen black (mature larva) or gray (immature), sometimes with broad brown stripe dorsally; skin with numerous coarse granules; prothoracic spiracle about twice as long as wide Black cutworm, Agrotis ipsilon (Hufnagel) Abdomen not black, usually indistinctly marked brown gray and tan, and often with a broad light stripe dorsally ........................................................... 20 20. Prothoracic spiracle oval, about 1.5 times as long as wide ..................................................................... 21 Prothoracic spiracle elongate, about twice as long as wide ............................................................. 22 21. Caterpillar mostly gray in color, but with indistinct brown markings; lacking coarse skin texture Dingy cutworm, Feltia jaculifera (Guenée) and F. subgothica (Haworth)

Caterpillar mostly yellowish-brown or sandy, but with indistinct brown markings; skin texture coarse, bearing conical projections Granulate cutworm, Agrotis subterranea (Fabricius) 22. Caterpillar mostly grayish-brown, but with a narrow wavy lateral line Darksided cutworm, Euxoa messoria (Harris) Caterpillar generally yellowish-brown, but with a broad dark band laterally Army cutworm, Euxoa auxiliaris (Grote) 23. Top of head lacking reticulations (net-like markings); prothoracic plate dark centrally; abdomen lacking distinct stripes laterally Western bean cutworm, Striacosta albicosta (Smith) Top of head bearing reticulations ............................. 24 24. Abdominal segment eight bearing four raised brown or black spots dorsally .............................................. 25 Abdominal segment eight lacking four brown or black spots dorsally .............................................. 27 25. Microspines absent from dorsal surface of abdomen; reticulations on head create dark-colored pattern on light background Fall armyworm, Spodoptera frugiperda (J.E. Smith) Minute microspines present on dorsal surface of abdomen, imparting a rough feel; reticulations on head create light-colored pattern on dark background ............................................................... 26 26. Small spines on the tubercles of the first, second, and eighth abdominal segments measure no more than one-fourth the height of the tubercle; a common vegetable pest, particularly on corn Corn earworm, Heliothis zea (Boddie) Small spines on the tubercles of the first, second, and eighth abdominal segments measure about half the height of the tubercle; not commonly a vegetable pest, and never on corn Tobacco budworm, Chloridea virescens (Fabricius)

Small spines on tubercle of tobacco budworm measuring about one-half the height of the tubercle. (Drawing by J. Capinera.)

27. Body color bronze, but with five distinct pale stripes (three when viewed from above), each about onehalf as wide as the bronze area separating them; reticulations on head weak Bronzed cutworm, Nephelodes minians (Guenée)

638   Appendix A

Body color and stripe pattern not as described above; reticulations on head distinct ................................... 28 28. Reticulations on head create light-colored pattern on dark background; body color variable but dark lateral stripe usually present; microspines present on abdomen Corn earworm, Heliothis zea (Boddie) Reticulations on head create dark-colored pattern on light background; lateral stripe variable; microspines not present on abdomen ........................................... 29 29. Abdominal color variable but usually with several distinct alternating dark and light stripes; spiracles dark brown or black throughout; submedial arcs bordering suture on head Armyworm, Mythimna unipuncta (Haworth) Abdominal color variable but usually lacking dark stripes; spiracles white or light brown and bordered by dark ring; suture not bordered by submedial arc;

similar to clover cutworm but lacking dark areas near abdominal spiracles Beet armyworm, Spodoptera exigua (Hübner)

Beet armyworm head capsule lacking complete submedial arcs; the “arc” is limited to the upper portion of the head. (Drawing by J. Capinera.)

Appendix A   639

KEY TO COMMON “CABBAGE WHITE” BUTTERFLIES 1. Upper surface of forewing with at least three to four (in male) or numerous (in female) dark spots centrally (do not consider pigmentation of the wing tip) Southern cabbageworm, Pontia protodice (Boisduval and LeConte) Upper surface of forewing with zero to two black spots centrally ....................................................................... 2 2. Largest of the “cabbage whites”; wingspan about 6–8 cm. May be white, tan, or brown. Dark band along outer margin of forewing, or the band bearing deep indentations of white resulting in a scalloped effect. Central forewing with zero to two spots. Southern white, Ascia monuste (Linnaeus) Forewing not marked with band along entire outer edge, or not deeply indented ........................................ 3 3. A small butterfly (3–5 cm wingspan), usually entirely white; rarely with black spots on, or at the tip of, the forewing. Wing veins, and especially veins of hindwing when viewed from below, marked with dusky stripes Mustard white, Pieris napi (Linnaeus) A butterfly of medium size (4.5–6.5 cm wingspan) with black at tip of forewing, and usually with pronounced black spot(s) centrally. Veins of hindwing not distinctly darkened. Imported cabbageworm, Pieris rapae (Linnaeus)

KEY TO COMMON “CABBAGEWORMS” 1. Larva with black transverse bands crossing body and that affects crucifers in the southeastern states) Cross-striped cabbageworm, Evergestis rimosalis (Guenée) Larva lacking transverse bands; larva either without pattern or with stripes running length of body ............ 2 2. Larvae green and velvety in appearance ...................... 3 Larva gray, bluish, or brownish, and not particularly velvet-like in appearance ............................................ 4 3. Larva with thin yellow line down center of back, and a broken yellow line or series of yellow spots on each side Imported cabbageworm, Pieris rapae (Linnaeus) Larva without yellow line dorsally; stripe, if present, is green. Yellow on sides surrounding spiracles Mustard white, Pieris napi (Linnaeus) 4. Larva with four yellow stripes running length of body Southern cabbageworm, Pontia protodice (Boisduval and LeConte) Larva with five orange or yellow-green stripes running length of body Southern white, Ascia monuste (Linnaeus)

Note: Armyworms and cutworms, loopers, and other caterpillars also affect cabbage.

640   Appendix A

KEY TO COMMON LOOPERS AFFECTING VEGETABLES 1. Larva with four to five pairs of prolegs ...................... 2 Larva with three pairs of prolegs ................................ 4 2. Larva with four pairs of prolegs ................................. 3 Larva with five pairs of prolegs This is not a looper; see armyworms and cutworms 3. Feeding on legumes Green cloverworm, Plathypea scabra (Fabricius) Feeding on okra Okra caterpillar, Anomis erosa Hübner 4. Very small, nipple-like structures (vestigial prolegs) found on abdominal segments three and four............. 5

Abdominal segments 3 and 4 with vestigial prolegs. (Drawing by J. Capinera.)

Nipple-like structures absent from abdominal segments three and four ............................................. 6 5. Thoracic legs usually dark; microspines present on body, but only observable under very high magnification Soybean looper, Chrysodeixis includens (Walker) Thoracic legs not dark; microspines absent Cabbage looper, Trichoplusia ni (Hübner)

6. Body color predominantly brown; longitudinal stripe pattern continuing onto head capsule; large, dark transverse spots usually occur dorsally between the segments at about the midpoint of the body Striped grass looper, Mocis latipes Guenée Body color predominantly green; stripes not continuing onto head capsule; transverse spots absent ................................................................ 7 7. Microspines absent from abdominal segments .......... 8 Microspines present on abdominal segments ............ 9 8. Found in Eastern North America Plantain looper, Autographa precationis (Guenée) Found in Western North America Alfalfa looper, Anagrapha californica (Speyer) 9. Head lacking black lateral bands, though spots may be present ...................................................................... 10 Head with black lateral bands extending through stemmata .................................................................. 11 10. Principally southern in distribution: California and Gulf Coast states; spiracles dark in color Bean leafskeletonizer, Autoplusia egena (Guenée) Principally northern in distribution; spiracles white, but with black rim Celery looper, Anagrapha falcifera (Kirby) 11. Thoracic legs generally black; lateral white line on abdomen is weak; dark bar on head is strong; tubercles above abdominal spiracles often black Bilobed looper; Megalographa biloba (Stephens) Thoracic legs pale; lateral line on abdomen bold; dark bar on head weak; abdominal tubercles not black Celery looper, Anagrapha falcifera (Kirby) Note: This key uses readily observable characters, but lacks the precision of keys based on mouthparts and setal patterns. For greater resolution, consult Crumb (1956), Eichlin and Cunningham (1978), and LaFontaine and Poole (1991).

Appendix A   641

KEY TO COMMON STALK BORERS AFFECTING CORN 1. Larva white or whitish, often with distinct dark spots ............................................................................. 2 Larva light brown or dark brown, with bands, stripes, or indistinct spots ............................................................. 5 2. Larva bearing dark spots dorsally and laterally; summer form of Diatraea spp ................................................... 3 Larva lacking distinct markings; overwintering form of Diatraea spp. 3. Spot adjacent to spiracle on abdominal segments elongate and bean-shaped, partly embracing spiracle Southwestern corn borer, Diatraea grandiosella (Dyar) Spot adjacent to spiracle on abdominal segments more circular, not embracing spiracle ......................... 4 4. Dorsal spots on abdominal segments surrounded by light brown or pink pigmentation Sugarcane borer, Diatraea saccharalis (Fabricius)

Dorsal spots on abdominal segments contrasting distinctly with background color, not surrounded by brown or pink pigmentation Southern cornstalk borer, Diatraea crambidoides (Grote) 5. Larva light brown with small, round, indistinct spots European corn borer, Ostrinia nubilalis (Hübner) Larva dark brown or gray-green with stripes or bands ....................................................................... 6 6. Larva dark brown with white stripe dorsally and broken white stripe laterally Stalk borer, Papaipema nebris (Guenée) Larva greenish or gray with reddish bands (early instars) or white stripes (late instars) Lesser cornstalk borer, Elasmopalpus lignosellus (Zeller) Note: This key is based on “typical” summer forms. Larval color changes as larvae near pupation or prepare for overwintering, and such larvae do not conform to the aforementioned descriptions. See Peterson (1948) and Stehr (1987) for more detailed keys.

642   Appendix A

GUIDE TO COMMON ADULT FLEA BEETLES (Vegetable-feeding Chaetocnema, Disonycha, Epitrix, Phyllotreta, Psylliodes, and Systena) Size moderate to large, usually 3.5 mm long or greater Beetles 5–6 mm long (Disonycha) Spinach flea beetle (preferred hosts: Chenopodiaceae) Three-spotted flea beetle (preferred hosts: Chenopodiaceae) Yellow-necked flea beetle (preferred hosts: Chenopodiaceae) Beetles 3.5–4.5 mm long (Systena) Elongate flea beetle (preferred hosts: variable) Palestriped flea beetle (preferred hosts: variable) Red-headed flea beetle (preferred hosts: variable) Smartweed flea beetle (preferred hosts: variable) Size small, usually 3 mm long or less Elytra covered with short hairs (Epitrix) Elytra color black Eggplant flea beetle (preferred hosts: Solanaceae) Potato flea beetle (preferred hosts: Solanaceae) Tuber flea beetle (preferred hosts: Solanaceae) Elytra color bronze, or with bronze luster Southern tobacco flea beetle (preferred hosts: Solanaceae) Tobacco flea beetle (preferred hosts: Solanaceae) Western potato flea beetle (preferred hosts: Solanaceae)

Elytra lacking layer of hairs Antennae with 10 segments (Psylliodes) Hop flea beetle (preferred hosts: variable) Antennae with 11 segments (Phyllotreta, Chaetocnema) Elytra color black, or blackish with yellow stripes or spots Elytra with yellow stripes or spots Horseradish flea beetle (preferred hosts: Cruciferae) Striped flea beetle (preferred hosts: Cruciferae) Western striped flea beetle (preferred hosts: Cruciferae) Zimmermann’s flea beetle (preferred hosts: Cruciferae) Elytra lacking yellow stripes or spots Desert corn flea beetle (preferred hosts: Gramineae) Elytra color bronze, or with bronze, blue, or green luster Cabbage flea beetle (preferred hosts: Cruciferae) Corn flea beetle (preferred hosts: Gramineae) Crucifer flea beetle (preferred hosts: Cruciferae) Sweetpotato flea beetle (preferred hosts: Convolvulaceae) Toothed flea beetle (preferred hosts: Gramineae) Western black flea beetle (preferred hosts: Cruciferae)

Note: Identification of flea beetles can be achieved by using keys provided by Blatchley (1910), Chittenden (1927), Hatch (1971), and Smith (1970, 1985). An excellent source of information on flea beetle biology, including some minor pests not discussed here, was provided by Campbell et al. (1989).

Appendix A   643

KEY TO GENERA OF WIREWORMS COMMONLY AFFECTING VEGETABLES 1. Antennae conspicuous, with distal segments enlarged; suture on top of head is Y-shaped False wireworm, family Tenebrionidae Antennae not conspicuous, distal segments reduced; suture on top of head not Y-shaped Wireworm, family Elateridae ................................... 2 2. Terminal abdominal segment lacking a notch at the tip ....................................................................... 3 Terminal abdominal segment with small or pronounced notch at tip ................................................................... 4 3. Terminal abdominal segment somewhat flattened and scalloped Melanotus spp. (M. communis, M. longulus oregonensis)

4. Notch at tip of abdomen wide, approximately one-third the width of the tip, and nearly closed; lateral margins and tip of abdomen bearing only dull or inconspicuous protuberances Limonius spp. (C. agonus, L. californicus, L. canus)

Terminal abdominal segments of sugarbeet wireworm. (Drawings by USDA.)

Terminal abdominal segments of Eastern field wireworm. (Drawings by USDA.) Terminal abdominal segment: corn wireworm. (Drawing by USDA.)

Terminal abdominal segment somewhat conical or bulbous, not flattened nor scalloped Agriotis spp. (A. mancus)

Terminal abdominal segments of Pacific Coast wireworm. (Drawings by USDA.)

Terminal abdominal segment: wheat wireworm. (Drawing by USDA.)

Notch at tip of abdomen not nearly closed, or if nearly closed then notch is minute; lateral margins and tip of abdomen often bearing sharply pointed protuberances ............................................................. 5

644   Appendix A

5. Notch at tip of abdomen small or “V”-shaped; found in the Southern United States Conoderus spp. (C. falli, C. vespertinus, C. amplicollis)

Terminal abdominal segment of grairie grain wireworm. (Drawings by USDA.) Terminal abdominal segment, southern potato (left), gulf (center), and tobacco wireworm (right). (Drawings by USDA.)

Notch at tip of abdomen large, not “V”-shaped; found in the Northern United States and Canada Selatosomus and Hadromorphus spp. (S. pruininus. S. aeripennis, S. destructor, H. glaucus).

Terminal abdominal segment of dryland wireworm. (Drawings by USDA.)

Terminal abdominal segment of Great Basin wireworm. (Drawings by USDA.)

Terminal abdominal segment of Puget Sound wireworm. (Drawings by USDA.)

Note: Keys for the identification of over 30 genera of wireworms were produced by Becker and Dogger (1991). Wilkinson (1963) provided an excellent key to distinguish among pest wireworms found in British Columbia and adjacent areas.

Appendix A   645

KEY TO COMMON THRIPS AFFECTING VEGETABLES 1. Body color grayish black, with front wings banded light and dark; mostly affecting beans ....................... 2 Body color variable, front wings uniform or shaded, but not banded ............................................................ 3 2. Dark bands diffuse, found in the Southern United States from California to Florida American bean thrips, Caliothrips phaseoli Hood Dark bands distinct, found in the Southwestern United States and Mexico Bean thrips, Caliothrips fasciatus (Pergande) (Note that thrips with three dark spots on each wing and a light body are likely six-spotted thrips, Scolothrips sexmaculatus (Pergande), a predator of phytophagous thrips.) 3. Front wings shaded near midpoint, but not distinctly banded; small brown spots or blotches located medially on dorsal surface of abdominal segments (tergites); body color yellow Chilli thrips, Scirtothrips dorsalis Hood

Interocellar seta

Ocellus Postocular seta

Anteromarginal seta Anteroangular seta Pronotum Posteroangular seta Posteromarginal seta

Setae (hairs) of anterior portion of body. (Drawing by J. Capinera.)

Anterior angles of pronotum bearing anteroangular hairs that are not discernably larger than hairs found centrally on the pronotum .......................................... 9 5. Pedicel (base) of antennal segment III (counting from base) gradually thickens near the midpoint, but lacks sharp increase in thickness: setal comb on the posterior margin of abdominal segment VIII complete or incomplete ............................................................. 6 (antennal pedicel)

Setae (hairs) of anterior portion of body. (Drawing by J. Capinera.)

Lacking bands or shaded wings; body color variable ....................................................................... 4 4. Anterior angles of pronotum bearing stout hairs (anteroangular setae), which are discernably larger than hairs located centrally on the pronotum ............. 5

Typical antennal segment, showing antennal numbering system. (Drawing by J. Capinera.)

646   Appendix A

7. Pronotum with anteromarginal setae distinctly shorter than anteoangular setae: body color brown, never yellow Tobacco thrips, Frankliniella fusca (Hinds)

Pedicel of third antennal segment of Frankliniella occidentalis (left); F. tritici (center), and F. bispinosa (right). (Drawing by J. Capinera.)

Pedicel (base) of antennal segment III sharply thickens near the midpoint; setal comb on posterior margin of abdominal tergite VIII incomplete (missing central area of comb) ................................... 8 6. Setal comb on posterior margin of abdominal tergite VIII complete (equal length, and equal spacing along margin); some anteromarginal and anteroangular setae on pronotum of similar length; postocular setae as long as, and as stout as, the interocellar setae Western flower thrips, Frankliniella occidentalis (Pergande)

Setae (hairs) of anterior portion of body. (Drawing by J. Capinera.)

Setal comb on posterior marigin of abdominal tergite VIII incomplete (not equal length, shorter centrally); anteromarginal setae usually shorter than anteroangular setae on pronotum; postocular setae shorter and more slender than the interocellar pair; body predominantly brown in color ........................... 7

Setae (hairs) of anterior portion of body: tobacco thrips. (Drawing by J. Capinera.)

Pronotum with anteromarginal setae only slightly shorter than anteroangular setae; body color normally brown in North America, though yellow in South America Common blossom thrips, Frankliniella schultzei (Tybom)

Setae (hairs) of anterior portion of body: common blossom thrips. (Drawing by J. Capinera.)

8. Pedicel of antennal segment III slightly diverging from base and gently curved; pair of dorsal setae on antennal segment III only slightly enlarged, not stout; not a pest of vegetable but very common in blossoms Eastern flower thrips, Frankliniella tritici (Fitch)

Appendix A   647

Setae (hairs) of anterior portion of body: eastern flower thrips. (Drawing by J. Capinera.)

Setae (hairs) of anterior portion of body: onion thrips. (Drawing by J. Capinera.)

Pedicel of antennal segment III strongly diverging and with sharp edges forming ring-like structure; pair of dorsal setae on antennal segment III prominent and stout Florida flower thrips, Frankliniella bispinosa (Morgan)

Hairs on top of head not originating within “triangle” formed by ocelli ....................................................... 10 10. Posterior margin of pronotum with stout hairs; body color yellow; hosts usually cucumbers, eggplant, and tomato; distribution limited to Hawaii, Southern Florida, and the Caribbean region Melon thrips, Thrips palmi Karny

Setae (hairs) of anterior portion of body: Florida flower thrips. (Drawing by J. Capinera.)

9. Hairs on top of head originating within “triangle” formed by ocelli; body color yellowish-brown; found throughout North America and on many crops, but especially on onion and cabbage Onion thrips, Thrips tabaci Lindeman

Setae (hairs) of anterior portion of body: melon thrips. (Drawing by J. Capinera.)

Posterior margin with uniformly small hairs; body color brown or pink; hosts usually corn and other grasses; found widely in North America Grass thrips, Anaphothrips obscurus (Müller) Note: More complete keys to thrips were supplied by Palmer et  al. (1989), Oetting et al. (1993), and Mound and Marullo (1996).

648   Appendix A

KEY TO COMMON MITES AFFECTING VEGETABLES 1. Mite with only two pairs of legs; body elongate and tapering from anterior to posterior; body measuring only about 0.2 mm long; orange-yellow in color; causing bronzing of leaves and deformed tissue growth of solanaceous plants, damage beginning low in the plant and progressing upward Tomato russet mite, Aculops lycopersici (Massee) Mite with four pairs of legs ......................................... 2 2. Mites moving rapidly in seemingly random directions; usually not abundant; not producing silk; often tan or light brown; adult body size 0.4 mm or larger These may be predatory mites in the family Phytoseiidae Mites rather sedentary or moving slowly; often abundant; sometimes producing silk; size variable ..........................................................................3 3. Body whitish and translucent; size variable; damage variable but not expressed as speckling; silk not apparent ....................................................................... 4 Body yellowish, greenish, or reddish, often with dark spots; body size usually 0.4–0.5 mm; feeding injury initially expressed as speckling, followed by bronzing of foliage; silk usually present on foliage .......................................................................... 5 4. Small mites, adult females measuring only 0.2–0.3 mm long; feeding on foliage, primarily of solanaceous crops; foliage usually distorted and plant response variable, but speckling is not a common response to feeding

Broad mite, Polyphagotarsonemus latus (Banks) Large mites, adult females measuring 0.5–0.8 mm or larger in length; feeding below-ground on bulbs and tubers or on injured, decaying tissue; largely a pest of onion and related crops Bulb mites, Rhizoglyphus echinopus (Fumouze and Robin) and Rhizoglyphus robini Claparede 5. During periods of feeding, body color reddish with dark lateral spots Tumid spider mite, Tetranychus tumidus Banks During periods of feeding, body color yellowish or greenish and usually with dark lateral spots ............................................................................. 6 6. Body usually with two dark irregular spots present laterally, with dark spots covering less than one-half the length of the body and not usually extending to the posterior tip of the body Two-spotted spider mite, Tetranychus urticae Koch Body usually with four dark spots present laterally, or spots extending to the posterior tip of the body ............................................................................. 7 7. Feeding on broadleaf plants Strawberry spider mite, Tetranychus turkestani Ugarov and Nikolski Feeding on grasses, particularly corn Banks grass mite, Oligonychus pratensis (Banks)

Note: Mites are difficult to distinguish, even for experts. Final determination should be made by an authority following collection of a large sample of mites to include males of the species.

Appendix A   649

KEY TO ADULT SLUGS COMMONLY AFFECTING VEGETABLES 1. Slug appearing to be hump-backed, bearing swollen area (mantle) immediately behind head ..................... 3 Slug not bearing swelling behind head, mantle extending the length of the slug’s body ..................... 2 2. Body blackish with a yellow stripe dorsally extending the length of the body Laevicaulis alte (Férussac) Body color brownish-gray mottled with black, and with dorsal stripe white and lateral stripes (if present) dark Leidyula floridana (Leidy) 3. Respiratory pore at lateral edge of mantle located anterior to midpoint of mantle; body without dorsal ridge or keel on body; body compact ......................... 4 Respiratory pore at lateral edge of mantle located posterior to midpoint of mantle; body with dorsal ridge or keel along at least a portion of the body; body elongate ............................................................. 6 4. Larger slugs, usually 100–130 mm long; generally uniform black Black slug, Arion ater rufus (Linnaeus) Smaller slugs, usually 40 mm or less in length .......... 5 5. Respiratory pore located below the dark band on the mantle; body length 30–40 mm; body mucus and sole (anterior region of ventral body surface) mucus colorless Banded slug, Arion circumscriptus Johnston Respiratory pore located within the dark band on the mantle; body length 25–30 mm; body mucus yellow to orange, sole mucus colorless Garden slug, Arion hortensis Férussac

6. Dorsal ridge or keel extending from mantle to posterior end .............................................................. 7 Dorsal ridge or keel not evident, or occurring along the posterior end of body only, not reaching mantle, or not apparent ................................................................ 8 7. Dorsal ridge or keel usually dark; body color variable but often yellowish-brown; when viewed from below, periphery is indistinct in color from the central area; length often 45–55 mm but sometimes attaining 70 mm Greenhouse slug, Milax gagates (Draparnaud) Dorsal ridge or keel light, usually yellow or orange; body color variable but usually dark brown, gray, or black; when viewed from below, periphery is distinctly different in color from the central area; length 60–70 mm Subterranean slug, Tandonia budapestensis (Hazay) 8. Larger slugs, measuring 75–150 mm long ................. 9 Smaller slugs, measuring 25–60 mm long ............... 10 9. Grayish or brownish with rows of darker spots or irregular stripes; mucus colorless Spotted slug, Limax maximus Linnaeus Brownish with irregular yellowish spots; mucus yellow Tawny garden slug, Limax flavus Linnaeus 10. Length 20–25 mm; color variable, ranging from gray to brown or brown-black, but with few or no distinct spots or markings; mucus colorless Marsh slug, Deroceras laeve (Müller) Length about 50 mm; color cream to gray or reddishbrown and usually well-marked with spots; mucus milky white Gray garden slug, Deroceras reticulatum (Müller)

650   Appendix A

KEY TO ADULT SNAILS COMMONLY AFFECTING VEGETABLES 1. Snail shell compact, flattened, or oval, not substantially taller than maximum diameter ..................................... 2 Snail shell elongate, at least twice as tall as maximum diameter ....................................................................... 9 2. Small snails, 19 mm or greater ........................... 4 4. Lip at shell opening reddish-brown to almost black, and turned back (reflected); shell 22–23 mm in diameter; color variable, usually yellow, reddish, or brown, sometimes with narrow dark bands; known principally from Northeastern North America and California Brown-lipped snail, Cepaea nemoralis (Linnaeus) Lip at shell opening white; shell 19–22 mm in diameter; color variable, usually ivory, yellow, light brown, or reddish ......................................................................... 5 5. Lip at shell opening turned back (reflected); shell usually yellow with reddish-brown bands; presently restricted to Northeastern North America White-lipped snail, Cepaea hortensis (Müller) Lip at shell opening not turned back; shell color ivory yellow or white with highly variable brown banding pattern; presently restricted to California

White garden snail, Theba pisana (Müller) 6. Shell whitish or gray, dark bands present or absent .... 7 Shell yellowish or brown, distinct dark bands usually absent or indistinct ....................................................... 8 7. Shell 32–45 mm in diameter; color whitish gray, sometimes with light brown bands; distribution presently restricted to Michigan Roman snail, Helix pomatia Linnaeus Shell 28–35 mm in diameter; color entirely white, sometimes with dark brown bands; distribution throughout southern states Milk snail, Otala lactea (Müller) 8. Shell about 20–30 mm in diameter and 20 mm in height; brown bands absent; lip of shell not turned back Singing snail, Helix aperta Born Shell about 28–38 mm in diameter and 35 mm in height; brown bands usually present; lip of shell turned back Brown garden snail, Cornu aspersum Müller 9. Shell elongate, tapering gradually, and with tip absent from adult specimens; color pinkish brown; measuring up to 45 mm in height and 14 mm in diameter Decollate snail, Rumina decollata (Linnaeus) Shell tapering rapidly to a point; color yellowish or grayish with reddish-brown transverse streaks; measuring up to 125 mm long and 60 mm in diameter Giant African snail, Lissachatina fulica Bowdich

Appendix B

Vegetable Plant Names Note that most of the vegetable crop family names have been revised in the recent years. The updated names shown here are, but in the “Introduction,” the section on

“Characteristics of the major vegetable crops” contains both the traditional and revised family names.

COMMON NAME, SCIENTIFIC NAME, AND PLANT FAMILY Common name

Scientific name

Plant family

Arugula

Eruca sativa

Brassicaceae

Artichoke, globe

Cynara scolymus

Asteraceae

Artichoke, Jerusalem

Helianthus tuberosus

Asteraceae

Asparagus

Asparagus officinalis

Asparagaceae

Bean, broad

Vicia faba

Fabaceae

Bean, lima

Phaseolus limensis

Fabaceae

Bean, mung

Phaseolus aureus

Fabaceae

Bean, snap

Phaseolus vulgaris

Fabaceae

Beet

Beta vulgaris

Amaranthaceae

Bok choy

Brassica campestris, var. pekinensis

Brassicaceae

Broccoli

Brassica oleracea, var. italica

Brassicaceae

Broccoli raab

Brassica campestris, var. ruvo

Brassicaceae

Brussels sprout

Brassica oleracea, var. gemmifera

Brassicaceae

Cabbage

Brassica oleracea, var. capitata

Brassicaceae

Cabbage, bok choy

Brassica campestris, var. pekinensis

Brassicaceae

Cabbage, Chinese

Brassica campestris, var. chinensis

Brassicaceae

Calabaza

Cucurbita moscata

Cucurbitaceae

Cantaloupe

Cucumis melo

Cucurbitaceae

Cardoon

Cynara cardunculus

Asteraceae

Carrot

Daucus carota

Apiaceae

Cauliflower

Brassica oleracea, var. botrytis

Brassicaceae

Celeriac

Apium graveolens, var. rapaceum

Apiaceae Continued

651

652   Appendix B

Common name

Scientific name

Plant family

Celery

Apium graveolens, var. dulce

Apiaceae

Celtuce

Lactuca sativa, var. asparagina

Asteraceae

Chard

Beta vulgaris, var. cicla

Amaranthaceae

Chayote

Sechium edule

Cucurbitaceae

Chervil, salad

Anthriscus cerefolium

Apiaceae

Chervil, turnip-rooted

Chaerophyllum bulbosum

Apiaceae

Chickpea

Cicer arietinum

Fabaceae

Chicory

Chicorium intybus

Asteraceae

Chili

Capsicum annuum

Solanaceae

Chive

Allium schoenoprasum

Amarylidaceae

Collard

Brassica oleracea, var. acephala

Brassicaceae

Coriander

Coriandrum sativum

Apiaceae

Corn, sweet

Zea mays

Poaceae

Cowpea

Vigna sinensis

Fabaceae

Cucumber

Cucumis sativus

Cucurbitaceae

Daikon

Raphanus sativus

Brassicaceae

Eggplant

Solanum melongena

Solanaceae

Endive

Cichorium endivia

Asteraceae

Escarole

Cichorium endivia

Asteraceae

Faba bean

Vicia faba

Fabaceae

Fennel

Foeniculum vulgare

Apiaceae

Garbanzo

Cicer arietinum

Fabaceae

Garlic

Allium sativum

Amarylidaceae

Garlic, elephant

Allium ampeloprasum

Amarylidaceae

Gherkin

Cucumis anguria

Cucurbitaceae

Globe artichoke

Cynara scolymus

Asteraceae

Horseradish

Amoracia lapathifolia

Brassicaceae

Husk tomato

Pysalis pruinosa

Solanaceae

Jerusalem artichoke

Helianthus tuberosus

Asteraceae

Kale

Brassica oleracea, var. cephala

Brassicaceae

Kale, sea

Crambe maritima

Brassicaceae

Kohlrabi

Brassica oleracea, var. gongylodes

Brassicaceae

Leek

Allium ampeolprasum, var. leek

Amarylidaceae

Lentil

Lepidium sativum

Brassicaceae

Lettuce, head

Lactuca sativa, var. capitata

Asteraceae

Lettuce, leaf

Lactuca sativa, var. crispa

Asteraceae

Lettuce, romaine

Lactuca sativa, var. longifolia

Asteraceae

Mushroom

Agaricus sp.

Agaricaceae Continued

Appendix B   653

Common name

Scientific name

Plant family

Muskmelon

Cucumis melo

Cucurbitaceae

Mustard

Brassica jundea, var. crispifolia

Brassicaceae

Okra

Hibuscus esculentus

Malvaceae

Onion

Allium cepa

Amarylidaceae

Parsley

Petroselinum crispum

Apiaceae

Parsnip

Pastinaca sativa

Apiaceae

Pea, edible-podded

Pisum sativum

Fabaceae

Pea, garden

Pisum sativum

Fabaceae

Pea, pigeon

Cajanus cajan

Fabaceae

Pea, southern

Vigna sinensis

Fabaceae

Pepper, bell

Capsicum annuum

Solanaceae

Pepper, chili

Capsicum annuum

Solanaceae

Potato

Solanum tuberosum

Solanaceae

Pumpkin

Cucurbita spp.

Cucurbitaceae

Purslane

Portulaca oleracea

Portulacaceae

Radicchio

Cichorium intybus

Asteraceae

Radish

Raphanus sativus

Brassicaceae

Radish, Chinese

Raphanus sativus

Brassicaceae

Rhubarb

Rheum rhabarbarum

Polygonaceae

Rutabaga

Brassica napus, var. napobrassica

Brassicaceae

Salsify

Tragopogon porrifolius

Asteraceae

Sea kale

Crambe maritima

Brassicaceae

Shallot

Allium cepa, var. aggregatum

Amarylidaceae

Southern pea

Vigna sinensis

Fabaceae

Spinach

Spinacia oleracea

Amaranthaceae

Spinach, New Zealand

Tetragonia tetragonioides

Aizoceae

Squash, summer

Cucurbita pepo

Cucurbitaceae

Squash, winter

Cucurbita maxima

Cucurbitaceae

Squash, winter

Cucurbita moschata

Cucurbitaceae

Sweet potato

Ipomoea batatas

Convolvulaceae

Swiss chard

Beta vulgaris, var. cicla

Amaranthaceae

Tomatillo

Physalis ixocarpa

Solanaceae

Tomato

Lycopersicon esculentum

Solanaceae

Tomato, husk

Physalis pruinosa

Solanaceae

Turnip

Brassica rapa, var. rapifera

Brassicaceae

Watercress

Nasturtium officinale

Brassicaceae

Watermelon

Citrullus lanatus

Cucurbitaceae

Yam

Dioscorea spp.

Dioscoreaceae

654   Appendix B

SCIENTIFIC NAME, COMMON NAME, AND PLANT FAMILY Scientific name

Common name

Plant family

Agaricus sp.

Mushroom

Agaricaceae

Allium ampeloprasum

Garlic, elephant

Amarylidaceae

Allium ampeolprasum, var. leek

Leek

Amarylidaceae

Allium cepa

Onion

Amarylidaceae

Allium cepa, var. aggregatum

Shallot

Amarylidaceae

Allium sativum

Garlic

Amarylidaceae

Allium schoenoprasum

Chive

Amarylidaceae

Amoracia lapathifolia

Horseradish

Brassicaceae

Anthriscus cerefolium

Chervil, salad

Apiaceae

Apium graveolens, var. dulce

Celery

Apiaceae

Apium graveolens, var. rapaceum

Celeriac

Apiaceae

Asparagus officinalis

Asparagus

Asparagaceae

Beta vulgaris

Beet

Amaranthaceae

Beta vulgaris, var. cicla

Chard

Amaranthaceae

Beta vulgaris, var. cicla

Swiss chard

Amaranthaceae

Brassica campestris, var. chinensis

Chinese cabbage, bok choy

Brassicaceae

Brassica campestris, var. pekinensis

Bok choy, nappa

Brassicaceae

Brassica rapa

Turnip greens, mustard

Brassicaceae

Brassica rapa, subsp. pekinensis

Napa cabbage

Brassicaceae

Brassica campestris, var. ruvo

Broccoli raab

Brassicaceae

Brassica jundea, var. crispifolia

Mustard

Brassicaceae

Brassica napus, var. napobrassica

Rutabaga

Brassicaceae

Brassica oleracea, var. acephala

Collard

Brassicaceae

Brassica oleracea, var. botrytis

Cauliflower

Brassicaceae

Brassica oleracea, var. capitata

Cabbage

Brassicaceae

Brassica oleracea, var. cephala

Kale

Brassicaceae

Brassica oleracea, var. gemmifera

Brussels sprout

Brassicaceae

Brassica oleracea, var. gongylodes

Kohlrabi

Brassicaceae

Brassica oleracea, var. italica

Broccoli

Brassicaceae

Brassica rapa, var. rapifera

Turnip

Brassicaceae

Cajanus cajan

Pea, pigeon

Fabaceae

Capsicum annuum

Chili

Solanaceae

Capsicum annuum

Pepper, bell

Solanaceae

Capsicum annuum

Pepper, chili

Solanaceae

Chaerophyllum bulbosum

Chervil, turnip-rooted

Apiaceae

Chicorium intybus

Chicory

Asteraceae Continued

Appendix B   655

Scientific name

Common name

Plant family

Cicer arietinum

Chickpea

Fabaceae

Cicer arietinum

Garbanzo

Fabaceae

Cichorium endivia

Endive

Asteraceae

Cichorium endivia

Escarole

Asteraceae

Cichorium intybus

Radicchio

Asteraceae

Citrullus lanatus

Watermelon

Cucurbitaceae

Coriandrum sativum

Coriander

Apiaceae

Crambe maritima

Kale, sea

Brassicaceae

Crambe maritima

Sea kale

Brassicaceae

Cucumis anguria

Gherkin

Cucurbitaceae

Cucumis melo

Cantaloupe

Cucurbitaceae

Cucumis melo

Muskmelon

Cucurbitaceae

Cucumis sativus

Cucumber

Cucurbitaceae

Cucurbita maxima

Squash, winter

Cucurbitaceae

Cucurbita moscata

Calabaza

Cucurbitaceae

Cucurbita moschata

Squash, winter

Cucurbitaceae

Cucurbita pepo

Squash, summer

Cucurbitaceae

Cucurbita spp.

Pumpkin

Cucurbitaceae

Cynara cardunculus

Cardoon

Asteraceae

Cynara scolymus

Artichoke, globe

Asteraceae

Cynara scolymus

Globe artichoke

Asteraceae

Daucus carota

Carrot

Apiaceae

Dioscorea spp.

Yam

Dioscoreaceae

Eruca sativa

Arugula

Brassicaceae

Foeniculum vulgare

Fennel

Apiaceae

Helianthus tuberosus

Artichoke, Jerusalem

Asteraceae

Helianthus tuberosus

Jerusalem artichoke

Asteraceae

Hibuscus esculentus

Okra

Malvaceae

Ipomoea batatas

Sweet potato

Convolvulaceae

Lactuca sativa, var. asparagina

Celtuce

Asteraceae

Lactuca sativa, var. capitata

Lettuce, head

Asteraceae

Lactuca sativa, var. crispa

Lettuce, leaf

Asteraceae

Lactuca sativa, var. longifolia

Lettuce, romaine

Asteraceae

Lepidium sativum

Lentil

Brassicaceae

Lycopersicon esculentum

Tomato

Solanaceae

Nasturtium officinale

Watercress

Brassicaceae

Pastinaca sativa

Parsnip

Apiaceae

Petroselinum crispum

Parsley

Apiaceae Continued

656   Appendix B

Scientific name

Common name

Plant family

Phaseolus aureus

Bean, mung

Fabaceae

Phaseolus limensis

Bean, lima

Fabaceae

Phaseolus vulgaris

Bean, snap

Fabaceae

Physalis ixocarpa

Tomatillo

Solanaceae

Physalis pruinosa

Tomato, husk

Solanaceae

Pisum sativum

Pea, edible-podded

Fabaceae

Pisum sativum

Pea, garden

Fabaceae

Portulaca oleracea

Purslane

Portulacaceae

Pysalis pruinosa

Husk tomato

Solanaceae

Raphanus sativus

Daikon

Brassicaceae

Raphanus sativus

Radish

Brassicaceae

Raphanus sativus

Radish, Chinese

Brassicaceae

Rheum rhabarbarum

Rhubarb

Polygonaceae

Sechium edule

Chayote

Cucurbitaceae

Solanum melongena

Eggplant

Solanaceae

Solanum tuberosum

Potato

Solanaceae

Spinacia oleracea

Spinach

Amaranthaceae

Tetragonia tetragonioides

Spinach, New Zealand

Aizoceae

Tragopogon porrifolius

Salsify

Asteraceae

Vicia faba

Bean, broad

Fabaceae

Vicia faba

Faba bean

Fabaceae

Vigna sinensis

Cowpea

Fabaceae

Vigna sinensis

Pea, southern

Fabaceae

Vigna sinensis

Southern pea

Fabaceae

Zea mays

Corn, sweet

Poaceae

PLANT FAMILY, COMMON NAME, AND SCIENTIFIC NAME Plant family

Common name

Scientific name

Agaricaceae

Mushroom

Agaricus sp.

Aizoceae

Spinach, New Zealand

Tetragonia tetragonioides

Amaranthaceae

Beet

Beta vulgaris

Amaranthaceae

Chard

Beta vulgaris, var. cicla

Amaranthaceae

Spinach

Spinacia oleracea

Amaranthaceae

Swiss chard

Beta vulgaris, var. cicla

Amarylidaceae

Chive

Allium schoenoprasum

Amarylidaceae

Garlic

Allium sativum Continued

Appendix B   657

Plant family

Common name

Scientific name

Amarylidaceae

Garlic, elephant

Allium ampeloprasum

Amarylidaceae

Leek

Allium ampeolprasum, var. leek

Amarylidaceae

Onion

Allium cepa

Amarylidaceae

Shallot

Allium cepa, var. aggregatum

Apiaceae

Carrot

Daucus carota

Apiaceae

Celeriac

Apium graveolens, var. rapaceum

Apiaceae

Celery

Apium graveolens, var. dulce

Apiaceae

Chervil, salad

Anthriscus cerefolium

Apiaceae

Chervil, turnip-rooted

Chaerophyllum bulbosum

Apiaceae

Coriander

Coriandrum sativum

Apiaceae

Fennel

Foeniculum vulgare

Apiaceae

Parsley

Petroselinum crispum

Apiaceae

Parsnip

Pastinaca sativa

Asparagaceae

Asparagus

Asparagus officinalis

Asteraceae

Artichoke, globe

Cynara scolymus

Asteraceae

Artichoke, Jerusalem

Helianthus tuberosus

Asteraceae

Cardoon

Cynara cardunculus

Asteraceae

Celtuce

Lactuca sativa, var. asparagina

Asteraceae

Chicory

Chicorium intybus

Asteraceae

Endive

Cichorium endivia

Asteraceae

Escarole

Cichorium endivia

Asteraceae

Globe artichoke

Cynara scolymus

Asteraceae

Jerusalem artichoke

Helianthus tuberosus

Asteraceae

Lettuce, head

Lactuca sativa, var. capitata

Asteraceae

Lettuce, leaf

Lactuca sativa, var. crispa

Asteraceae

Lettuce, romaine

Lactuca sativa, var. longifolia

Asteraceae

Radicchio

Cichorium intybus

Asteraceae

Salsify

Tragopogon porrifolius

Brassicaceae

Arugula

Eruca sativa

Brassicaceae

Bok choy

Brassica campestris, var. pekinensis

Brassicaceae

Broccoli

Brassica oleracea, var. italica

Brassicaceae

Broccoli raab

Brassica campestris, var. ruvo

Brassicaceae

Brussels sprout

Brassica oleracea, var. gemmifera

Brassicaceae

Cabbage

Brassica oleracea, var. capitata

Brassicaceae

Cabbage, bok choy

Brassica campestris, var. pekinensis

Convolvulaceae

Sweet potato

Ipomoea batatas

Cucurbitaceae

Cucumber

Cucumis sativus

Cucurbitaceae

Gherkin

Cucumis anguria Continued

658   Appendix B

Plant family

Common name

Scientific name

Cucurbitaceae

Muskmelon

Cucumis melo

Cucurbitaceae

Pumpkin

Cucurbita spp.

Cucurbitaceae

Squash, summer

Cucurbita pepo

Cucurbitaceae

Squash, winter

Cucurbita maxima

Cucurbitaceae

Squash, winter

Cucurbita moschata

Cucurbitaceae

Watermelon

Citrullus lanatus

Dioscoreaceae

Yam

Dioscorea spp.

Fabaceae

Bean, broad

Vicia faba

Fabaceae

Bean, lima

Phaseolus limensis

Fabaceae

Bean, mung

Phaseolus aureus

Fabaceae

Bean, snap

Phaseolus vulgaris

Fabaceae

Chickpea

Cicer arietinum

Fabaceae

Cowpea

Vigna sinensis

Fabaceae

Faba bean

Vicia faba

Fabaceae

Garbanzo

Cicer arietinum

Fabaceae

Pea, edible-podded

Pisum sativum

Fabaceae

Pea, garden

Pisum sativum

Fabaceae

Pea, pigeon

Cajanus cajan

Fabaceae

Pea, southern

Vigna sinensis

Fabaceae

Southern pea

Vigna sinensis

Malvaceae

Okra

Hibuscus esculentus

Poaceae

Corn, sweet

Zea mays

Polygonaceae

Rhubarb

Rheum rhabarbarum

Portulacaceae

Purslane

Portulaca oleracea

Solanaceae

Chili

Capsicum annuum

Solanaceae

Eggplant

Solanum melongena

Solanaceae

Husk tomato

Pysalis pruinosa

Solanaceae

Pepper, bell

Capsicum annuum

Solanaceae

Pepper, chili

Capsicum annuum

Solanaceae

Potato

Solanum tuberosum

Solanaceae

Tomatillo

Physalis ixocarpa

Solanaceae

Tomato

Lycopersicon esculentum

Solanaceae

Tomato, husk

Physalis pruinosa

Journal Abbreviations and Journal Titles Acarologia Acta Agro. Acta Agric. Scandinavica Acta Entomol. Sinica Acta Hort. Acta Protozool. African J. Agric. Res. African J. Biotech. Agric. Canada Tech. Bull. Agric. Ecosyst. Environ. Agric. Forest Entomol. Agric. Zool. Rev. Am. Entomol. Am. J. Agric. Biol. Sci. Am. J. Pot. Res. Am. Midl. Nat. Am. Nat. Am. Pot. J. An. Soc. Entomol. Brasil Ann. Agric. Fenn. Ann. Appl. Biol. Ann. Entomol. Fennici Ann. Entomol. Soc. Am. Ann. Entomol. Soc. Brasil Ann. Entomol. Soc. Quebec Ann. Hist. Nat. Mus. Natl. Hung. Annu. Mtg. Florida State Hort. Soc. Annu. Rev. Entomol. Apidologie Appl. Agric. Res. Appl. Entomol. Zool. Appl. Environ. Microbiol. Appl. Soil Ecol. Arch. Phytopath. Plant Prot. Arthro. Manage. Tests Arthropod-Plant Inter. Aust. J. Agric. Res. Aust. J. Zool. Behav. Ecol. Sociobiol.

Acarologia Acta Agrobotanica Acta Agriculturae Scandinavica Acta Entomologica Sinica Acta Horticulturae Acta Protozoologica African Journal of Agricultural Research African Journal of Biotechnology Agriculture Canada Technical Bulletin Agriculture, Ecosystems and Environment Agricultural and Forest Entomology Agricultural Zoology Reviews American Entomologist American Journal of Agricultural and Biological Sciences American Journal of Potato Research American Midland Naturalist American Naturalist American Potato Journal Anais do Sociedade Entomologica do Brasil Annales Agriculture Fenniae Annals of Applied Biology Annales Entomologici Fennici Annals of the Entomological Society of America Annais do Sociedade Entomologica do Brasil Annals of the Entomological Society of Quebec Annales Musei Historico-Naturalis Hungarici Annual Meeting of the Florida State Horticultural Society Annual Review of Entomology Apidologie Applied Agricultural Research Applied Entomology and Zoology Applied and Environmental Microbiology Applied Soil Ecology Archives of Phytopathology and Plant Protection Arthropod Management Tests Arthropod-Plant Interactions Australian Journal of Agricultural Research Australian Journal of Zoology Behavioral Ecology and Sociobiology

659

660   Journal Abbreviations and Journal Titles

BioControl Biocontrol News Infor. Biocontrol Sci. Tech. Biol. Agric. Hort. Biol. Control Bull. Brooklyn Entomol. Soc. Bull. Entomol. Res. Bull. Entomol. Soc. Am. Bull. So. California Acad. Sci. Bull. Soc. Entomol. Egypte California Agric. Can. Entomol. Can. J. Bot. Can. J. Plant Sci. Can. J. Res. Can. J. Zool. Clin. All. Coleop. Bull. Crop Prot. Crop Sci. Ecol. Entomol. Ecology Entomol. Am. Entomol. Exp. Appl. Entomol. Mon. Mag. Entomol. News Entomophaga Environ. Entomol. Evolution Exp. Appl. Acarol. Experientia FAO Plant Prot. Bull. Fieldiana Florida Entomol. Fund. Appl. Nematol. Great Lakes Entomol. Hilgardia HortScience Indian J. Agric. Sci. Indian J. Entomol. Indust. Crops Prod. Insect Biochem. Molec. Biol. Insect Life Insect Sci. Applic. Insecta Matsu. Insecta Mun. Insects Integr. Pest Manage. Rev. Inter. J. Agric. Biol.

BioControl Biocontrol News and Information Biocontrol Science and Technology Biological Agriculture and Horticulture Biological Control Bulletin of the Brooklyn Entomological Society Bulletin of Entomological Research Bulletin of the Entomological Society of America Bulletin Southern California Academy of Sciences Bulletin of the Entomological Society of Egypt California Agriculture Canadian Entomologist Canadian Journal of Botany Canadian Journal of Plant Science Canadian Journal of Research Canadian Journal of Zoology Clinical Allergy The Coleopterists Bulletin Crop Protection Crop Science Ecological Entomology Ecology Entomologica Americana Entomologia Experimentalis et Applicata Entomologist’s Monthly Magazine Entomological News Entomophaga Environmental Entomology Evolution Experimental and Applied Acarology Experientia Food and Agriculture Organization of the United Nations, Plant Protection Bulletin Fieldiana Florida Entomologist Fundamentals of Applied Nematology Great Lakes Entomologist Hilgardia HortScience Indian Journal of Agricultural Science Indian Journal of Entomology Industrial Crops and Products Insect Biochemistry and Molecular Biology Insect Life Insect Science and its Application Insecta Matsumurana Insecta Mundi Insects Integrated Pest Management Reviews International Journal of Agriculture and Biology

Journal Abbreviations and Journal Titles   661

Inter. J. Curr. Micro. Appl. Sci. Inter. J. Pest Manage. Inter. J. Trop. Insect Sci. Inter. J. Veg. Sci. IOBC, WPRS Bull. IOBC/WPRS Bull. IPM Pract. Irrigation Sci. J. Acadian Entomol. Soc. J. Agric. Entomol. J. Agric. Food Chem. J. Agric. Res. J. Agric. Sci. J. Agric. Univ. Puerto Rico J. Agric. Urban Entomol. J. Am Soc. Sugar Cane Technol. J. Am. Soc. Hort. Sci. J. Anim. Plt. Sci. J. Animal Ecol. J. Appl. Biol. J. Appl. Ecol. J. Appl. Entomol. J. Asia Pacific Entomol. J. Aust. Entomol. Soc. J. Biol. Control J. Biol. Sci. Res. J. Biopest. J. Chem. Ecol. J. Econ. Entomol. J. Entomol. J. Entomol. Acarol. Res. J. Entomol. Sci. J. Entomol. Soc. Brit. Columbia J. Entomol. Soc. Iran J. Environ. Entomol. J. Essential Oil-Bearing Plt. J. Ethol. J. Eucary. Micro. J. Evol. Biol. J. Exp. Bot. J. Exp. Zool. J. Georgia Entomol. Soc. J. Hymen. Res. J. Insect Behav. J. Insect Pathol. J. Insect Physiol.

International Journal of Current Microbiology and Applied Sciences International Journal of Pest Management International Journal of Tropical Insect Science International Journal of Vegetable Science International Organization of Biological Control, Western Palearctic Regional Section Bulletin IOBC/WPRS Bulletin The IPM Practitioner Irrigation Science Journal of the Acadian Entomological Society Journal of Agricultural Entomology Journal of Agricultural and Food Chemistry Journal of Agricultural Research Journal of Agricultural Science Journal of Agriculture of the University of Puerto Rico Journal of Agricultural and Urban Entomology Journal of the American Society of Sugar Cane Technologists Journal of the American Society for Horticultural Science Journal of Animal and Plant Science Journal of Animal Ecology Journal of Applied Biology Journal of Applied Ecology Journal of Applied Entomology Journal of Asia-Pacific Entomology Journal of the Australian Entomological Society Journal of Biological Control Journal of Biological Science Research Journal of Biopesticides Journal of Chemical Ecology Journal of Economic Entomology Journal of Entomology Journal of Entomological and Acarological Research Journal of Entomological Science Journal of the Entomological Society of British Columbia Journal of Entomological Society of Iran Journal of Environmental Entomology Journal of Essential Oil-Bearing Plants Journal of Ethology Journal of Eucaryotic Microbiology Journal of Evolutionary Biology Journal of Experimental Botany Journal of Experimental Zoology Journal of the Georgia Entomological Society Journal of Hymenoptera Research Journal of Insect Behavior Journal of Insect Pathology Journal of Insect Physiology

662   Journal Abbreviations and Journal Titles

J. Insect Sci. J. Integr. Agric. J. Integr. Pest Manage. J. Invertebr. Pathol. J. Kansas Entomol. Soc. J. Lepid. Soc. J. Nat. Hist. J. Nat. Prod. J. Nematol. J. New York Entomol. Soc. J. Parasit. J. Pest Sci. J. Prod. Agric. J. Range Manage. J. Res. Lep. J. Rio Grande Valley Hort. Soc. J. Root Crops J. Stored Prod. Res. J. Sugar Beet Res. J. Trinidad Tobago Field Nat. J. Zool. Japanese Agric. Res. Quar. Japanese J. Appl. Entomol. Zool. Japanese J. Entomol. Korean J. Appl. Entomol. Manitoba Entomol. Maydica Mem. Am. Entomol. Soc. Mem. Entomol. Inter. Mem. Entomol. Soc. Canada Microb. Ecol. Midsouth Entomol. Molec. Ecol. Molec. Phylo. Evol. Nat. Biotech. Nature Naturwissenschaften Nematology Neotrop. Entomol. Netherlands J. Zool. New Phytol. New Zealand Entomol. New Zealand J. Crop Hort. Sci. New Zealand Plt. Prot. Oecologia Ohio J. Sci. Oikos Outl. Agric. Pacific Sci. Pakistan J. Biol. Sci.

Journal of Insect Science Journal of Integrative Agriculture Journal of Integrated Pest Management Journal of Invertebrate Pathology Journal of the Kansas Entomological Society Journal of the Lepidopterists’ Society Journal of Natural History Journal of Natural Products Journal of Nematology Journal of the New York Entomological Society Journal of Parasitology Journal of Pest Science Journal of Production Agriculture Journal of Range Management Journal of Research on the Lepidoptera Journal of the Rio Grande Horticultural Society Journal of Root Crops Journal of Stored Product Research Journal of Sugar Beet Research Living World: Journal of the Trinidad and Tobago Field Naturalist’s Club Journal of Zoology (London) Japanese Agricultural Research Quarterly Japanese Journal of Applied Entomology and Zoology Japanese Journal of Entomology Korean Journal of Applied Entomology Manitoba Entomologist Maydica Memoirs of the American Entomological Society Memoirs on Entomology, International Memoirs of the Entomological Society of America Microbial Ecology Midsouth Entomologist Molecular Ecology Molecular Phylogenetics and Evolution Nature Biotechnology Nature Naturwissenschaften Nematology Neotropical Entomology Netherlands Journal of Zoology New Phytologist New Zealand Entomologist New Zealand Journal of Crop and Horticultural Science New Zealand Plant Protection Oecologia Ohio Journal of Science Oikos Outlook on Agriculture Pacific Science Pakistan Journal of Biological Sciences

Journal Abbreviations and Journal Titles   663

Pakistan J. Zool. Pan-Pac. Entomol. PANS Pedobiologia Persian Gulf Crop Prot. Pest Manage. Hort. Ecosys. Pest Manage. Sci. Pest. Sci. Physiol. Entomol. Phytoparasitica Phytopathology Phytoprotection Plant Dis. Plant Dis. Rep. Plant Pathol. PLOS ONE Plt. Breed. Pop. Ecol. Potato Res. Prairie Soils Crops J. Proc. Entomol. Soc. Brit. Columbia Proc. Entomol. Soc. Ontario Proc. Entomol. Soc. Philadelphia Proc. Entomol. Soc. Washington Proc. Florida State Hort. Soc. Proc. Hawaiian Entomol. Soc. Proc. Iowa Acad. Sci. Proc. Nat. Acad. Sci. USA Proc. R. Entomol. Soc. Lond. Proc. Third Inter. Conf. Urban Pests Proc. U.S. Nat. Mus. Prot. Ecol. Psyche Quaes. Entomol. Quebec Soc. Prot. Plants Rev. Entomol. Soc. Argentina Revista Bras. Entomol. Revista Bras. Zool. Rivista Instit. Colombiano Agropec. Sarhad J. Aric. Sci. Agr. Sci. Agric. Sociobiology Soil Tillage Res. Southeastern Nat. Southwestern Entomol. Southwestern Nat.

Pakistan Journal of Zoology Pan-Pacific Entomologist Pest Articles and News Summaries Pedobiologia Persian Gulf Crop Protection Pest Management in Horticultural Ecosystems Pest Management Science Pesticide Science Physiological Entomology Phytoparasitica Phytopathology Phytoprotection Plant Disease Plant Disease Reporter Plant Pathology PLOS ONE Plant Breeding Population Ecology Potato Research Prairie Soils and Crops Journal Proceedings of the Entomological Society of British Columbia Proceedings of the Entomological Society of Ontario Proceedings of the Entomological Society of Philadelphia Proceedings of the Entomological Society of Washington Proceedings of the Florida State Horticultural Society Proceedings of the Hawaiian Entomological Society Proceedings of the Iowa Academy of Science Proceedings of the National Academy of Science USA Proceedings of the Royal Entomological Society of London Proceedings of the Third International Conference on Urban Pests Proceedings of the United States National Museum Protection Ecology Psyche Quaestiones entomologicae Quebec Society for the Protection of Plants, Report Revista de la Sociedad Entomológica Argentina Revista Brasileira de Entomologia Revista Brasileira de Zoologia Rivista Instituto Colombiano Agropecuario Sarhad Journal of Agriculture Scientia Agricola Scientific Agriculture Sociobiology Soil and Tillage Research Southeastern Naturalist Southwestern Entomologist Southwestern Naturalist

664   Journal Abbreviations and Journal Titles

Symbiosis Syst. Appl. Acarl. Spec. Pub. Syst. Entomol. Theor. Appl. Genet. Trans. R. Entomol. Soc. Trends Agric. Sci. Trop. Agric. Trop. Pest Manage. Trop. Sci. Univ. California Pub. Entomol. Univ. Sci. Veg. Crops Res. Bull. Z. Angew. Entomol. ZooKeys Zool. Studies Zoologia

Symbiosis Systematic and Applied Acarology Special Publications Systematic Entomology Theoretical and Applied Genetics Transactions of the Royal Entomological Society Trends in Agricultural Science Tropical Agriculture Tropical Pest Management Tropical Science University of California Publications in Entomology Universitas Scientiarum Vegetable Crops Research Bulletin Zeitschrift fur Angewandte Entomologie ZooKeys Zoological Studies Zoologia

Glossary

Abaxial  Facing toward the stem of a plant. Abdomen  The posterior of the three main body divisions of an insect. Abiotic disease  A disease caused by factors other than pathogens (e.g., weather or chemicals). Acaricide  A pesticide applied to manage mite populations, also called miticide. Acid  Having a pH of less than 7. Action threshold  A level of pest abundance that stimulates action to protect plants from serious damage. Adaxial  Facing away from the stem of a plant. Adult  The sexually mature stage of an animal; usually the winged stage in insects; this stage does not molt. Aestivation  A state of inactivity during the summer months. Alate  Bearing wings. Alkaline  Basic, having a pH greater than 7. Allelopathy  The ability of a plant species to produce substances that are toxic to certain other plants. Anal plate  The shield-like plate or dorsal covering on the terminal segment in caterpillars, usually dark in color. Annual  A plant that normally completes its life cycle of seed germination, vegetative growth, reproduction, and death in a single growing season or year. Antenna (pl., antennae)  The paired segmented sensory organs, borne one on each side of the head, commonly protruding forward and termed horns or feelers. Anterior  In front; usually used to refer to the end of the body containing the head. Apical  Pertaining to the apex or outer end. Apterous  Lacking wings. Arista  A large hair or bristle on the antenna of flies. Axil  The upper angle where a leaf or twig joins the stem from which it grows. Bacillus thuringiensis  A bacterium that causes disease in insects; formulations of the bacteria are used as insecticides, with different strains used for suppression of caterpillars, beetles, and flies. Band  A transverse line, usually wide, crossing the body (often confused with “stripe,” a term used to designate a longitudinal line running the length of the body). Band application  An application in which an insecticide is applied in strips, usually to the bed or seed row. Basal  At or pertaining to the base or point of attachment, or nearest the main body.

Beneficials  Organisms that provide a benefit to crop production, especially natural enemies of pests and plant pollinators such as bees. Biennial  A plant that completes its life cycle in more than 1 year and usually does not flower until the second growing season. Binomial sampling  A sampling method that involves recording only the presence or the absence of members of the population being sampled (such as an insect pest) on a sample unit (such as a leaf), rather than counting the numbers of individuals; the p­ resence-absence sampling. Biological control  The action of parasites, predators, or pathogens in maintaining another organism’s population density at a lower average level than would occur in their absence. Biological control may occur naturally in the field or result from manipulation or introduction of biological control agents. Biotic disease  Disease caused by a pathogen, such as a bacterium, fungus, or virus. Botanical  Derived from plants or plant parts; often used to describe insecticides derived from plant. Brachypterous  Having short wings or elytra. Bract  A modified leaf at the base of a flower. Broadcast application  The application of a material such as an insecticide to the entire surface of a field. Brood  All the individuals of a generation; insects that hatch at about the same time. Bt  Acronym for Bacillus thuringiensis. C  Centigrade; a unit of temperature on the centigrade thermometer. Calibrate  To standardize or correct the measuring devices on instruments; to adjust nozzles on a spray apparatus. Canker  A dead, discolored, often sunken area (lesion) on a root, trunk, stem, or branch. Canopy  The leafy part of plants or trees. Carcinogen  A substance or agent capable of causing ­cancer. Carnivorous  Feeding on animals. Caterpillar  The larva of a butterfly, moth, sawfly, or scorpionfly. Catkin  A cluster of flowers in the form of a spike. Cauda  The pointed tip of the abdomen in aphids. 665

666 Glossary

Caudal  Pertaining to the anal end of the insect body. Cersus  A process located laterally near the tip of the abdomen; often an important diagnostic character in identification. Cervical shield  Plate on the dorsal surface of caterpillars just behind the head; also known as thoracic plate. Chelicerae  The pincerlike first pair of appendages in arachnids. Chlorinated hydrocarbons  A class of synthetic insecticides containing chlorine as one of the constituents; chlorinated hydrocarbon insecticides are typically very persistent and formerly were used widely for soil and seed treatments; also known as organochlorines. Chlorophyll  The green pigment of plants that captures, from sunlight, the energy necessary for photosynthesis. Chlorosis  Yellowing or bleaching of normal green plant tissue usually caused by the loss of chlorophyll. Chorion  The outer layer of an insect egg. Circulative virus  A virus that systemically infects its insect vector and usually is transmitted for the remainder of the vector’s life. Cladophyll  In plants, a branchlet that functions as a leaf. Claw  A hollow, sharp organ, located at the tip of the tarsus (foot). Clublike  A structure with a knob or swelling distally, usually used in reference to antennae. cm  Centimeter; a unit of length equivalent to 1/100 of a meter or 0.3937 in. Coccoon  A sheath, usually of silk, formed by an insect larva as a chamber for pupation. Collophore  A tubelike structure located ventrally on the first abdominal segment of springtails (Collembola). Companion planting  The practice of planting certain plant species often herbs on close association with crop plants to repel the practice of planting certain pests. Complete metamorphosis  Change in body form in which the insect displays the egg, larval, pupal, and adult stages. Corium  The thickened basal region of the front wing in Hemiptera. Cornicle  Two tubular structures located on the posterior part of an aphid’s abdomen. Cortex  Tissue between the phloem and the epidermis in roots and stems. Costa  The basal segment of the leg, articulating with the body. Cotyledon  A leaf formed within the seed and present on a seedling at germination; seed leaf. Cover crops  Cultivation of a second type of crop primarily to improve the production system for a primary crop (e.g.) legumes maintained during the winter season to improve soil condition. Crawler  The active first instar of a scale insect. Crochets  Tiny hooks on the prolegs of caterpillars.

Cross-resistance  In pest management, resistance of a pest population to a pesticide to which it has not been exposed that accompanies the development of resistance to a pesticide to which it has been exposed. Crown  The basal region of a plant, usually at the juncture of the stem and roots. Cultivar  An agricultural plant variety or strain developed for specific horticultural properties. Day-degree  A unit combining temperature and time, and used to measure and predict growth of organisms; sometimes called degree-days. Developmental threshold  The lowest temperature at which growth will occur. Diapause  A period of dormancy (“hiberation”) in arthropods. Disk  A soil cultivator made up of many circular blades that act to break up the soil into smaller aggregates. Distal  Pertaining to the part of an appendage furthest from the body. Dormant  To become inactive during periods of cold weather. Dorsal  Referring to the upper surface. Economic threshold  A level of pest abundance or damage at which the cost of control equals the crop value gained from instituting the control procedure. Ectoparasite  A parasite that lives outside (on the surface) of the host. Egg  The first free-living stage of most animals, contained within a chorion (shell). Elytron (pl., elytra)  The thickened front wing of beetles, serving primarily for protection of the hind wings or flight wings. Endoparasite  A parasite that lives inside the host. Entomopathogenic  Causing disease and/or death in insects. Epicranium  The upper portion of the head. Epidermis  The outermost layer of living cells on a plant or animal. Extrafloral nectary  A nectar-secreting gland found outside the flower. F  Fahrenheit; a unit of temperature on the Fahrenheit thermometer. Fallow  Cultivated land allowed to remain free of crops during the normal growing season. Femur (pl., femora)  A segment of the leg between the trochanter and tibia; of one of the two largest portions of the insect leg, often expanded to enhance hopping. Frass  The solid fecal material produced by insects. Fumigation  Treatment that uses a pesticide applied in a gaseous form. Fungicide  A pesticide used to manage growth of fungi. Furcula  A forked structure found beneath the abdomen of springtails (Colembolla), attached apically, and used for leaping.

Glossary  667

Gaster  The swollen, terminal abdominal segments of ants behind the constriction or pedicel. Genitalia  The modified abdominal segments used in copulation. h  Hour; a unit of time equivalent to 1/24 of a day. Halter (pl., halteres)  The vestigial, modified hind wings of Diptera that function as balancing organs. Hemelytra  The front wings in Hemiptera, in which the basal portion is thickened and the distal portion membranous. Herbivore  Animal that consumes plant tissue. Herbivory  Consumption of plant material. Honeydew  Sugary anal secretions of aphids and some other insects. Horticultural oil  Petroleum or botanical oil used to control pests on plants. Host  An organism that provides food and/or shelter for another organism. Hyphae  The thread-like mycelial tissue of fungi. Immature  The feeding stage of insects after birth but before adulthood, includes both larvae and nymphs. Incomplete metamorphosis  Change in body form in which the insect displays the egg, nymphal, and adult stages. Infection  The entry and establishment of a pathogen into a host. Infestation  The presence of a number of pests in a field, on plants, or in the soil. Inoculum  A small number of pests or pathogens that can, or does, lead to a greater abundance, and subsequent damage. Inorganic  Not containing the element carbon; usually derived from naturally occurring minerals. Insecticide  A pesticide applied to manage insect populations. Instar  The stage of larvae and nymphs between molts. Integrated pest management  An approach to prevention of damage by pests that focuses on the long-term suppression of pest abundance through a combination of environmentally sound techniques such as biological control and resistant plant varieties, rather than depending solely on chemical insecticides; insecticides are applied only when needed rather than routinely; also known as “IPM.” Integument  The outer covering or cuticle of an insect. Invertebrate  Animals lacking and internal skeleton or “backbone,” such as insects, mites, and worms. IPM  See integrated pest management. Juvenile  The immature form of a nematode; the stages between the egg and adult form. Larva (pl., larvae)  The growing stage of an insect with complete metamorphosis; the feeding stage between the egg and pupal stages; in mites the immature stages are also called larvae; larval insects usually bear little resemblance to the adult stage.

Lesion  A localized area of damage or infection, usually discolored or deformed. Lodging  Toppling of crop plants, often from wind or rain, and ultimately due to destruction of roots by insects. m  Meter; a unit of length equivalent to 39.37 in. Mandible  One of the pair of jaws in insects, attached to the head; usually stout and tooth-like in appearance. Median  At or near the middle. Meristem  The cells capable of division located at the growing point of a plant. Mesothorax  The second or middle thoracic segment; the segment bearing the middle pair of legs; the segment bearing the first pair of wings. Metamorphosis  A change in body form as the insect grows from the immature to the adult stage. Microbial pesticide  Microbial organisms that are applied like chemical pesticides for the suppression of pest abundance; various bacterial, fungal, viral, and microsporidian pathogens registered with the Environmental Protection Agency for use as a pest control agent. Middorsal  Refers to the middle of the upper region or “back” (as opposed to the subdorsal or slightly to the side or lateral). Mineral oil  Horticultural oil that is derived from petroleum. Miticide  A pesticide used to control mites; also called acaricide. mm  Millimeter; a unit of length equivalent to 1/10 of a centimeter or 0.0394 in. Molt  To shed or cast off the outer body covering, a necessary prerequisite to grow and attain the adult stage. Monitoring  Careful observation of pest abundance and damage; pest scouting. Motile  Active, able to move freely. Mulch  A layer of material placed on the soil to retard growth of weeds; in commercial crop production plastic is often used, whereas in the home garden organic materials such as leaves, straw, wood chips, and pine needles are often used. Mummy  The remains of an aphid whose insides have been consumed by a parasitoid. Natural enemies  Organisms normally killing arthropods that people consider to be pests without human intervention; for example, include predatory insects or vertebrates, insect parasitoids, and microbial pathogens causing disease. Neem  A botanical insecticide derived from the tropical tree Azadirachta indica; extracts of the foliage confer both feeding deterrent and toxic properties. Neonate  Recently hatched. Nocturnal  Active at night. Nonpersistent virus  A virus carried on the mouthparts of a vector, and usually lost after the insect feeds a few times; a stylet-borne virus.

668 Glossary

Nymph  The immature stage of insects with incomplete metamorphosis; the nymph usually resembles the adult in form except for undeveloped wings. Oil  See horticultural oil. Omnivorous  Having a broad diet, consisting of both plant and animal material. Organic  Chemically, this is a material that contains the elements carbon and hydrogen; agriculturally, this refers to plants cultured without synthetic fertilizers or pesticides. Organophosphate  A class of synthetic insecticides derived from phosphoric acid, capable of disrupting neurotransmission in insects and vertebrates. Oviparous  Producing eggs. Ovipositor  The structure, usually in the shape of a tube, that is used to deposit eggs. Palps  Small antennalike sensory appendages attached to the mouth. Parasite  An organism that obtains its food by feeding on the body of another organism, its host. Parasitoid  A parasite that kills its host at about the time the parasite completes its development. Parthenogenesis  Development from an egg that has not been fertilized. Pathogen  A disease-causing microbial organism. Pedicel  The constricted region of the abdomen in Hymenoptera; in ants the pedicel bears one or more upright lobes. Perennial  A plant that lives at least 3  years and reproduces at least twice. Persistent virus  A virus that passes through the body of the vector, and that usually persists for the remainder of the vector’s life. Pesticide  A material that kills pests; this term is often used to describe insecticides, which are pesticides that kill insects. Petiole  Stalk that connects the leaf to a stem; in Hymenoptera it is sometimes used in place of pedicel. Pheromone  Chemical substance secreted into the environment that affects the behavior or physiology of other members of the same species. Phytophagous  Feeding on plants or plant products. Phytotoxicity  Damage to a plant due to contact with a chemical toxin. Plumose  Feather-like in structure with a single thick stem and numerous parallel branches; usually used to describe antennae of Lepidoptera. Pollinator  The agent of pollen transfer in plants, often bees. Posterior  The hind region of the body, or referring to the end containing the anus. Postplant  Refers to treatments applied to a crop after planting. Preplant  Refers to treatments applied to a crop before planting.

Prepupa  An active but nonfeeding stage of insects; the period immediately preceding the molt to the adult stage. Proleg  A fleshy, unsegmented leg found on the abdomen of caterpillars. Pronotum  The upper or dorsal surface of the prothorax. Prothoracic plate  Equivalent to thoracic plate. Prothorax  The most anterior of the three thoracic segments. Proximal  Pertaining to the part of an appendage closer to the body. Pubescence  A covering of setae (hairs). Pubescent  Covered with hairlike structures (setae in insects, trichomes in plants). Punctate  Containing impressed points, punctures, or dimples. Pupa  The nonfeeding, immobile stage between the larval and adult stages in insects with complete metamorphosis; a stage where major reorganization of the body take place. Puparium  The hardened, thickened integument of the last instar larva of Diptera, in which the pupa is formed (plural, puparia). Pupate  To molt from the larval stage to the pupa. Pyrethroids  Synthetic insecticides that are structurally similar to the toxic components of pyrethrum; also called synthetic pyrethroids. Pyrethrum  Natural insecticide derived from certain plants in the genus Chrysanthemum; it is highly valued for its rapid effects on insects and low toxicity to mammals. Reservoir  A site where organisms can survive, usually in relatively small numbers, and then invade or repopulate an area. Resistant  Tolerant of conditions that are deleterious to other strains of the same species; usually applied to plant tolerance of pest damage, or arthropod tolerance of pesticides. Reticulations  A net-like structure, usually referring to the pigmented pattern on eyes of Lepidoptera. Rostrum  In weevils, the snoutlike prolongation of the head containing the mouthparts distally; in Hemiptera, this sometimes refers to the beak or piercing-sucking mouthparts. Rotation  In agriculture, purposeful alternation of crops grown on the same plot of land; in pest control, purposeful alternation of insecticides used to control a pest population. Row covers  A covering, usually consisting of spun-­ bonded polyester, that is placed over crops to protect them from adverse weather or pests. Sanitation  The practice of eliminating pests, or materials or sites that might harbor pests. Saprophagous  Feeding on dead or dying plant or animal tissue.

Glossary  669

Scale  A modified, flattened seta on the surface of an insect. Scientific name  A Latin or Latinized name given to all biological organisms and consisting of two parts, a genus and species; the scientific name often includes the name of the individual(s) originally describing the species. Scutellum  In Hemiptera, the triangular mesothoracic region between the base of the wings. Segment  A major subdivision of the body or appendage, separated from other segments by areas of flexibility. Selective pesticide  Pesticides that are toxic principally to the target pest; having few adverse effects on nontarget organisms. Semicircular  Like the half of a circle. Senescence  The period of life after maturity. Sequential sampling  A sampling protocol where the sampling continues only until a decision can be reached, rather than requiring a fixed number of samples to be taken. Serrate  Notched, like the teeth of a saw. Sessile  Immobile; incapable of moving. Sex pheromone  Pheromones that attract the opposite sex for mating; these are often used, in conjunction with traps, to monitor abundance of insects. Sheath  A structure enclosing others. Skeletonize  To remove the tissue of a leaf except for the veins, leaving a “skeleton.” Sooty mold  A dark-colored fungus growing on the honeydew secreted by insects, usually aphids or scales. Spermatophore  A covering or capsule around the sperm. Spindle-shaped  Elongate-cylindrical, thicker in the middle and tapering to each end. Spine  A large, stout seta, or thorn-like process. Spiracle  An external opening of the system of ducts used to transfer atmospheric gases into, and out of, the body of arthropods; they are commonly found along each side of the body. Spore  A reproductive stage of fungi, usually somewhat resistant to adverse environmental conditions, that is capable of growing into a new organism. sq  Square; a quantity multiplied by itself. Stemmata  The small, simple eyes found on some insects, usually on the side of the head. Stolon  A basal branch or stem that is inclined to root; a plant runner. Stridulate  Produce a noise by rubbing together two surfaces. Stripe  A line that runs horizontally or lengthwise on an insect (often confused with “band” or transverse line). Stylet  The elongated needle-like portions of the piercingsucking type of insect mouthparts. Stylet-borne virus  See nonpersistent virus. Subdorsal  A region between the dorsal and lateral areas. Submedial arc  Pigmentation in the form of an arc occurring on the face or top of the head in caterpillars.

Subspiracular  The area immediately below the spiracles. Supraspiracular  The area immediately above the spiracles. Taproot  A large primary root growing downwards, and giving rise to numerous, smaller lateral roots. Tarsus  The portion of the insect distal to the tibia, and often bearing claws; the foot. Tegmen (pl., tegmina)  The thickened front wing of Orthoptera and related insect, and the thickened basal portion of the front wings of Hemiptera. Thoracic plate  Shield-like dorsal covering or plat on the body segment immediately behind the head, usually dark in color; also known as cervical shield. Thorax  The second or middle of the three major body regions of insects, and the section bearing wings and jointed (true) legs. Thread-like  Long, thin structure, approximately equal in diameter throughout; usually used in reference to the antennae. Tibia  The section of the insect leg between the femur and the tarsus, usually one of the largest sections and often bearing spines or spurs. Trap crop  A crop or portion of a crop that is intended to lure insects away from the main crop. Trapezoidal  A four-sided figure in which two sides are parallel and two are not. Trichomes  Hairs or small spines on the surface of a plant. Trochanter  A small section of the leg connecting the coxa and femur. Tubercle  A small raised area or pimple; in caterpillars a hair often originates from these raised areas. Tympanum  A membrane-covered cavity on the thorax, abdomen, or leg of insects that functions like an ear. Variety  An identifiable strain of a species, usually bred for a particular horticultural purpose; also called a cultivar. Vascular system  The system of plant tissues that conducts water, minerals, and products of photosynthesis within the plant. Vein  A tube running through the wings of insect, through which blood is pumped. Ventral  The lower surface. Virulence  The ability of a pathogen to infect a host and cause disease. Viviparous  Bearing living young, as opposed to eggs. Vestigial  Small or degenerate; the remains of a previously functioning organ. Volunteer plants  The unexpected and undesired emergence of plants, usually self-seeded by the previous plants. Whorl  The arrangement of leaves in a circle around the stem. Wing  Paired membranous flight organs of insects, originating at the mesothorax and metathorax; the number varies (0, 2, or 4) and they may or may not be functional. Wing pad  The underdeveloped wings of nymphs.

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Index Note: Page numbers followed by f indicate figures and t indicate tables.

A

abbreviatus, Diaprepes, 156–158 abbreviatus, Neoscapteriscus, 573 abrupta, Empoasca, 322–323 absoluta, Tuta, 442–443 Acalymma trivittatum, 120–122 Acalymma vittatum, 120 Acanthoscelides obtectus, 89–90 Acari banks grass mite, 601–602 broad mite, 602–604 bulb mites, 604–605 strawberry spider mite, 605 tomato russet mite, 608–609 tumid spider mite, 605–608 twospotted spider mite, 605 Acaridae, 604–605 Achyra rantalis, 409–410 acrea, Estigmene, 434–436 Acrididae American grasshopper, 555–557 differential grasshopper, 557–559 eastern lubber grasshopper, 559–562 migratory grasshopper, 562–567 redlegged grasshopper, 567 southern redlegged grasshopper, 567–569 two-striped grasshopper, 569–571 Acrolepiidae, 389–390 Acrolepiopsis assectella, 389–390 Aculops lycopersici, 608–609 Acyrthosiphon kondoi, 297–300 Acyrthosiphon pisum, 297 Adelphocoris lineolatus, 333–335 Adelphocoris rapidus, 338–339 Adoretus sinicus, 187–188 Adult flea beetles, 642 Adult slugs, 649 Adult snails, 650 aegopodii, Cavariella, 308–310 aeneus, Chalcodermus, 140–141 aeripennis, Selatosomus, 165–167 African earwig damage, 209 distribution, 207 host plants, 208 life cycle, 208 management, 209 natural enemies, 208 agonus, Limonius, 170 Agriotes mancus, 172–174 Agrius cingulatus, 546–547

Agroiconota bivittata, 97–98 Agromyza parvicornis, 216–217 Agromyzidae American serpentine leafminer, 211–213 asparagus miner, 213–215 cabbage leafminer, 215–216 characteristics, 9 corn blotch leafminer, 216–217 pea leafminer, 217–219 vegetable leafminer, 219–221 Agrotis ipsilon, 462–465 Agrotis orthogonia, 491–494 Agrotis subterranea, 487–489 albicosta, Striacosta, 516–518 albionica, Phyllotreta, 64–65 albopunctatus, Arvelius, 368–369 Aleyrodidae characteristics, 11 greenhouse whitefly, 259–262 sweetpotato whitefly, 262–266 Alfalfa caterpillar, 525–527 Alfalfa looper, 449–450 Alfalfa plant bug, 333–335 Alfalfa webworm, 393–395 Alliaceae, 5 Alticinae cabbage flea beetle, 64–65 characteristics, 8 corn flea beetle, 65 crucifer flea beetle, 67–69 desert corn flea beetle, 69–71 eggplant flea beetle, 71–72 elongate flea beetle, 74–76 hop flea beetle, 72–73 horseradish flea beetle, 73–74 palestriped flea beetle, 74 potato flea beetle, 76 redheaded flea beetle, 78 smartweed flea beetle, 78–79 southern tobacco flea beetle, 83–85 spinach flea beetle, 79 striped flea beetle, 81 sweetpotato flea beetle, 82–83 three-spotted flea beetle, 79 tobacco flea beetle, 83 toothed flea beetle, 65–67 tuber flea beetle, 85–87 western black flea beetle, 87–88 western potato flea beetle, 76–78 western striped flea beetle, 81–82 yellow-necked flea beetle, 80 Zimmermann’s flea beetle, 88–89

Amaranthaceae, 3 Amaryllidaceae, 5 Amblyomma americanum, 387 americana, Entomoscelis, 114–115 americana, Schistocerca, 555–557 American bean thrips, 581–600 American grasshopper, 555–557 American serpentine leafminer, 211–213 americanum, Amblyomma, 387 amplicollis, Conoderus, 167–170 Anabrus simplex, 576 Anagrapha falcifera, 471–473 Anaphothrips obscurus, 588–589 Anasa andresii, 326–329 Anasa armigera, 326–329 Anasa scorbutica, 326–329 Anasa tristis, 326–329 anastomasalis, Omphisa, 428–429 andresii, Anasa, 326–329 Anise swallowtail, 523–525 anisopliae, Metarhizium, 205–206 annonae, Euxesta, 255–257 annulipes, Euborellia, 207 Anomala orientalis, 194–196 Anomis erosa, 433–434 Anthomyiidae bean seed maggot, 221–222 beet leafminer, 222 cabbage maggot, 224–227 onion maggot, 227–231 radish root maggot, 231–232 seedcorn maggot, 232–234 spinach leafminer, 222–224 turnip root maggot, 235–236 Anthonomus eugenii cano, 146–148 Anthonomus grandis, 387 antiqua, Delia, 227–231 Apamea devastator, 486–487 Aphididae artichoke aphid, 266–267 asparagus aphid, 267–268 bean aphid, 268–271 bean root aphid, 271 bird cherry-oat aphid, 271–273 blue alfalfa aphid, 297–300 buckthorn aphid, 273–275 cabbage aphid, 275–278 carrot root aphids, 278 characteristics, 11 coriander aphid, 278 corn leaf aphid, 279–281 corn root aphid, 281–283

789

790  Index

Aphididae (Continued) cowpea aphid koch, 283–285 foxglove aphid, 285–286 green peach aphid, 286–290 honeysuckle aphid, 278–279 lettuce aphid, 290–292 lettuce root aphid, 292–294 melon aphid, 294–297 pea aphid, 297 potato aphid, 300–303 rice root aphid, 303–304 sugarbeet root aphids, 304–306 turnip aphid, 306–308 willow-carrot aphid, 308–310 Aphid abdomen with dorsal projections, 629f characteristics, 11 immature, 630f membranous wings and wing vines, 627f Aphis fabae, 268–271 Aphis gossypii, 294–297 Aphis maidiradicis, 281–283 Aphis nasturtii, 273–275 Apiaceae, 4 arboricola, Strigoderma, 197–198 Argidae, 12, 383–384 Argid sawflies, 12 Argus tortoise beetle, 97 Armadillidiidae, 613 Armadillidium vulgare, 613 armigera, Anasa, 326–329 armigera, Helicoverpa, 480–482 armoraciae, Phyllotreta, 73–74 Army cutworm, 450–452 Armyworms, 13, 452–455, 635–639 Artichoke aphid, 266–267 Artichoke plume moth, 538–539 Arvelius albopunctatus, 368–369 asahinai, Blattella, 55–57 Ascia monuste, 532–534 Asian cockroach, 55–57 Asiatic garden beetle, 184–186 Asparagaceae, 3 asparagi, Brachycorynella, 267–268 asparagi, Crioceris, 99–101 Asparagus, pest identification, 3, 43–44 Asparagus aphid, 267–268 Asparagus beetle, 99–101 Asparagus miner, 213–215 assectella, Acrolepiopsis, 389–390 Aster leafhopper, 312–315 Aulacorthum solani, 285–286 auricularia, Forficula, 205–209 Autographa californica, 449–450 Autographa precationis, 494 Autoplusia egena, 455–456 auxiliaris, Euxoa, 450–452

B

Bactericera cockerelli, 377–379 Bactrocera cucurbitae, 247–249 Bactrocera dorsalis, 249–250 Bagrada hilaris, 357–360

balteata, Diabrotica, 101–103 Banded cucumber beetle, 101–103 Banded woollybear, 429–431 Banks grass mite, 601–602 barberi, Diabrotica, 111–114 bassiana, Beauveria, 384 Bean, pest identification, 3, 44–45 Bean aphid, 268–271 Bean leaf beetle, 103–105 Bean leafroller, 445–447 Bean leafskeletonizer, 455–456 Bean root aphid, 271 Bean seed beetles, 8, 89–90 Bean seed maggot, 221–222 Bean thrips, 581–583 Bean weevil. See Bean seed beetles Beauveria bassiana, 60, 384 Bedellia orchilella, 391–392 Bedellia somnulentella, 391 Bedellidae morningglory leaf miner, 391 sweetpotato leaf miner, 391–392 Bedellid moths, 391–392 Beet, pest identification, 45 Beet and related crops, 3 Beet armyworm, 56–57, 456–459 Beet leafhopper, 315–318 Beet leafminer, 222 Beet webworm, 395–397 Bemisia tabaci, 262–266 Bertha armyworm, 459–461 betae, Pegomya, 222 betae, Pemphigus, 304–306 betae, Smynthurodes, 271 bicolor, Charidotella, 97 Bigonicheta spinipennis, 205–206 Big-Stem Jersey, 384 bilineatus, Pangaeus, 329–330 biloba, Megalographa, 461–462 Bilobed looper, 461–462 bipunctalis, Herpetogramma, 410–413 Bird cherry-oat aphid, 271–273 bispinosa, Frankliniella, 586–588 bivittata, Agroiconota, 97–98 bivittatus, Melanoplus, 569–571 Blaberidae damage, 58 management, 59 natural history, 57–58 Black blister beetle, 174–176 Black cutworm, 462–465 Blacklegged tortoise beetle, 97 Black swallowtail, 523 blanda, Systena, 74 Blapstinus spp., 203–204 Blattella asahinai, 55–57 Blattodea, 7 Blissidae, chinch bug, 310–312 Blissid bugs, 310–312 Blissus leucopterus, 310–312 Blister beetles, 8–9 Blue alfalfa aphid, 297–300 Blue butterflies, 13 Boethus schizoceri, 383

borealis, Epilachna, 132–134 borellii, Neoscapteriscus, 573 Borers, 13–14, 393–429 Bourletiella hortensis, 609–611 Brachycorynella asparagi, 267–268 bractatus, Microtechnites, 336–338 Bradybaena spp., 619–622 Brassicae, 3–4 brassicae, Brevicoryne, 275–278 brassicae, Liriomyza, 215–216 Brentidae characteristics, 8 sweetpotato weevil, 151–154 Brevicoryne brassicae, 275–278 Broadbean seed beetle, 90–92 Broadbean weevil. See Broadbean seed beetle Broad mite, 602–604 Bronzed cutworm, 465–467 Brown marmorated stink bug, 345–348 Brown stink bug, 348–350 Bruchinae bean seed beetle, 89–90 broadbean seed beetle, 90–92 characteristics, 8 cowpea seed beetle, 92 cultural manipulations, 89–96 pea seed beetle, 94–96 southern cowpea seed beetle, 92–94 Bruchus pisorum, 94–96 Bruchus rufimanus, 90–92 brunnea, Colaspis, 109–111 Buckthorn aphid, 273–275 Budworms, 13, 393–429 Bugs, 10–12 Bulb mites, 604–605 Burrower bugs, 10, 329–330 bursarius, Pemphigus, 292–294

C

Cabbage, pest identification, 45–46 Cabbage and related crops, 3–4 Cabbage aphid, 275–278 Cabbage budworm, 398 Cabbage crane fly, 253–255 Cabbage curculio, 134–135 Cabbage flea beetle, 64–65 Cabbage leafminer, 215–216 Cabbage looper, 467–471 Cabbage maggot, 224–227 Cabbage seedpod weevil, 135–137 Cabbage webworm, 398 Cabbage white butterflies, 639 Cabbage worms, 13, 525–534, 639 calabaza, Melittia, 543–546 californica, Autographa, 449–450 californicus, Limonius, 170 Caliothrips fasciatus, 581–583 Caliothrips phaseoli, 581–600 Callosobruchus chinensis, 92–94 Callosobruchus maculatus, 92 callosus, Sphenophorus, 141–143 canus, Limonius, 170–172 capitata, Ceratitis, 244–247

Index  791

Capitophorus elaeagni, 266–267 Carabidae characteristics, 7 ground beetles, 7, 63–64 seedcorn beetle, 63 slender seedcorn beetle, 63–64 Carcinophoridae, 207–209 carduidactyla, Platyptilia, 538–539 Carpophilus lugubris, 181–183 Carrot, pest identification, 46–47 Carrot and related crops, 4 Carrot beetle, 186–187 Carrot plant bug, 335–336 Carrot root aphids, 278 Carrot rust fly, 240–243 Carrot weevil, 137 cassidea, Chelymorpha, 97 Cassidinae argus tortoise beetle, 97 blacklegged tortoise beetle, 97 characteristics, 8 golden tortoise beetle, 97 mottled tortoise beetle, 97 striped tortoise beetle, 97–98 castanea, Maladera, 184–186 Caterpillars, 12–14 Cavariella aegopodii, 308–310 Cecidomyiidae, 236–238 Celery leaftier, 400 Celery looper, 471–473 Celeryworms, 13, 523–525 Cepaea spp., 619–622 Ceratitis capitata, 244–247 cereralis, Loxostege, 393–395 Cerotoma trifurcata, 103–105 Ceutorhynchus obstrictus, 135–137 Ceutorhynchus rapae, 134–135 Chaetocnema confinis, 82–83 Chaetocnema denticulata, 65 Chaetocnema ectypa, 69–71 Chaetocnema pulicaria, 65 Chaetopsis massyla, 255–257 Chalcodermus aeneus, 140–141 Charidotella bicolor, 97 Chelymorpha cassidea, 97 Chenopodiaceae, 3 Chilli thrips, 583–585 Chinavia hilaris, 351–354 Chinch bug, 310–312 chinensis, Callosobruchus, 92–94 Chinese rose beetle, 187–188 Chloridea virescens, 511–513 Chlorochroa sayi, 363 Chlorochroa uhleri, 363–365 Chrysodeixis includens, 500–502 Chrysomelidae argus tortoise beetle, 97 asparagus beetle, 99–101 banded cucumber beetle, 101–103 bean leaf beetle, 103–105 bean seed beetle, 89–90 blacklegged tortoise beetle, 97 broadbean seed beetle, 90–92 cabbage flea beetle, 64–65

characteristics, 7–8 colorado potato beetle, 105–109 corn flea beetle, 65 cowpea seed beetle, 92 crucifer flea beetle, 67–69 desert corn flea beetle, 69–71 eggplant flea beetle, 71–72 elongate flea beetle, 74–76 golden tortoise beetle, 97 grape colaspis, 109–111 hop flea beetle, 72–73 horseradish flea beetle, 73–74 mottled tortoise beetle, 97 northern corn rootworm, 111–114 palestriped flea beetle, 74 pea seed beetle, 94–96 potato flea beetle, 76 redheaded flea beetle, 78 red turnip beetle, 114–115 smartweed flea beetle, 78–79 southern cowpea seed beetle, 92–94 southern tobacco flea beetle, 83–85 spinach flea beetle, 79 spotted asparagus beetle, 115–116 spotted cucumber beetle, 116–120 striped cucumber beetle, 120 striped flea beetle, 81 striped tortoise beetle, 97–98 sweetpotato flea beetle, 82–83 sweetpotato leaf beetle, 122–123 three-spotted flea beetle, 79 tobacco flea beetle, 83 toothed flea beetle, 65–67 tuber flea beetle, 85–87 western black flea beetle, 87–88 western corn rootworm, 123–127 western potato flea beetle, 76–78 western striped cucumber beetle, 120–122 western striped flea beetle, 81–82 yellowmargined leaf beetle, 127–129 yellow-necked flea beetle, 80 Zimmermann’s flea beetle, 88–89 Cicadellidae aster leafhopper, 312–315 beet leafhopper, 315–318 corn leafhopper, 318–320 potato leafhopper, 320–322 western potato leafhopper, 322–323 cicticollis, Euborellia, 207 cingulatus, Agrius, 546–547 Circulifer tenellus, 315–318 Clearwing moths, 543–546 Click beetles, 8 Clivina impressifrons, 63–64 Clover cutworm, 473–474 clypealis, Leptoglossus, 323–326 c-nigrum, Xestia, 502–504 Coccinellidae lady beetle, 8 Mexican bean beetle, 129–132 squash beetle, 132–134 cockerelli, Bactericera, 377–379 Colaspis brunnea, 109–111 Coleoptera, 7–9, 629f

Colias eurytheme, 525–527 Colinus virginianus, 58 Collembola, 609–611 Colorado potato beetle, 105–109 comma, Stenolophus, 63 Common blossom thrips, 585–586 Common name, vegetables, 651–658t Common pillbug, 613 communis, Melanotus, 162 Compositae, 4–5 concavus, Lixus, 150–151 configurata, Mamestra, 459–461 confinis, Chaetocnema, 82–83 Coniontis spp., 203–204 Conoderus amplicollis, 167–170 Conoderus falli, 167 Conoderus vespertinus, 167 Consperse stink bug, 350–351 conspersus, Euschistus, 350–351 Contarinia nasturtii, 236–238 Convolvulaceae, 6 Copper butterflies, 13 Coptotermes formosanus, 59–62 Coreidae leaffooted bugs, 323–326 squash bugs, 326–329 Coriander aphid, 278 coriandri, Hyadaphis, 278 Corimelaena pulicaria, 379–380 Corn blotch leafminer, 216–217 Corn delphacid, 330–332 Corn earworm, 474–479 Corn flea beetle, 65 Corn leaf aphid, 279–281 Corn leafhopper, 318–320 Corn root aphid, 281–283 Cornsilk flies, 255–257 Cornu spp., 619–622 Corn wireworm, 162 Cotinis nitida, 189–191 Cotton mealybug, 369–371 Cotton stainer, 375–376 Coulee cricket, 576–579 Cowpea aphid koch, 283–285 Cowpea curculio, 140–141 Cowpea seed beetle, 92 Cowpea weevil. See Cowpea seed beetle Crambidae alfalfa webworm, 393–395 beet webworm, 395–397 cabbage budworm, 398 cabbage webworm, 398 celery leaftier, 400 characteristics, 13 cross-striped cabbage worm, 402–403 cucumber moth, 413–416 European corn borer, 403–407 European pepper moth, 407–409 false celery leaftier, 400–401 garden webworm, 409–410 hawaiian beet webworm, 410 melonworm, 413 oriental cabbage webworm, 398–400 pickleworm, 416–418

792  Index

Crambidae (Continued) purplebacked cabbage worm, 418–419 southern beet webworm, 410–413 southern cornstalk borer, 421–422 southwestern corn borer, 422–425 spotted beet webworm, 410 sugarcane borer, 425–427 sweetpotato leaf folder, 427–428, 510–511 sweetpotato vine borer, 428–429 crambidoides, Diatraea, 421–422 Crambus spp., 419–421 Crane flies, 253–255 crataegi, Dysaphis, 278 Crickets, vegetable-feeding, 38t Crioceris asparagi, 99–101 Crioceris duodecimpunctata, 115–116 Cross-striped cabbage worm, 402–403 Cruciferae, 3–4 cruciferae, Phyllotreta, 67–69 Crucifer flea beetle, 67–69 cubae, Faustinus, 134–135 Cuban pepper weevil, 134–135 Cucumber moth, 413–416 cucumeris, Epitrix, 76 Cucurbitaceae, 5 cucurbitae, Bactrocera, 247–249 cucurbitae, Melittia, 543 Curculionidae anthonomus eugenii cano, 146–148 cabbage curculio, 134–135 cabbage seedpod weevil, 135–137 carrot weevil, 137 cowpea curculio, 140–141 cuban pepper weevil, 134–135 maize billbug, 141 pea leaf weevil, 143–146 pepper weevil, 146–148 potato stalk borer, 148–150 rhubarb curculio, 150–151 southern corn billbug, 141–143 texas carrot weevil, 137–140 vegetable weevil, 154–156 west Indian sugarcane rootstalk borer weevil, 156–158 west Indian sweetpotato weevil, 158–160 whitefringed beetle, 160–162 Curculios, 8 Cutworms, 13, 635–639 Cydia nigricana, 551–553 Cydnidae, 10, 329–330 Cylas formicarius, 151–154 Cyrtopeltis notata, 343–345

D

daguerrei, Solenopsis, 384–385 Dalbulus maidis, 318–320 Darkling beetles, 9 Darksided cutworm, 479–480 decemlineata, Leptinotarsa, 105–109 Delia antiqua, 227–231 Delia floralis, 235–236 Delia florilega, 221–222 Delia planipalpis, 231–232

Delia platura, 232–234 Delia radicum, 224–227 Deloyala guttata, 97 Delphacidae, 11–12, 330–332 denticulata, Chaetocnema, 65 Depressaria radiella, 392–393 Depressariidae, 392–393 Depressariid moths, 392–393 Dermaptera African earwig, 207–209 European earwig, 205–209 ringlegged earwig, 207 Deroceras spp., 615–619 Desert corn flea beetle, 69–71 destructor, Selatosomus, 165 devastator, Apamea, 486–487 Diabrotica balteata, 101–103 Diabrotica barberi, 111–114 Diabrotica undecimpunctata, 116–120 Diabrotica virgifera, 123–127 Diamondback moth, 534–538 Diaphania hyalinata, 413 Diaphania indica, 413–416 Diaphania nitidalis, 416–418 Diaphorina citri, 56–57 Diaprepes abbreviatus, 156–158 Diatraea crambidoides, 421–422 Diatraea grandiosella, 422–425 Diatraea saccharalis, 387, 425–427 Dicyphus hesperus, 343–345 Differential grasshopper, 557–559 differentialis, Melanoplus, 557–559 difficilis, Listroderes, 154–156 Dingy cutworm, 480–482 Diplopoda, 611–613 Diptera, 9–10 agromyzidae, 211–221 anthomyiidae, 221–236 cecidomyiidae, 236–238 drosophilidae, 238–240 psilidae, 240–243 syrphidae, 243–244 tephritidae, 244–253 tipulidae, 253–255 ulidiidae, 255–258 Discestra trifolii, 473–474 Disonycha mellicollis, 80 Disonycha triangularis, 79 Disonycha xanthomelas, 79 dolichos, Spodoptera, 508 dolosa, Xestia, 502–504 Dooryard sowbug, 613–615 dorsalis, Bactrocera, 249–250 dorsalis, Scirtothrips, 583–585 Drosophila spp., 238–240 Drosophilidae, 9, 238–240 Dryland wireworm, 165 duodecimpunctata, Crioceris, 115–116 Duponchelia fovealis, 407–409 Dusky sap beetle, 181–183 Dysaphis crataegi, 278 Dysaphis foeniculus, 278 Dysdercus suturellus, 375–376

E

Eastern field wireworm, 170 Eastern lubber grasshopper, 559–562 Ebony bugs, 11 echinopus, Rhizoglyphus, 604–605 Ectobiidae damage, 56–57 management, 57 natural history, 55–56 ectypa, Chaetocnema, 69–71 egena, Autoplusia, 455–456 Eggplant flea beetle, 71–72 Eggplant lace bug, 380–382 Eggplant leaf miner, 438–439 Eggplant tortoise beetle, 99 elaeagni, Capitophorus, 266–267 Elasmopalpus lignosellus, 539–542 Elateridae corn wireworm, 162 dryland wireworm, 165 eastern field wireworm, 170 great basin wireworm, 165 gulf wireworm, 167–170 oregon wireworm, 162–165 pacific coast wireworm, 170–172 prairie grain wireworm, 165 puget sound wireworm, 165–167 southern potato wireworm, 167 sugarbeet wireworm, 170 tobacco wireworm, 167 wheat wireworm, 172–174 electa, Zonosemata, 251–253 Eleodes spp., 203–204 elisus, Lygus, 339–343 elongata, Systena, 74–76 Elongate flea beetle, 74–76 eluta, Euxesta, 255–257 Empoasca abrupta, 322–323 Empoasca fabae, 320–322 Engytatus modestus, 343–345 Entomoscelis americana, 114–115 Epicauta immaculata, 177–178 Epicauta maculata, 178–179 Epicauta pensylvanica, 174–176 Epicauta vittata, 179–181 Epilachna borealis, 132–134 Epilachna varivestis, 129–132 Epitrix cucumeris, 76 Epitrix fasciata, 83–85 Epitrix fuscula, 71–72 Epitrix hirtipennis, 83 Epitrix subcrinita, 76–78 Epitrix tuberis, 85–87 Erebidae banded woollybear, 429–431 characteristics, 12–13 green cloverworm, 431–433 okra caterpillar, 433–434 saltmarsh caterpillar, 434–436 yellow woollybear, 436–438 eridania, Spodoptera, 498–500 Eriophyidae, tomato russet mite, 608–609 erosa, Anomis, 433–434 Estigmene acrea, 434–436

Index  793

Etiella zinckenella, 542–543 Euborellia annulipes damage, 209 distribution, 207 host plants, 208 life cycle, 208 management, 209 natural enemies, 208 Euborellia cicticollis damage, 209 distribution, 207 host plants, 208 life cycle, 208 management, 209 natural enemies, 208 Euetheola humilis, 198–200 Euleia fratria, 251 Eumerus strigatus, 243 Eumerus tuberculatus, 243–244 euphorbiae, Macrosiphum, 300–303 European corn borer, 403–407 European earwig cultural practices, 207 damage, 207 distribution, 205 host plants, 205 insecticides, 207 life cycle, 206 natural enemies, 205–206 sampling, 207 European marsh crane fly, 253 European pepper moth, 407–409 eurytheme, Colias, 525–527 Euscepes postfasciatus, 158–160 Euschistus conspersus, 350–351 Euschistus servus, 348–350 Euschistus variolarius, 356–357 Euxesta annonae, 255–257 Euxesta eluta, 255–257 Euxesta stigmatias, 255–257 Euxoa auxiliaris, 450–452 Euxoa messoria, 479–480 Euxoa ochrogaster, 496–498 Evergestis pallidata, 418–419 Evergestis rimosalis, 402–403 exigua, Spodoptera, 456–459

F

Fabaceae, 3 fabae, Aphis, 268–271 fabae, Empoasca, 320–322 falcifera, Anagrapha, 471–473 Fall armyworm, 482–486 Fall field cricket, 571 falli, Conoderus, 167 False celery leaftier, 400–401 False chinch bug, 332–333 False diamondback moths, 389–390 False wireworms, 9, 203–204 fasciata, Epitrix, 83–85 fasciatus, Caliothrips, 581–583 Faustinus cubae, 134–135 Feltia jaculifera, 480–482 Feltia subgothica, 480–482

femurrubrum, Melanoplus, 567 Field crickets, 571–573 Flea beetle cabbage, 64–65 corn, 65 crucifer, 67–69 desert corn, 69–71 eggplant, 71–72 elongate, 74–76 hop flea, 72–73 horseradish, 73–74 palestriped, 74 potato, 76 redheaded, 78 smartweed, 78–79 southern tobacco, 83–85 spinach, 79 striped, 81 sweetpotato, 82–83 three-spotted, 79 tobacco, 83 toothed, 65–67 tuber, 85–87 western black, 87–88 western potato, 76–78 western striped, 81–82 yellow-necked, 80 Zimmermann, 88–89 Flies, vegetable-feeding, 33–34t floralis, Delia, 235–236 Florida flower thrips, 586–588 florilega, Delia, 221–222 Flower and bulb flies, 243–244 Flower flies, 9–10 foeniculi, Hyadaphis, 278–279 foeniculus, Dysaphis, 278 Forficula auricularia, 205–209 forficulae, Erynia, 205–206 Forficulidae, 207–209 formicarius, Cylas, 151–154 Formicidae, 12, 384–388 formosanus, Coptotermes, 59–62 Four-spotted sap beetle, 183–184 fovealis, Duponchelia, 407–409 Foxglove aphid, 285–286 Frankliniella bispinosa, 586–588 Frankliniella fusca, 595–597 Frankliniella occidentalis, 597–600 Frankliniella schultzei, 585–586 fratria, Euleia, 251 frontalis, Systena, 78 frugiperda, Spodoptera, 482–486 Fruit flies, 244–253 fusca, Frankliniella, 595–597 fuscula, Epitrix, 71–72

G

Gall midges, 236–238 Garden fleahopper, 336–338 Garden millipede, 611–613 Garden springtail, 609–611 Garden symphylan, 622–624 Garden webworm, 409–410 Gargaphia solani, 380–382

Gastropoda slugs, 615–619 snails, 619–622 Gelechid moths, 13 Gelechiidae characteristics, 13 eggplant leaf miner, 438–439 potato tuberworm, 439–442 tomato leaf miner, 442–443 tomato pinworm, 443–445 geminata, Solenopsis, 384 German cockroach, 56 gibbosus, Tomarus, 186–187 Glassy cutworm, 486–487 glaucus, Hadromorphus, 165 Glischrochilus quadrisignatus, 183–184 Golden tortoise beetle, 97 gossypii, Aphis, 294–297 gracilis, Oxidus, 611–613 Graminae, 5–6 grandiosella, Diatraea, 422–425 grandis, Anthonomus, 387 Granulate cutworm, 487–489 Grape colaspis, 109–111 Grasshoppers chewing mouthparts, 628f vegetable-feeding, 38t Grass thrips, 588–589 Gratiana pallidula, 99 Gray hairstreak, 447–449 Great basin wireworm, 165 Green cloverworm, 431–433 Greenhouse whitefly, 259–262 Green june beetle, 189–191 Green peach aphid, 286–290 Green stink bug, 351–354 Ground beetles, 7, 63–64 Gryllidae fall field cricket, 571 southeastern field cricket, 571–573 spring field cricket, 571 Gryllotalpidae shortwinged mole cricket, 573 southern mole cricket, 573 tawny mole cricket, 573–576 Gryllus pennsylvanicus, 571 Gryllus rubens, 571–573 Gryllus veletis, 571 guildinii, Piezodorus, 360–361 Gulf wireworm, 167–170 guttata, Deloyala, 97

H

Hadromorphus glaucus, 165 hageni, Reticulitermes, 59–62 Hairstreak butterflies, 13, 447–449 Halyomorpha halys, 345–348 halys, Halyomorpha, 345–348 Harlequin bug, 354–356 Hawaiian beet webworm, 410 Hawk moths, 13–14 Helicoverpa armigera, 480–482 Helicoverpa zea, 56–57, 474–479 Hellula phidilealis, 398

794  Index

Hellula rogatalis, 398 Hellula undalis, 398–400 Hemiptera aleyrodidae, 259–266 aphididae, 266–310 blissidae, 310–312 cicadellidae, 266–310 coreidae, 323–329 cydnidae, 329–330 delphacidae, 330–332 miridae, 333–345 vegetable-feeding, 35–36t Herpetogramma bipunctalis, 410–413 Hesperiidae, 13, 445–447 hesperus, Dicyphus, 343–345 hesperus, Lygus, 339 hilaris, Bagrada, 357–360 hilaris, Chinavia, 351–354 hirsutus, Maconellicoccus, 373–375 hirtipennis, Epitrix, 83 histrionica, Murgantia, 354–356 Honeysuckle aphid, 278–279 Hop flea beetle, 72–73 Hop vine borer, 494–496 Hornworms, 13–14, 546–551 Horseradish flea beetle, 73–74 hortensis, Bourletiella, 609–611 hudsonias, Systena, 78–79 humilis, Euetheola, 198–200 Hyadaphis coriandri, 278 Hyadaphis foeniculi, 278–279 hyalinata, Diaphania, 413 Hydraecia immanis, 494–496 Hydraecia micacea, 494 Hyles lineata, 550–551 Hymenia perspectalis, 410 Hymenoptera, 12 hyoscyami, Pegomya, 222–224 Hypena scabra, 431–433

I

immaculata, Epicauta, 177–178 immaculata, Scutigerella, 622–624 Immaculate blister beetle, 177–178 immanis, Hydraecia, 494–496 Immature psyllid, 630f Immature sawfly, 631f Imported cabbage worm, 527–530 impressifrons, Clivina, 63–64 includens, Chrysodeixis, 500–502 inconspicuella, Keiferia, 438–439 Indian cockroach, 57–59 indica, Diaphania, 413–416 indica, Pycnoscelus, 57 Insects, vegetable-feeding, 24–28t Invertebrate pests acari, 601–609 collembola, 609–611 diplopoda, 611–613 gastropoda, 615–622 isopoda, 613–615 symphyla, 622–624 invicta, Solenopsis, 384–388

ipsilon, Agrotis, 462–465 isabella, Pyrrharctia, 429–431 Isopoda common pillbug, 613 dooryard sowbug, 613–615

J

jaculifera, Feltia, 480–482 Japanese beetle, 191–194 japonica, Popillia, 191–194 Jonthonota nigripes, 97

K

Keiferia inconspicuella, 438–439 Keiferia lycopersicella, 443–445 Kneallhazia solenopsae, 384, 387 kondoi, Acyrthosiphon, 297–300

L

Lacebugs, 11 Lady beetles, 8 langei, Liriomyza, 217–219 latifascia, Spodoptera, 508–510 latipes, Mocis, 506–508 latus, Polyphagotarsonemus, 602–604 Leaf beetles, 7–8 Leaffooted bugs, 10, 323–326 Leafhopper aster, 312–315 beet, 315–318 corn, 318–320 pointed head and wing veins, 628f potato, 320–322 western potato, 322–323 Leafminer flies American serpentine, 211–213 asparagus miner, 213–215 cabbage, 215–216 corn blotch, 216–217 pea, 217–219 vegetable, 219–221 Leafminer moths, 438–445 Leafminers, 13 Leaf mining flies, 9 Leafroller moths, 551–553 Leafrollers, 14 Leaftiers, 13, 393–429 leconti, Stenolophus, 63–64 Leek moth, 389–390 Leguminosae, 3 leibyi, Schizocerophaga, 383 Leidyula spp., 615–619 Lepidoptera acrolepiidae, 1–3 crambidae, 6–11 depressariidae, 5 lyonetiidae, 4 Leptinotarsa decemlineata, 105–109 Leptoglossus clypealis, 323–326 Leptoglossus oppositus, 323–326 Leptoglossus phyllopus, 323–326 Leptoglossus zonatus, 323–326

Lesser bulb fly, 243–244 Lesser cornstalk borer, 539–542 Lettuce, pest identification, 47–48 Lettuce and related crops, 4–5 Lettuce aphid, 290–292 Lettuce root aphid, 292–294 leucopterus, Blissus, 310–312 lignosellus, Elasmopalpus, 539–542 Liliaceae, 3 Limabean pod borer, 542–543 Limax spp., 615–619 Limonius agonus, 170 Limonius californicus, 170 Limonius canus, 170–172 lineata, Hyles, 550–551 lineatus, Sitona, 143–146 lineolaris, Lygus, 339 lineolatus, Adelphocoris, 333–335 Lipaphis pseudobrassicae, 306–308 Liriomyza brassicae, 215–216 Liriomyza langei, 217–219 Liriomyza sativae, 219–221 Liriomyza trifolii, 211–213 Listroderes difficilis, 154–156 Listronotus oregonensis, 137 Listronotus texanus, 137–140 Little ebony bug, 379–380 Lixus concavus, 150–151 Loopers, 13, 640 Loxostege cereralis, 393–395 Loxostege sticticalis, 395–397 lugubris, Carpophilus, 181–183 Lycaenidae, 13, 447–449 lycopersicella, Keiferia, 443–445 lycopersici, Aculops, 608–609 lycopersici, Aculopsa, 608–609 Lygaeidae, 10, 332–333 Lygropia tripunctata, 427–428, 510–511 Lygus elisus, 339–343 Lygus hesperus, 339 Lygus lineolaris, 339

M

Maconellicoccus hirsutus, 373–375 Macrodactylus subspinosus, 196 Macrodactylus uniformis, 196–197 Macrosiphum euphorbiae, 300–303 Macrosteles quadrilineatus, 312–315 maculata, Epicauta, 178–179 maculatus, Callosobruchus, 92 Madeira mealybug, 371–373 madeirensis, Phenacoccus, 371–373 Maggots, 9–10 maidiradicis, Aphis, 281–283 maidis, Dalbulus, 318–320 maidis, Peregrinus, 330–332 maidis, Rhopalosiphum, 279–281 maidis, Sphenophorus, 141 Maize billbug, 141 Maladera castanea, 184–186 Malvaceae, 5 Mamestra configurata, 459–461 mancus, Agriotes, 172–174

Index  795

Manduca quinquemaculata, 547–550 Manduca sexta, 547 massyla, Chaetopsis, 255–257 Mealybugs, 11–12 Mediterranean fruit fly, 244–247 Megalographa biloba, 461–462 Melanchra picta, 522–523 Melanoplus bivittatus, 569–571 Melanoplus differentialis, 557–559 Melanoplus femurrubrum, 567 Melanoplus propinquus, 567–569 Melanoplus sanguinipes, 562–567 Melanotus communis, 162 Melanotus longulus oregonensis, 162–165 melinus, Strymon, 447–449 Melittia calabaza, 543–546 Melittia cucurbitae, 543 mellicollis, Disonycha, 80 Meloidae black blister beetle, 174–176 characteristics, 8–9 immaculate blister beetle, 177–178 spotted blister beetle, 178–179 striped blister beetle, 179–181 Melon aphid, 294–297 Melon fly, 247–249 Melon thrips, 589–591 Melonworm, 413 messoria, Euxoa, 479–480 Metarhizium anisopliae, 60, 205–206 Mexican bean beetle, 129–132 micacea, Hydraecia, 494 microptera, Romalea, 559–562 Microtechnites bractatus, 336–338 Microtheca ochroloma, 127–129 Migratory grasshopper, 562–567 Milax spp., 615–619 Millipedes, 611–613 minians, Nephelodes, 465–467 Miridae alfalfa plant bug, 333–335 carrot plant bug, 335–336 characteristics, 10 garden fleahopper, 336–338 pale legume bug, 339–343 rapid plant bug, 338–339 tarnished plant bug, 339 tomato plant bug, 343–345 western tarnished plant bug, 339 Mites banks grass, 601–602 broad mite, 602–604 bulb, 604–605 strawberry spider, 605 tomato russet, 608–609 tumid spider, 605–608 twospotted spider, 605 Mocis latipes, 506–508 modestus, Engytatus, 343–345 Mole crickets, 573–576 monuste, Ascia, 532–534 Mormon cricket, 576 Morningglory leaf miner, 391 Moths, vegetable-feeding, 36–37t

Mottled tortoise beetle, 97 Murgantia histrionica, 354–356 Mustard white, 530–531 myopaeformis, Tetanops, 257–258 Mythimna unipuncta, 452–455 Myzus persicae, 286–290

N

napi, Pieris, 530–531 Nasonovia ribisnigri, 290–292 nasturtii, Aphis, 273–275 nasturtii, Contarinia, 236–238 Naupactus spp., 160–162 nebris, Papaipema, 504–506 Neoscapteriscus abbreviatus, 573 Neoscapteriscus borellii, 573 Neoscapteriscus vicinus, 573–576 Nephelodes minians, 465–467 Nesidiocoris tenuis, 343–345 Nezara viridula, 365–368 niger, Nysius, 332–333 nigricana, Cydia, 551–553 nigripes, Jonthonota, 97 nigritus, Typophorus, 122–123 nitida, Cotinis, 189–191 nitidalis, Diaphania, 416–418 Nitidulidae characteristics, 9 dusky sap beetle, 181–183 four-spotted sap beetle, 183–184 ni, Trichoplusia, 467–471 Noctua pronuba, 518–519 Noctuidae alfalfa looper, 449–450 army cutworm, 450–452 armyworm, 452–455 bean leafskeletonizer, 455–456 beet armyworm, 456–459 bertha armyworm, 459–461 bilobed looper, 461–462 black cutworm, 462–465 bronzed cutworm, 465–467 cabbage looper, 467–471 celery looper, 471–473 characteristics, 13 clover cutworm, 473–474 corn earworm, 474–479 darksided cutworm, 479–480 dingy cutworm, 480–482 fall armyworm, 482–486 glassy cutworm, 486–487 granulate cutworm, 487–489 hop vine borer, 494–496 old world bollworm, 480–482 pale western cutworm, 491–494 plantain looper, 494 potato stem borer, 494 redbacked cutworm, 496–498 southern armyworm, 498–500 soybean looper, 500–502 spotted cutworm, 502–504 stalk borer, 504–506 striped grass looper, 506–508

sweetpotato armyworm, 508 tobacco budworm, 511–513 variegated cutworm, 513–516 velvet armyworm, 508–510 western bean cutworm, 516–518 western yellow-striped armyworm, 520–522 winter cutworm, 518–519 yellow-striped armyworm, 520 zebra caterpillar, 522–523 Noctuid moths, 13 Noninsect vegetable pests, 7 North American vegetable crops biological control, 19 characteristics, 3–6 cultural manipulations, 19 direct pests, 15 economic injury level, 15–16 economic pests, 15 esthetic/cosmetic pest, 15 estimated crop losses, 16, 16–17t farm gate value, 1 indirect pests, 15 insecticide use, 16–17, 20 insect injury, 17–18 insects and insect relatives, 6–15 pest management philosophy, 18–19 physical manipulations, 19–20 production data, 1, 2t species identification, 20–21 Northern corn rootworm, 111–114 notata, Cyrtopeltis, 343–345 nubilalis, Ostrinia, 403–407 Nysius niger, 332–333

O

obscurus, Anaphothrips, 588–589 obstrictus, Ceutorhynchus, 135–137 obtectus, Acanthoscelides, 89–90 occidentalis, Frankliniella, 597–600 ochrogaster, Euxoa, 496–498 ochroloma, Microtheca, 127–129 Ocytata pallipes, 205–206 Okra, pest identification, 48 Okra caterpillar, 433–434 Old world bollworm, 480–482 oleracea, Tipula, 253–255 Oligonychus pratensis, 601–602 Omphisa anastomasalis, 428–429 Onespotted stink bug, 356–357 Onion, pest identification, 48–49 Onion and related plants, 5 Onion bulb fly, 243 Onion maggot, 227–231 Onion thrips, 591–595 operculella, Phthorimaea, 439–442 Ophiomyia simplex, 213–215 oppositus, Leptoglossus, 323–326 Orange butterflies, 13 orchilella, Bedellia, 391–392 oregonensis, Listronotus, 137 Oregon wireworm, 162–165 Oriental beetle, 194–196 Oriental cabbage webworm, 398–400

796  Index

Oriental fruit fly, 249–250 orientalis, Anomala, 194–196 ornithogalli, Spodoptera, 520 orthogonia, Agrotis, 491–494 Orthops scutellatus, 335–336 Orthoptera acrididae, 555–571 gryllidae, 571–573 gryllotalpidae, 573–576 tettigoniidae, 576–579 Ostrinia nubilalis, 403–407 Oxidus gracilis, 611–613 Oxyspirura mansoni, 58 Oxyspirura parvorum, 58

P

Pacific coast wireworm, 170–172 padi, Rhopalosiphum, 271–273 Painted bug, 357–360 Pale legume bug, 339–343 Palestriped flea beetle, 74 Pale western cutworm, 491–494 pallidata, Evergestis, 418–419 pallidula, Gratiana, 99 pallipes, Ocytata, 205–206 palmi, Thrips, 589–591 paludosa, Tipula, 253 Pangaeus bilineatus, 329–330 Papaipema nebris, 504–506 Papilionidae anise swallowtail, 523–525 black swallowtail, 523 characteristics, 13 Papilio polyxenes, 523 Papilio zelicaon, 523–525 Paradoxosomatidae, 611–613 Parsnip leafminer, 251 Parsnip webworm, 392–393 parvicornis, Agromyza, 216–217 Pea aphid, 297 Pea leafminer, 217–219 Pea leaf weevil, 143–146 Pea moth, 551–553 Pea seed beetles, 8, 94–96 Pea weevil. See Pea seed beetle Pegomya betae, 222 Pegomya hyoscyami, 222–224 Pemphigus betae, 304–306 Pemphigus bursarius, 292–294 Pemphigus populivenae, 304–306 pennsylvanicus, Gryllus, 571 pensylvanica, Epicauta, 174–176 Pentatomidae brown marmorated stink bug, 345–348 brown stink bug, 348–350 consperse stink bug, 350–351 green stink bug, 351–354 harlequin bug, 354–356 onespotted stink bug, 356–357 painted bug, 357–360 redbanded stink bug, 360–361 redshouldered stink bug, 361–363 say stink bug, 363

southern green stink bug, 365–368 tomato stink bug, 368–369 uhler stink bug, 363–365 Pepper maggot, 251–253 Pepper weevil, 146–148 Peranabrus scabricollis, 576–579 Peregrinus maidis, 330–332 Peridroma saucia, 513–516 persicae, Myzus, 286–290 perspectalis, Hymenia, 410 Pest identification asparagus, 43–44 bean, 44–45 beet, 45 behavior, 23 cabbage, 45–46 carrot, 46–47 Cooperative Extension System, 24 county agents, 24 easy-to-discern characters, 23 lettuce, 47–48 noninsect groups, 39–41t okra, 48 onion, 48–49 rhubarb, 49 squash, 49–50 sweet corn, 50–51 sweet potato, 51 tomato, 51–52 vegetable-feeding bugs, 34–35t vegetable-feeding flies, 33–34t vegetable-feeding grasshoppers and crickets, 38t vegetable-feeding Hemiptera, 35–36t vegetable-feeding insects, 24–30t vegetable-feeding moths and butterflies, 36–37t phaseoli, Caliothrips, 581–600 Phenacoccus madeirensis, 371–373 Phenacoccus solenopsis, 369–371 phidilealis, Hellula, 398 Phthia picta, 323–326 Phthorimaea operculella, 439–442 Phyllophaga, 200–203 phyllopus, Leptoglossus, 323–326 Phyllotreta albionica, 64–65 Phyllotreta armoraciae, 73–74 Phyllotreta cruciferae, 67–69 Phyllotreta pusilla, 87–88 Phyllotreta ramosa, 81–82 Phyllotreta striolata, 81 Phyllotreta zimmermanni, 88–89 Pickleworm, 416–418 picta, Melanchra, 522–523 picta, Phthia, 323–326 Picture-winged flies, 9, 255–258 Pieridae alfalfa caterpillar, 525–527 characteristics, 13 imported cabbage worm, 527–530 mustard white, 530–531 southern cabbage worm, 531–532 southern white, 532–534 Pieris napi, 530–531

Pieris rapae, 527–530 Piezodorus guildinii, 360–361 Pillbugs, 613–615 Pink hibiscus mealybug, 373–375 pisorum, Bruchus, 94–96 pisum, Acyrthosiphon, 297 planipalpis, Delia, 231–232 Plantain looper, 494 Plant bug alfalfa, 333–335 carrot, 335–336 garden fleahopper, 336–338 pale legume bug, 339–343 rapid plant bug, 338–339 tarnished, 339 tomato, 343–345 western tarnished, 339 Plant family, vegetables, 651–658t Planthoppers, 11–12 platura, Delia, 232–234 Platyptilia carduidactyla, 538–539 Plutella xylostella, 534–538 Plutellidae, 534–538 Poaceae, 5–6 Polygonaceae, 5 Polyphagotarsonemus latus, 602–604 polyxenes, Papilio, 523 Pomace flies, 9, 238–240 Pontia protodice, 531–532 Popillia japonica, 191–194 populivenae, Pemphigus, 304–306 Porcellionidae, 613–615 Porcellio scaber, 613–615 postfasciatus, Euscepes, 158–160 Potato aphid, 300–303 Potato flea beetle, 76 Potato leafhopper, 320–322 Potato psyllid, 377–379 Potato stalk borer, 148–150 Potato stem borer, 494 Potato tuberworm, 439–442 praefica, Spodoptera, 520–522 Prairie grain wireworm, 165 pratensis, Oligonychus, 601–602 precationis, Autographa, 494 profundalis, Udea, 400–401 pronuba, Noctua, 518–519 propinquus, Melanoplus, 567–569 proteus, Urbanus, 445–447 protodice, Pontia, 531–532 pruininus, Selatsomus, 165 Pseudacteon tricuspis, 387 pseudobrassicae, Lipaphis, 306–308 Pseudococcidae characteristics, 11–12 cotton mealybug, 369–371 madeira mealybug, 371–373 pink hibiscus mealybug, 373–375 Psila rosae, 240–243 Psilidae, 9, 240–243 Psyllidae, 12, 377–379 Psyllids, 12 Psylliodes punctulata, 72–73 Pterophoridae, 538–539

Index  797

Puget sound wireworm, 165–167 pulicaria, Chaetocnema, 65 pulicaria, Corimelaena, 379–380 punctulata, Psylliodes, 72–73 Purplebacked cabbage worm, 418–419 pusilla, Phyllotreta, 87–88 Pycnoscelus indica, 57 Pycnoscelus surinamensis, 57 Pyemotes tritici, 387–388 Pyralidae lesser cornstalk borer, 539–542 limabean pod borer, 542–543 sod and root webworms, 419–421 Pyralid moths, 539–541 Pyrrharctia isabella, 429–431 Pyrrhocoridae cotton stainer, 375–376

Q

quadrilineatus, Macrosteles, 312–315 quadrisignatus, Glischrochilus, 183–184 quinquemaculata, Manduca, 547–550

R

radicum, Delia, 224–227 radiella, Depressaria, 392–393 Radish root maggot, 231–232 ramosa, Phyllotreta, 81–82 rantalis, Achyra, 409–410 rapae, Ceutorhynchus, 134–135 rapae, Pieris, 527–530 raphanus, Nysius, 332–333 Rapid plant bug, 338–339 rapidus, Adelphocoris, 338–339 recurvalis, Spoladea, 410 Redbacked cutworm, 496–498 Redbanded stink bug, 360–361 Redheaded flea beetle, 78 Red imported fire ant biological control, 387–388 cultural practices, 387 damage, 386–387 distribution, 384 host plants, 384 insecticides, 387 life cycle, 385 natural enemies, 384 sampling, 387 Redlegged grasshopper, 567 Redshouldered stink bug, 361–363 Red turnip beetle, 114–115 Reticulitermes flavipes, 61 Reticulitermes hageni, 59–62 Reticulitermes virginicus, 61 Rhinotermitidae damage, 62 management, 62 natural history, 59–62 Rhizoglyphus echinopus, 604–605 Rhizoglyphus robini, 604–605 Rhopalosiphum maidis, 279–281 Rhopalosiphum padi, 271–273 Rhopalosiphum rufiabdominalis, 303–304

Rhubarb, pest identification, 49 Rhubarb curculio, 150–151 ribisnigri, Nasonovia, 290–292 Rice root aphid, 303–304 richteri, Solenopsis, 384 rimosalis, Evergestis, 402–403 Ringlegged earwig damage, 209 distribution, 207 host plants, 208 life cycle, 208 management, 209 natural enemies, 208 robini, Rhizoglyphus, 604–605 rogatalis, Hellula, 398 Romalea microptera, 559–562 Root and seed maggots, 221–236 rosae, Psila, 240–243 Rose chafer, 196 rubens, Gryllus, 571–573 rubigalis, Udea, 400 rufiabdominalis, Rhopalosiphum, 303–304 rufimanus, Bruchus, 90–92 Rumina spp., 619–622 Rust flies, 9, 240–243

S

saccharalis, Diatraea, 387, 425–427 Saltmarsh caterpillar, 434–436 sanguinipes, Melanoplus, 562–567 Sap beetles, 9 sativae, Liriomyza, 219–221 saucia, Peridroma, 513–516 sayi, Chlorochroa, 363 Say stink bug, 363 scaber, Porcellio, 613–615 scabra, Hypena, 431–433 scabricollis, Peranabrus, 576–579 Scarabaeidae asiatic garden beetle, 184–186 carrot beetle, 186–187 characteristics, 9 chinese rose beetle, 187–188 green june beetle, 189–191 japanese beetle, 191–194 oriental beetle, 194–196 rose chafer, 196 spring rose beetle, 197–198 sugarcane beetle, 198–200 western rose chafer, 196–197 white grubs, 200–203 Scarab beetles, 9 Schistocerca americana, 555–557 schizoceri, Boethus, 383 Schizocerophaga leibyi, 383 schultzei, Frankliniella, 585–586 Scientific name, vegetables, 651–658t Scirtothrips dorsalis, 583–585 scorbutica, Anasa, 326–329 scutellatus, Orthops, 335–336 Scutigerella immaculata, 622–624 Scutigerellidae, 622–624

Seed bugs, 10 Seedcorn beetle, 63 Seedcorn maggot, 232–234 Selatosomus aeripennis, 165–167 Selatosomus destructor, 165 Selatsomus pruininus, 165 servus, Euschistus, 348–350 Sesiidae southwestern squash vine borer, 543–546 squash vine borer, 543 sexta, Manduca, 547 Shield-backed katydids, 576–579 Shortwinged mole cricket, 573 simplex, Anabrus, 576 simplex, Ophiomyia, 213–215 sinicus, Adoretus, 187–188 Sitona lineatus, 143–146 Skipper butterflies, 13 Slugs, 615–619 Small fruit flies, 238–240 Smartweed flea beetle, 78–79 Sminthuridae, 609–611 Smynthurodes betae, 271 Snails, 619–622 Snout moths, 13, 393–429 Sod and root webworms, 419–421 Solanaceae, 6 solani, Aulacorthum, 285–286 solani, Gargaphia, 380–382 solenopsae, Kneallhazia, 384, 387 Solenopsis daguerrei, 384–385 Solenopsis geminata, 384 Solenopsis invicta, 384–388 solenopsis, Phenacoccus, 369–371 Solenopsis richteri, 384 Solenopsis xyloni, 384 somnulentella, Bedellia, 391 Southeastern field cricket, 571–573 Southern armyworm, 498–500 Southern beet webworm, 410–413 Southern cabbage worm, 531–532 Southern corn billbug, 141–143 Southern cornstalk borer, 421–422 Southern cowpea seed beetle, 92–94 Southern cowpea weevil. See Southern cowpea seed beetle Southern green stink bug, 365–368 Southern mole cricket, 573 Southern potato wireworm, 167 Southern redlegged grasshopper, 567–569 Southern tobacco flea beetle, 83–85 Southern white, 532–534 Southwestern corn borer, 422–425 Southwestern squash vine borer, 543–546 Sowbugs, 613–615 Soybean looper, 500–502 Sphenophorus callosus, 141–143 Sphenophorus maidis, 141 Sphingidae characteristics, 13–14 sweetpotato hornworm, 546–547 tobacco hornworm, 547 tomato hornworm, 547–550 whitelined sphinx, 550–551

798  Index

Sphinx moths, 546–551 Spilosoma virginica, 436–438 Spinach flea beetle, 79 Spinach leafminer, 222–224 spinipennis, Bigonicheta, 205–206 Spodoptera dolichos, 508 Spodoptera eridania, 498–500 Spodoptera exigua, 56–57, 456–459 Spodoptera frugiperda, 482–486 Spodoptera latifascia, 508–510 Spodoptera ornithogalli, 520 Spodoptera praefica, 520–522 Spoladea recurvalis, 410 Spotted asparagus beetle, 115–116 Spotted beet webworm, 410 Spotted blister beetle, 178–179 Spotted cucumber beetle, 116–120 Spotted cutworm, 502–504 Spring field cricket, 571 Spring rose beetle, 197–198 Springtails, 609–611, 629f Squash, pest identification, 49–50 Squash and related crops, 5 Squash beetle, 132–134 Squash bugs, 10, 326–329 Squash vine borer, 543 Stalk borers, 13, 504–506, 641 Stenolophus comma, 63 Stenolophus leconti, 63–64 sticticalis, Loxostege, 395–397 stigmatias, Euxesta, 255–257 Stink bugs, 10–11, 632–634 Strawberry spider mite, 605 Striacosta albicosta, 516–518 strigatus, Eumerus, 243 Strigoderma arboricola, 197–198 striolata, Phyllotreta, 81 Striped blister beetle, 179–181 Striped cucumber beetle, 120 Striped flea beetle, 81 Striped grass looper, 506–508 Striped tortoise beetle, 97–98 Strymon melinus, 447–449 subcrinita, Epitrix, 76–78 subgothica, Feltia, 480–482 subspinosus, Macrodactylus, 196 subterranea, Agrotis, 487–489 Subterranean termites distribution, 59 host plants, 59 life cycle and description, 60 natural enemies, 59–60 Sugarbeet root aphids, 304–306 Sugarbeet root maggot, 257–258 Sugarbeet wireworm, 170 Sugar cane, 56 Sugarcane beetle, 198–200 Sugarcane borer, 425–427 Sulfur butterflies, 13, 525–534 Surinam cockroach, 57 surinamensis, Pycnoscelus, 57 suturellus, Dysdercus, 375–376 Swallowtail butterflies, 13, 523–525 Swede midge, 236–238

Sweet corn, pest identification, 50–51 Sweet potato, pest identification, 51 Sweetpotato armyworm, 508 Sweetpotato flea beetle, 82–83 Sweetpotato hornworm, 546–547 Sweetpotato leaf beetle, 122–123 Sweetpotato leaf folder, 427–428, 510–511 Sweetpotato leaf miner, 391–392 Sweetpotato sawfly damage, 384 distribution, 383 host plants, 383 life cycle, 383 management, 384 natural enemies, 383 Sweetpotato vine borer, 428–429 Sweetpotato weevil, 151–154 Sweetpotato whitefly, 262–266 Symphyla, 622–624 Syrphidae characteristics, 9–10 lesser bulb fly, 243–244 onion bulb fly, 243 Systena blanda, 74 Systena elongata, 74–76 Systena frontalis, 78 Systena hudsonias, 78–79

T

tabaci, Bemisia, 262–266 tabaci, Thrips, 591–595 Tarnished plant bug, 339 Tarsonemidae, 602–604 Tawny mole cricket, 573–576 Tenebrionidae characteristics, 9 false wireworms, 203–204 tenellus, Circulifer, 315–318 tenuis, Nesidiocoris, 343–345 Tephritidae mediterranean fruit fly, 244–247 melon fly, 247–249 oriental fruit fly, 249–250 parsnip leafminer, 251 pepper maggot, 251–253 Tetanops myopaeformis, 257–258 Tetranychidae banks grass mite, 601–602 strawberry spider mite, 605 tumid spider mite, 605–608 twospotted spider mite, 605 Tetranychus tumidus, 605–608 Tetranychus turkestani, 605 Tetranychus urticae, 605 Tettigoniidae coulee cricket, 576–579 mormon cricket, 576 texanus, Listronotus, 137–140 Texas carrot weevil, 137–140 Three-spotted flea beetle, 79 Thripidae american bean thrips, 581–600 bean thrips, 581–583

chilli thrips, 583–585 common blossom thrips, 585–586 Florida flower thrips, 586–588 grass thrips, 588–589 melon thrips, 589–591 onion thrips, 591–595 tobacco thrips, 595–597 western flower thrips, 597–600 Thrips immature, 630f narrow wings and fringe, 628f Thrips palmi, 589–591 Thrips tabaci, 591–595 Thyanta spp., 361–363 Thyrecoridae, 379–380 Thyreocoridae, 11 Thysanoptera american bean thrips, 581–600 bean thrips, 581–583 chilli thrips, 583–585 common blossom thrips, 585–586 Florida flower thrips, 586–588 grass thrips, 588–589 melon thrips, 589–591 onion thrips, 591–595 tobacco thrips, 595–597 western flower thrips, 597–600 Tiger moths, 12–13, 429–438 Tingidae characteristics, 11 eggplant lace bug, 380–382 Tipula oleracea, 253–255 Tipula paludosa, 253 Tipulidae cabbage crane fly, 253–255 european marsh crane fly, 253 Tobacco budworm, 511–513 Tobacco flea beetle, 83 Tobacco hornworm, 547 Tobacco thrips, 595–597 Tobacco wireworm, 167 Tomarus gibbosus, 186–187 Tomato, pest identification, 51–52 Tomato hornworm, 547–550 Tomato leaf miner, 442–443 Tomato pinworm, 443–445 Tomato plant bugs, 343–345 Tomato russet mite, 608–609 Tomato stink bug, 368–369 Toothed flea beetle, 65–67 Tortoise beetle argus, 97 blacklegged, 97 golden, 97 mottled, 97 striped, 97–98 Tortricidae characteristics, 14 pea moth, 551–553 Toxoptera citricida, 56–57 Trialeurodes vaporariorum, 259–262 triangularis, Disonycha, 79 Trichobaris trinonotata, 148–150 Trichoplusia ni, 467–471

Index  799

tricuspis, Pseudacteon, 387 trifolii, Discestra, 473–474 trifolii, Liriomyza, 211–213 trifurcata, Cerotoma, 103–105 trinonotata, Trichobaris, 148–150 tripunctata, Lygropia, 427–428, 510–511 tristis, Anasa, 326–329 tritici, Pyemotes, 387–388 trivittatum, Acalymma, 120–122 tuberculatus, Eumerus, 243–244 Tuber flea beetle, 85–87 tuberis, Epitrix, 85–87 Tumid spider mite, 605–608 tumidus, Tetranychus, 605–608 turkestani, Tetranychus, 605 Turnip aphid, 306–308 Turnip root maggot, 235–236 Tuta absoluta, 442–443 Twospotted spider mite, 605 Two-striped grasshopper, 569–571 Typophorus nigritus, 122–123

U

Udea profundalis, 400–401 Udea rubigalis, 400 uhleri, Chlorochroa, 363–365 Uhler stink bug, 363–365 Ulidiidae cornsilk flies, 255–257 characteristics, 9 sugarbeet root maggot, 257–258 Ulus spp., 203–204 Umbelliferae, 4 undalis, Hellula, 398–400 undecimpunctata, Diabrotica, 116–120 uniformis, Macrodactylus, 196–197 unipuncta, Mythimna, 452–455 Urbanus proteus, 445–447 urticae, Tetranychus, 605

V

vaporariorum, Trialeurodes, 259–262 Variegated cutworm, 513–516 variolarius, Euschistus, 356–357 varivestis, Epilachna, 129–132

Vegetable-feeding bugs, 34–35t butterflies, 36–37t crickets, 38t flies, 33–34t grasshoppers, 38t hemiptera, 35–36t insects, 24–28t moths, 36–37t Vegetable leafminer, 219–221 Vegetable plant names, 651–658t Vegetable weevil, 154–156 veletis, Gryllus, 571 Velvet armyworm, 508–510 vespertinus, Conoderus, 167 vicinus, Neoscapteriscus, 573–576 Vine borers, 543–546 virescens, Chloridea, 511–513 virgifera, Diabrotica, 123–127 virginica, Spilosoma, 436–438 viridula, Nezara, 365–368 vittata, Epicauta, 179–181 vittatum, Acalymma, 120 vulgare, Armadillidium, 613

W

Webworms, 13, 393–429 Weevils, 8 Western bean cutworm, 516–518 Western black flea beetle, 87–88 Western corn rootworm, 123–127 Western flower thrips, 597–600 Western potato flea beetle, 76–78 Western potato leafhopper, 322–323 Western rose chafer, 196–197 Western striped cucumber beetle, 120–122 Western striped flea beetle, 81–82 Western tarnished plant bug, 339 Western yellow-striped armyworm, 520–522 West Indian sugarcane rootstalk borer weevil, 156–158 West Indian sweetpotato weevil, 158–160 Wheat wireworm, 172–174 White butterflies, 13, 525–534 Whiteflies, 11, 628f, 630f Whitefringed beetle, 160–162

White grubs, 9, 200–203 Whitelined sphinx, 550–551 Willow-carrot aphid, 308–310 Winter cutworm, 518–519 Wireworm corn, 162 dryland, 165 eastern field, 170 great basin, 165 gulf, 167–170 oregon, 162–165 pacific coast, 170–172 prairie grain, 165 puget sound, 165–167 southern potato, 167 sugarbeet, 170 tobacco, 167 wheat, 172–174 Woodlice, 613–615 Woollybear caterpillars, 429–438 Wooly caterpillars, 12–13

X

xanthomelas, Disonycha, 79 Xestia c-nigrum, 502–504 Xestia dolosa, 502–504 xyloni, Solenopsis, 384 xylostella, Plutella, 534–538

Y

Yellowmargined leaf beetle, 127–129 Yellow-necked flea beetle, 80 Yellow-striped armyworm, 520 Yellow woollybear, 436–438

Z

Zachrysia spp., 619–622 zea, Helicoverpa, 474–479 Zebra caterpillar, 522–523 zelicaon, Papilio, 523–525 Zimmermann’s flea beetle, 88–89 zimmermanni, Phyllotreta, 88–89 zinckenella, Etiella, 542–543 zonatus, Leptoglossus, 323–326 Zonosemata electa, 251–253

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