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Vims-Insect-Plant Interactions
Vims-Insect-Plant Interactions
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Virus-Insect-Plant Interactions EDITED BY
Kerry R Harris Virus-Vector Research Center Department of Entomology Texas A&M University College Station, Texas
Oney P. Smith Department of Biology Hood College Frederick, Maryland
James E. Duffus Crop Improvement Research Unit USDA-ARS Salinas, California
ACADEMIC PRESS San Diego
London
Boston New York
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Toronto
This book is printed on acid-free paper. ® Copyright © 2001 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. Requests for permission to make copies of any part of the work should be mailed to: Permissions Department, Harcourt Inc., 6277 Sea Harbor Drive, Orlando, Florida 32887-6777 Academic Press a Harcourt Science and Technology Company 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http://www.academicpress.com Academic Press Harcourt Place, 32 Jamestown Road, London NWl 7BY, UK http://www.academicpress.com
Library of Congress Catalog Card Number: 2001093315
International Standard Book Number: 0-12-327681-0
PRINTED IN THE UNITED STATES OF AMERICA 01 02 03 04 05 06 SB 9 8 7 6 5 4
3 2
1
Contents
Contributors Preface
xi XV
Acknowledgments
xvii
PART I: VIRUS LOCALIZATION IN PLANTS AND VECTORS
1. Tomato Yellow Leaf Curi Virus: A Disease Sexually Transmitted by Whiteflies HENRYK CZOSNEK, SHAI MORIN, GALINA RUBINSTEIN, VIVIANE FRIDMAN, MUHAMAD ZEIDAN, AND MURAD GHANIM
I. II. III. IV V VI.
Introduction 1 Geminiviruses Transmitted by The Whitefly Bemisia tabaci 2 Role of Whitefly Endosymbiotic Chaperonins in Virus Transmission Deleterious Effects of Virus on Whiteflies 9 Sexual Transmission of Virus among Whiteflies 12 Concluding Remarks 18 References 23
4
2. Possible Etiology of Eriophyid Mite-Borne Pathogens Associated with Double Membrane-Bound Particles KYUNG-SOO KIM, KYUNG-KY AHN, ROSE C . GERGERICH, AND SUNG-BOON KIM
L Introduction 29 n. Groups ofEriophyid Mite-Associated Diseases III. Concluding Remarks 42 References 44
30
3. An Anatomical Perspective of Tospovirus Transmission T. NAGATA AND D, PETERS
L Introduction 51 II. Tospovirus Morphology and Composition
52
vi
CONTENTS
III. Thrips Vectors and Tospovirus Transmission IV Thrips as Tospvirus Hosts 56 V Concluding Remarks 63 References 63
52
PART II: ELUCIDATION OF TRANSMISSION MECHANISMS
4. Analysis of Circulative Transmission by Electrical Penetration Graphs W. FRED TJALLINGII AND ERNESTO PRADO
I. II. III. IV. V VI.
Introduction 69 The Electrical Penetration Graph Technique 70 Barley Yellow Dwarf Virus Transmission by Rhopalosiphon padi Studies of Other Circulative Viruses 80 Vector Resistance in Plants 81 Concluding Remarks 82 References 83
74
5. Analysis of Noncirculative Transmission by Electrical Penetration Graphs ALBERTO FERERES AND JOSE LUIS COLLAR
I. II. III. IV V
Introduction 87 Noncirculative Transmission: Properties and Vector Participants Electronic Analysis of Nonpersistent Transmission 90 Electronic Analysis of Semipersistent Transmission 100 Concluding Remarks 102 References 103
88
6. Ingestion-Egestion Theory of Cuticula-Bome Virus Transmission KERRY F. HARRIS AND LISA JEAN HARRIS
I. II. III. IV V VI. VII. VIII.
Introduction 111 Terminology 112 Mechanism of Nonpersistent Transmission Site of Virus Retention 117 Electrical Penetration Graph Analysis Role of Watery Saliva in Transmission Semipersistent Transmission 127 Concluding Remarks 129 References 129
115 120 125
7. Mechanism of Virus Transmission by Leaf-Feeding Beetles ROSE C. GERGERICH
I. Introduction
133
CONTENTS
II. III. IV V VI.
Vll
Vims Acquisition: Beetle-Plant Interactions Plant Virus-Beetle Interactions 134 Deposition of Virus in Beetle Regurgitant Virus-Host Plant Interactions 136 Concluding Remarks 140 References 140
133 135
PART III: MOLECULAR ASPECTS OF VIRUS-VECTOR INTERACTIONS
8. Caulimoviruses STEPHANE BLANC, EUGENIE HEBRARD, MARTIN DRUCKER, AND REMY FROISSART
I. II. III. IV V VI. VII. VIII.
Introduction 143 The Virus 145 Biology of Caulimovirus Transmission by Aphids Identification of Aphid Transmission Factor(s) Characterization of Aphid Transmission Factor(s) Mode of Action of Aphid Transmission Factor(s) Regulation of Aphid Transmission Factor's Function(s) Concluding Remarks 160 References 162
148 150 152 156 159
9. Cucumoviruses KEITH L. PERRY
I. II. III. IV V VI.
Introduction 167 Viral Genome and Cucumovirus Transmission Vector Transmission of Cucumber Mosaic Virus StractuTQ of Cucumber Mosaic Virus 170 Mechanisms of Aphid Transmission of Cucumoviruses Concluding Remarks 175 References 176
168 169 173
10. Potyviruses BENJAMIN RACCAH, HERVE HUET, AND STEPHANE BLANC
I. II. III. IV V VI. VII.
Introduction 181 Biology of Potyvims Transmission 182 Role of Coat Protein in Potyvirus Transmission Role of Helper Component 189 Potyvirus Transmission by Aphids 195 Specificity of Potyvirus Transmission by Aphids Concluding Remarks 198 References 200
185
197
viii
CONTENTS
11. Viral Determinants Involved in Luteovirus-Aphid Interactions VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND KENNETH E. RICHARDS
I. II. III. IV V
Introduction 207 Viral Passage through the Aphid 208 Identifying Viral Proteins Involved in Transmission Virus-Symbionin Interactions 225 Concluding Remarks 226 References 228
213
12. Approaches to Genetic Engineering of Potato for Resistance to Potato Leafroll Virus CHARLES R. BROWN AND ONEY P. SMITH
I. II. III. IV V
Introduction 233 The Virus 235 Approaches to Pathogen-Derived Resistance Resistance Mechanisms 240 Concluding Remarks 242 References 243
236
PART IV: ECOLOGY, EPIDEMIOLOGY, AND CONTROL
13. Bemisia: Pest Status, Economics, Biology, and Population Dynamics T. J. HENNEBERRY AND S. J. CASTLE
I. II. III. IV V
Introduction 247 Economic Impact and Pest Status Taxonomy Flux 250 Population Dynamics 252 Concluding Remarks 268 References 270
248
14. Whitefly-Bome Viruses in Continental Europe PiERO C. CACIAGLI
I. II. III. IV
Introduction 279 Virus and Virus-Like Diseases Vectors 285 Concluding Remarks 287 References 287
280
15. Transmission Properties of Whitefly-Bome Criniviruses and Their Impact on Virus Epidemiology GAIL C. WISLER AND JAMES E . BUFFUS
I. Introduction
293
CONTENTS
II. III. IV V VI. VII. VIII.
ix
Vector Transmission and Virus-Vector Relationships Criniviruses Infecting Cucurbits 296 Criniviruses Infecting Lettuce 298 Criniviruses Infecting Tomatoes 301 Criniviruses Infecting Sweet Potato 304 Criniviruses Infecting Weed Hosts 304 Concluding Remarks 305 References 306
294
16. Classical Biological Control of Bemisia and Successful Integration of Management Strategies in the United States A. A. KIRK, L. A. LACEY, AND J. A. GOOLSBY
I. II. III. IV V
Introduction 309 Foreign Exploration 311 Pathogenic Fungi for Biological Control of Silverleaf Whitefly Evaluation and Release of Silverleaf Whitefly Parasitoids Concluding Remarks 323 References 324
315 318
17. Interference with Ultraviolet Vision of Insects to Impede Insect Pests and Insect-Borne Plant Viruses YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
I. II. III. IV V
Introduction 331 StructureandFunctionof the Insect Compound Eye Ultraviolet-Dependent, Vision-Related Behavior Ultraviolet-Vision Based Management Strategies Concluding Remarks 347 References 347
332 334 336
18. Bionomics of Micrutalis malleifera Fowler and Its Transmission of Pseudo-Curly Top Virus JAMES H . TSAI
I. II. III. IV
Introduction 351 Biology of Pseudo-Curly Top Disease Vector Biology 353 Virus Transmission 360 References 361
Index
363
3 51
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Contributors
Numbers in parentheses indicate the pages on which the authors' contributions begin.
(29), Syngenta Seeds Co., Ltd., R&T Center, Changhowon-Eup, Ichon-Si, Kyunggi-Do, Korea.
KYUNG-KY AHN
(331), Agricultural Research Organization, The Volcani Center, Bet Dagan 50250, Israel.
YEHEZKEL ANTIGNUS
(143), Station de Recherches de Pathologic Comparee, Institut National de la Recherche Agronomique, Centre National de la Recherche Scientifique Unite Associee, 30380 Saint-Christol-les-Ales, France.
STEPHANE BLANC
VERONIQUE BRAULT
(207), Unite de Recherche Vigne et Vin, INRA, 68021 Colmar
Cedex, France. (233), USDA-ARS, Vegetable and Forage Crop Research Unit, Prosser, Washington 99350, USA.
CHARLES R. BROWN
PiERO C. CACIAGLI (279), Istituto di Fitovirologia Applicata, CNR, 10135 Torino, Italy. J. CASTLE (247), USDA-ARS, Western Cotton Research Laboratory, Phoenix, Arizona 85040-8803, USA.
STEVE
(331), Agricultural Research Organization, The Volcani Center, Bet Dagan 50250, Israel.
SHLOMO COHEN
JOSE LUIS COLLAR
(87), Aragonesas Agro S. A., 28004 Madrid, Spain.
(1), Department of Field Crops and Genetics, Faculty of Agricultural, Food and Environmental Quality Sciences, The Hebrew University of Jerusalem, Rehovot 76100, Israel.
HENRYK CZOSNEK
(143), Station de Recherches de Pathologic Comparee, Institut National de la Recherche Agronomique, Centre National de la Recherche Scientifique Unite Associee, 30380 Saint-Christol-les-Ales, France.
MARTIN DRUCKER
XI
xii
CONTRIBUTORS
(293), USDA-ARS, Crop Improvement Research Unit, Salinas, California 93905, USA.
JAMES E. DUFFUS
(87), Consejo Superior de Investigaciones Cientificas (CSIC), Centro de Ciencias Medioambientales, 28006 Madrid, Spain.
ALBERTO FERERES
ViviANE FRIDMAN (1), Institute of Life Science, The Hebrew University of Jerusalem, Rehovot 76100, Israel. (143), Station de Recherches de Pathologic Comparee, Institut National de la Recherche Agronomique, Centre National de la Recherche Scientifique Unite Associee, 30380 Saint-Christol-les-Ales, France.
REMY FROISSART
(29, 133), Department of Plant Pathology, University of Arkansas, Fayetteville, Arkansas 72701, USA.
ROSE C. GERGERICH
MuRAD GHANIM (1), Department of Field Crops and Genetics, Faculty of Agricultural, Food and Environmental Quality Sciences, The Hebrew University of Jerusalem, Rehovot 76100, Israel. A. GOOLSBY (309), USDA-ARS/CSIRO, Australian Biological Control Laboratory, Indooroopilly, Queensland, Australia 4068.
JOHN
(111), Virus-Vector Research Center, Department of Entomology, Texas A&M University, College Station, Texas 77843, USA.
KERRY F. HARRIS LISA JEAN HARRIS
(111), 4109 Viceroy Drive, Bryan, Texas 77802, USA.
(143), Station de Recherches de Pathologic Comparee, Institut National de la Recherche Agronomique, Centre National de la Recherche Scientifique Unite Associee, 30380 Saint-Christol-les-Ales, France.
EUGENIE HEBRARD
TOM J. HENNEBERRY (247), USDA-ARS, Western Cotton Research Laboratory, Phoenix, Arizona 85040, USA. HERVE HUET (181), Agricultural Research Organization, Department of Virology, The Volcani Center, Bet Dagan 50250, Israel. (29), Department of Plant Pathology, University of Arkansas, Fayetteville, Arkansas 72701, USA.
KYUNG-SOO KIM SUNG-BOON KIM
(29), 2405 Karyn Avenue, Fayetteville, Arizona 72703, USA.
ALAN A. KIRK (309), European Biological Control Laboratory, USDA-ARS, CILBA, Montferrier sur Lez, 34061 France. A. LACEY (309), USDA-ARS, Yakima Agricultural Research Laboratory, Wapata, Washington 98951, USA.
LAWRENCE
MosHE LAPIDOT (331), Agricultural Research Organization, The Volcani Center, Bet Dagan 50250, Israel. SHAI MORIN (1), Department of Plant Sciences, University of Arizona, Tucson, Arizona, USA.
CONTRIBUTORS
xiii
(51), Department of Cellular Biology, University of Brazil, Brazilia 70919-970, Brazil.
TATSUYA NAGATA
KEITH L. PERRY (167), Department of Botany and Plant Pathology, Purdue University, West Lafayette, Indiana 47907, USA. (51) Department of Virology, Wageningen Agricultural University, 6709 PD Wageningen, The Netherlands.
DICK PETERS
ERNESTO PRADO (69), Instituto de Investigaciones Agropecuarias, La Platina. Santiago, Chile. (181), Agricultural Research Organization, Department of Virology, The Volcani Center, Bet Dagan 50250, Israel.
BENJAMIN RACCAH
(207), Institut de Biologic Moleculaire des Plantes du CNRS et de TUniversite Louis Pasteur, 67084 Strasbourg Cedex, France.
KENNETH E. RICHARDS
(1), Department of Field Crops and Genetics, Faculty of Agricultural Food and Environmental Quality Sciences, The Hebrew University of Jerusalem, Rehovot 76100, Israel.
GALINA RUBINSTED^
(233), Department of Biology, Hood College, Frederick, Maryland 21701, USA.
ONEY P. SMITH
W.
(69), Department of Entomology, Wageningen University, Wageningen, The Netherlands.
FRED TJALLINGII
(351), University of Florida, Institute of Food and Agricultural Sciences (IFAS), Fort Lauderdale, Florida, USA.
JAMES H . TSAI
(293), Department of Plant Pathology, University of Florida, Gainesville, Florida 32611, USA.
GAIL C. WISLER
(207), Institut de Biologic Moleculaire des Plantes du CNRS et de TUniversite Louis Pasteur, 67084 Strasbourg Cedex, France.
VERONIQUE ZIEGLER-GRAFF
MuHAMAD ZEIDAN (1), Ministry of Agriculture, Plant Protection and Inspection Services, Bet-Dagan, Israel.
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Preface
Insect-transmitted plant viruses continue to plague man's attempts to grow plants for food, fiber, and other needs. The many biological and ecological factors that define these complex transmission systems are not readily apparent. In the past two decades, researchers have made strides in understanding virus-insect-plant interactions greater than the totality of knowledge gained during the previous 50 years. Much of this progress has resulted from the development and application of new technologies, as well as novel improvements in and applications of old ones. The greater one's understanding of the fundamentals of any pathogen-vectorhost transmission system, the greater the likelihood of one's discerning weak points in the system that might be manipulated to man's advantage. In the present volume, 42 highly respected researchers of virus-vector-plant interactions present and discuss the nature and implications of the most recent and significant breakthroughs. The information presented brings us to the brink of producing novel, user and environmentally friendly approaches to controlling insect-transmitted plant viruses. The book is comprised of 18 chapters organized under four subtitles: Virus Localization in Plants and Vectors; Elucidating Transmission Mechanisms; Molecular Aspects of Virus-Vector Interactions; and Ecology, Epidemiology, and Control. The primary topics treated in individual chapters and the pages on which each begins are listed in the Contents, to guide readers to topics of particular interest. Individual chapter bibliographies will help readers delve into topics with greater depth. This volume could serve as a usefril supplement in plant pathology, entomology, and other courses that treat insects as transmitters of pathogens. Kerry Harris, Senior Editor
XV
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Acknowledgments
In August of 1996, Jim Duffus and I traveled to the Rockefeller Foundation's Bellagio Study and Conference Center overlooking beautiful Lake Como in Bellagio, Italy to chair a week-long International Conference on "Whiteflies and Viruses: Menace to World Agriculture." The Center's staff, resident scholars, and picturesque environs inspired an abundant exchange of ideas among the conference participants. The latter exchange was to ignite the spark that would grow to be Virus-Insect-Plant Interactions, I speak for all when I thank the Rockefeller Foundation for their "birth-stage" involvement. Ms. Susan Garfield, Manager of the Bellagio Center Office in New York City, provided direction and help in planning the conference. Ms. Gianna Celli, Manager-in-Residence of the Study and Conference Center in Bellagio, looked after our every need, allowing us to devote full attention to the objectives and goals of the conference. Finally, we acknowledge and thank Dr. John J. McKelvey, Jr., retired Deputy Director of Agricultural Sciences for the Rockefeller Foundation, for serving as Conference Moderator. The editors thank the contributing authors and the staff of Academic Press for transforming Virus-Insect-Plant Interactions from concept to reality. I especially thank my wife, Lisa Jean Harris, and my co-editors, Oney Smith and Jim Duffus, for their support, help, patience, and perseverance. Kerry Harris, Senior Editor
xvu
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About the Editors
'^:'^'ii';'fii'i/&f > Kerry Francis Harris is Professor and Director of the Virus-Vector Research Center, Department of Entomology at Texas A & M University in College Station, Texas. He earned a Bachelor of Science degree at Louisiana State University in New Orleans and his Master of Science Degree in Biological Sciences at Loyola University in New Orleans where his master's dissertation on a genus of biting flies, Culicoides (Diptera: Ceratopogonidae) served as his introduction to virus-vector research. Twice awarded Fulbright fellowships, he also had accepted both National Institutes of Health and National Science Foundation fellowships for studies on virus-vector studies at Michigan State University where he earned his Ph.D. Dr. Harris has earned the respect of peers around the world as an innovative teacher, researcher, thinker, and scholar in virus-vector research. Having served on numerous national and international scientific panels and committees, as well as organizing and chairing over 30 international congresses, conferences, workshops, and symposia on pathogen-vector-host interactions, he has also authored over 80 journal publications, 16 book chapters, and 4 annual reviews. Former posts include Editor-in-Chief, Senior Editor, or Editorial Board Member of several book series and professional journals: Current Topics in Vector Research, Advances in Disease Vector Research, Plant Disease, and Journal of Economic Entomology. Dr. Harris' favorite occupation is to love and be loved by his wife Lisa Jean Patricia Harris, a registered nurse. In his spare time he can be found motorcycling or racing his Corvette - among his favorite hobbies.
XIX
XX
ABOUT THE EDITORS
Oney P. Smith is a Collaborating Scientist with USDAARS in Frederick, Maryland and an Assistant Professor of Biology at Hood College. Dr. Smith's education includes a Ph.D. in Entomology from Texas A & M University, a M.S. in Entomology from the University of Maine, and a B.S. in Biology from the University of Vermont. He is interested in the biologies, gene-expression strategies, and control of insect-transmitted plant viruses. His research experience includes a Postdoctoral Research Associateship with the Foreign Disease-Weed Science Research Unit of the USDA-ARS at Fort Detrick, Maryland where he conducted studies on the identification, molecular biology, and control of soybean dwarf and potato leafroll luteoviruses. Dr. Smith's research, supported by the National Potato Council, the USDA-ARS, and the undergraduate Summer Science Institute of Hood College, has been published in Archives of Virology, Biotechniques, Phytopathology, Plant Disease, and Virus Genes. Dr. Smith enjoys coaching youth basketball and trout fishing in western Maryland. Most importantly, he treasures his wife Marcia and their children, Andrew andAimee. James E. Duffus, Research Plant Pathologist with the USDA-ARS, US. Agricultural Research Station, Salinas, California recently retired after more then 42 years of service. A continuing Collaborating Scientist with the ARS, he has formerly served on various committees of the American Phytopathological Society, American Societies of Sugar Beet Technologists, the International Society for Plant Pathology, and the International Society of Horticultural Science. He earned a Ph.D. in Plant Pathology at the University of Wisconsin, with his B.S. earlier at Michigan State University. In recognition of his role in vector-insect-plant interactions he has been invited to present invitational papers for international meetings and organizations in 20 countries on beet virus diseases, virus-vector relationships, epidemiology, virus diseases of vegetable crops, and disease management practices. He resides in the Pastures of Heaven of the Monterey Bay region with his wife of about 50 years, Rachel. His long association with aphids and whiteflies has given him important insight on the importance of "Honey-do," which should take a good portion of his time.
CHAPTER 1
Tomato Yellow Leaf Curl Virus: A Disease Sexually Transmitted by Whiteflies HENRYK CZOSNEK SHAI MORIN GALINA RUBINSTEIN ViVIANE F R I D M A N MUHAMAD Z E I D A N
MURAD G H A N I M
/. Introduction Relationships between plant viruses and their arthropod vectors are complex and much more than passive associations [1]. Some plant viruses are carried in the insect feeding apparatus and can be acquired and inoculated within seconds or minutes (noncirculative transmission). Others circulate through the body of the insect and once acquired can be transmitted only after a latent or incubation period of hours to days (circulative transmission) [2, 3]. The passage of large amounts of plant virus in insect vectors is seldom innocuous. Several plant viruses replicate in their insect host (propagative circulative transmission) [4] and can be considered as plant as well as insect viruses [1]. The phytoarbovirus group contains members that replicate in their aphid or leafhopper vectors [1, 5], for example, replication of tomato spotted wilt tospovirus in thrips [6, 7]. A number of phytoarboviruses impair vector longevity and fecundity. Some are even transmitted from parent insect vector to progeny [8-10]. Horizontal venereal transmission of a plant virus among insects has never been reported. Transmission of virus through gametes of insects has been documented, however, especially for Drosophila spp. [11]. The Drosophila S virus (DSV), a reolike virus [12], invades differentiating male and female germ cells [13], causing developmental malformation [14]. The baculovirus-like gonad-specific virus (GSV) causes abnormalities in the male and female reproductive systems of two Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
\
2
HENRYK CZOSNEK ET AL.
closely related moth species, Helicoverpazae and H. armigera [ 15]. In both cases, the sperm and the egg could carry virions that could be transmitted to progeny at the time of fertilization. It is generally believed that geminiviruses do not affect their insect vectors [16]. Nonetheless, the relationship between the whitefly Bemisia tabaci and an isolate of tomato yellow leaf curl geminivirus from Israel (TYLCV-Is) is reminiscent of an insect-pathogen relationship [17]. In this review, we will present and discuss some of our recent investigations on whether TYLCV is a sexually transmitted pathogen ofB. tabaci.
IL Geminiviruses Transmitted by ttie Wliitefiy Bemisia tabaci A.
Geminiviruses in the Family Geminiviridae
Geminiviruses have a single-stranded DNA (ssDNA) genome encapsidated in about 20 X 30-nm geminate particles [18, 19]. Geminiviridae comprise three subgroups. Mastreviruses and curtoviruses possess a single genomic component of about 2,600 to 2,900 nucleotides (nt) encoding five to seven genes. They are transmitted by several species of leafhoppers. One mastrevirus is transmitted by a treehopper. Begomoviruses possess either one (monopartite) or two (bipartite) genomic components and are transmitted by a single whitefly species, B. tabaci. Monopartite begomoviruses have a genome of about 2,800 nt encoding six genes. The genome of bipartite begomoviruses is split between two DNA molecules of about 2,600 nt each: DNA A (basically similar to the genome of monopartite begomoviruses) and DNA B encoding two genes [20, 21]. Geminiviruses replicate in host plants via double-stranded DNA (dsDNA) intermediates and assemble in the nuclei of host plant cells [22]. In monopartite geminiviruses, the DNA A-like genomic molecule encodes the information required for replication, gene expression, particle assembly, and virus spread. In bipartite geminiviruses, the functions for virus spread are encoded by DNA B. B.
Whitefly-Transmitted Geminiviruses
Begomoviruses are all transmitted by the whitefly B. tabaci. The past two decades have witnessed a dramatic expansion of this insect from tropical and subtropical regions to more temperate ones [23-25]. Probably owing to human activities, some B. tabaci biotypes and variants have invaded new regions, whereas others have been almost eradicated. As a striking example, the B biotype of B. tabaci probably was introduced to the New World from the Middle East during the last
1.
TOMATO YELLOW LEAF CURL VIRUS
3
decade, displacing the endogenous A biotype [25, 26]. It has been proposed that the B biotype constitutes a new species, Bemisia argentifolii [27]. The spread of B. tabaci has been accompanied by a formidable increase in the economical importance of begomoviruses, which today infect many important agricultural plants worldwide, including bean, cassava, cotton, melon, pepper, potato, squash, tobacco, tomato, and watermelon. Sequence comparisons of begomovirus genomes and open reading frames (ORFs) has allowed grouping of these viruses according to their geographical origin: (1) New World, with subgroups including Central and South America and the Caribbean Islands (except the newly introduced Middle Eastern TYLCV); (2) western Mediterranean basin; (3) Middle East; (4) Indian subcontinent; and (5) East and Southeast Asia and Australia [20, 21, 28]. Similarly, the B. tabaci complex can be resolved into five major groups based on mitochondrial DNA markers; these groups essentially overlap the geographical groupings of begomoviruses [29]. The begomovirus capsid plays a crucial role in transmission by B. tabaci. Begomoviral particles are antigenically related, and the amino acid sequence of their coat protein (CP) is highly conserved. Swapping the CP between whiteflyand leafhopper-transmitted geminiviruses resulted in a swapping of vectors [30]. Similarly replacing the CP from a nontransmissible begomovirus with that from a transmissible one restores transmission by B. tabaci [31]. A single mutation resulting in the replacement of one amino acid in the CP of a TYLCV isolate from Sardinia, Italy (TYLCV-Sar) abolished the isolate's vector transmissibility but not its ability to assemble into virions and to replicate and spread in plants [32]. C.
Tomato Yellow Leaf Curl Virus
Tomato yellow leaf curl virus was one of the first begomoviruses characterized according to its relationship with the B. tabaci vector and its host range [33, 34]. It is also one of the most economically important begomoviruses worldwide [35]. Molecular comparisons of isolates from distinct geographical regions have revealed that the name TYLCV was given to closely related as well as to distantly related tomato begomoviruses [35-37]. The genome of TYLCV is either monopartite (Mediterranean isolates) or bipartite (Thailand isolate). The role of virus ORFs has been established by investigating TYLCV-Sar. Similarly to other geminiviruses, monopartite TYLCV isolates encode six genes, two on the virion (+) strand (VI and V2) and four on the complementary (-) strand (C1-C4). The VI gene encodes the CP, and V2 may control symptom expression and viral movement. The CI gene encodes the Rep protein necessary for virus replication, C2 encodes a transcription activator protein, C3 encodes a replication enhancer protein, and C4 may affect host range, symptom severity, and movement [3 8-41 ].
4
HENRYK CZOSNEK ET AL.
///. Role of Whitefly Endosymbiotic Chaperonins in Virus Transmission A.
Virus Acquisition and Transmission
Whitefly-mediated transmission to tomato test plants and subsequent observation of induced disease symptoms indicated that the minimal acquisition access periods (AAPs) and inoculation feeding periods (IFPs) for TYLCV-Is transmission were approximately 15 minutes [33, 34]. Similar values were obtained with TYLCV isolates from other regions. In a one insect-one plant inoculation assay, female B. tabaci transmitted TYLCV-Is with higher efficiency than did males [34]. By applying polymerase chain reaction (PCR) technology, one can detect amounts of TYLCV DNA in single insects that are below the threshold of infectivity [42]. By use of the print-capture PCR technique, TYLCV-Is DNA was found in 20% of individual whiteflies tested after AAPs as brief as 5 minutes and in all insects given a 10-minute AAP. With the same technique, we have determined that TYLCV-Is can be transmitted to about 15% of tomato test plants after a 5-minute IFP and to all of them after a 30-minute one [43]. The minimal AAP and IFP obtained by PCR are much briefer than the 15-minute ones established previously by examining stylet penetration under the light microscope [44], by whitefly-mediated transmission [34], or by analyzing electronic waveforms produced during whitefly feeding [45]. B.
Circulative Transmission
Since geminiviruses are transmitted in a circulative manner, whiteflies are unable to inoculate begomoviruses immediately after acquiring them. Geminivims particles ingested with phloem sap through the stylets into the gut are subsequently translocated through the gut wall into the hemocoel, where they are carried by hemolymph to the salivary glands. Virions that selectively traverse salivary gland cells to the salivary duct system are inoculated to plants in virus-laden saliva during feeding [46-48]. The latent period of TYLCV-Is is 8 to 10 hours, measured as the time interval from the start of an AAP to the whitefly's ability to inoculate virus to a test plant [34]. The pol)mierase chain reaction has been instrumental in tracking geminivirus translocation over time in B. tabaci. Squash leaf curl virus (SLCV) is detectable in insect extracts 30 minutes after the beginning of an AAP, in the hemolymph after 2 hours, and in the saliva and honeydew after 8 hours [49]. We followed the translocation of TYLCV-Is in B. tabaci using stylets, midgut, hemolymph, and salivary glands dissected from individual insects as PCR templates [50]. Virus was detected in the insect head 10 minutes after the beginning of an AAP, in the midgut after 60 minutes, and in the hemolymph after 90 minutes. TYLCV was detected in the salivary glands approximately 5 V2 hours after it was first detected
1.
TOMATO YELLOW LEAP CURL VIRUS
5-10 min
5-10 min
1.5 h
4
Salivary glands Test Plant
Fig, 1 Schematic representation of the circulative pathway of an Israel strain of tomato yellow leaf curl vims (TYLCV-Is) and the velocity of translocation of the virus in selected organs of Bemisia tabaci. The values were obtained by amplifying TYLCV-Is DNA using, as PCR templates, insect and plant squashes [43] and isolated insect organs [50].
in the hemolymph or 7 hours after the start of an AAP (Fig. 1). Since the latent period is approximately 8 hours, it seems that once the virus reaches and penetrates the salivary glands, it is almost immediately ejected by the salivary pump into the target plant. C. Involvement of a GroEL Homologue from Vector Endosymbionts in Transmission Once ingested, viral particles movefi*omthe digestive tract lumen through the gut wall to the hemolymph. This transit is a long (Fig. 1) and hazardous phase in circulative transmission. The question of how viruses are protected in the hemolymph is fundamental to our understanding of how circulative transmission
6
HENRYK CZOSNEK ET AL.
is ensured. The role of chaperonins synthesized by insect endosymbiotic bacteria in mediating survival of viruses was first demonstrated in aphids. Potato leaf roll luteovirus (PLRV) survival depends on a 63-kDa GroEL homologue [51] produced by the primary endosymbiont, a Buchnera sp., of the aphid Myzus persicae [52]. The symbionts are harbored in the hemocoel in specialized polyploid cells called mycetocytes [53]. Buchnera GroEL homologues are very similar to GroEL of Escherichia coli and other free-living bacteria with which they share putative functional domains [54-56]. Buchnera GroEL is not restricted to symbiont cytosol and is present in aphid hemolymph [52, 55, 57]. Inhibition of prokaryotic protein synthesis following antibiotic treatment of aphid nymphs results in decreased levels of GroEL in the hemolymph and destruction of PLRV As a result of the latter, transmission of PLRV is markedly inhibited [52]. Collectively, these and other experimental data indicate that some kind of interaction between luteovirus and endosymbiotic GroEL is essential for virus retention in the aphid vector (see Chapter 11). The survival of geminiviruses in the hemolymph ofB. tabaci is ensured by a similar strategy. Whiteflies, like aphids and most other homopterous insects, contain endosymbiotic microorganisms in their mycetocytes. Two morphologically distinct types of microorganisms are present in each mycetocyte. The predominant endosymbiont in B. tabaci biotype B [58] is highly pleomorphic (P type) and unrelated to the aphid primary endosymbiont [59]. The second type of B. tabaci endosymbiont, a coccoid bacterium (C type) found in lower numbers than the P type, morphologically resembles the aphid primary endosymbiont [59, 60]. We have immunolocalized GroEL in the cytoplasm of whitefly coccoid bacterium [61]. This protein is conspicuous in the insect hemolymph as a native tetradecamer unit, each subunit having a mass of 63 kDa (Fig. 2). No GroEL is detectable in the digestive tract. The approximately 63-kDa GroEL homologue of B. tabaci is a member of the chaperonin-60 family [62]. Particles of TYLCV displayed affinity for the B. tabaci GroEL homologue in a virus overlay assay. The biological significance of the TYLCV-GroEL interaction in the hemolymph was demonstrated by membrane feeding experiments. Whiteflies were fed through membranes with an antiserum to aphid Buchnera GroEL. The insects were fed anti-GroEL antiserum either before, during, or before and during membrane feeding on a suspension of partially purified TYLCV in sucrose solution. Control groups of whiteflies were prefed on preimmune serum prior to TYLCV acquisition. Antibody-treated and control insects were caged with tomato test plants for an inoculation feeding period of 5 days. Feeding whiteflies with anti-GroEL antiserum reduced TYLCV transmission to tomato test plants by more than 80%. The hemolymph concentrations of TYLCV DNA in anti-GroEL-treated whiteflies were reduced to levels undetectable by Southern blot hybridization, whereas viral DNA was readily detected in the hemolymph of insects fed preimmune serum. Active GroEL antibodies were recovered from the hemolymph of whiteflies fed anti-GroEL
1.
TOMATO YELLOW LEAF CURL VIRUS
W H
W H kDa • 83 • 62 • 47.5
Non-denaturing
SDS-PAGE
Fig. 2 Presence of a GroEL homologue in Bemisia tabaci. Protein extracts of whole whiteflies (W) and their hemolymph (H) were subjected to electrophoresis in nondenaturing polyacrylamide gels and in denaturing SDS-containing gels. The proteins were blotted and immunodetected by using an antibody against Buchnera GroEL from aphids [61].
antiserum. These results suggest that GroEL antibody altered the interaction between TYLCV and GroEL homologue in the hemolymph, leading to TYLCV degradation and markedly reduced virus transmission. D.
Interaction between Vector GroEL and Viral Coat Protein
In both aphids and whiteflies, it is postulated that interaction between GroEL and virions mediates the safe translocation of virions in the hemocoel. In order to investigate this hypothesis, interaction between TYLCV-Is CP and insect GroEL was studied in the yeast two-hybrid system [63]. A prerequisite for this work was cloning the GroEL gene from B. tabaci. Native GroEL was partially purified from B. tabaci homogenates, and the 63-kDa subunit was isolated from a SDS-PAGE gel. The sequence of the 30 Nterminal amino acids [61] was then used to design oligonucleotides specific to the 5' end of the GroEL gene coding sequence. To achieve PCR-aided progress
8
HENRYK CZOSNEK ET AL.
toward the 3' end of the gene, additional oligonucleotides were designed based on GroEL conserved regions from endosymbiotic as well as free bacteria [54, 56]. Four consecutive PCR amplification sequencing steps were necessary to obtain the sequence of the full-length structural gene. Two additional reactions were necessary to obtain the 5' and the 3' flanking regions of the gene. The full-length GroEL gene was amplified by using oligonucleotides covering the 5' and 3' untranslated regions. Sequence analysis of the B. tabaci GroEL gene revealed that it encodes 555 amino acids [64]. The aphid GroEL N-terminal (amino acids 1-121) and C-terminal (amino acids 409-474) regions of the equatorial domain that bind luteoviruses [55, 56] are homologous with the corresponding regions in the B. tabaci protein [64]. Therefore, it is likely that the equatorial domain of 5. tabaci GroEL is also involved in binding TYLCV The TYLCV-Is CP and the B. tabaci GroEL ORFs were cloned in yeast cells (Fig. 3). The CP structural gene was fused in-frame to the LexA binding domain (BD) in the yeast expression vector pLexABD. The GroEL gene was fused inframe to the B42 transcriptional activation domain (AD) in the yeast expression vector pB42AD, Both the GroEL plasmid (pB42AD-GroEL) and the CP plasmid (pLexABD-CP) were then co-introduced into yeast cells containing the LacZ reporter plasmid pAop-LacZ. The expression of GroEL and CP in the transformed yeast cells was assessed by Western blot analysis using antiserum to LexA (for the LexA-CP fusion) and to GroEL. The plasmids were maintained in yeast using a medium lacking uracyl, tryptophan, and histidine. Yeast co-transformed with pB42AD-GroEL and pLexABD-CP were able to grow vigorously in either the presence or absence of leucine (Fig. 3). In order to verify that only association of LexA BD and B42 AD (as a result of interaction between CP and GroEL) induces the expression of the LEU2 and LacZ promoters, the GroEL or CP coding sequences or both, were deleted from their vectors. Yeast transformed with plasmids that did not contain either the CP or the GroEL ORFs did not grow in the absence of leucine. All yeast colonies that grew in the absence of leucine stained blue in a colony-lift assay for p-galactosidase activity (Fig. 3, see color insert). These results indicated that, at least in this cell system, TYLCV CP and B. tabaci GroEL interact physically. It remains to be seen whether the virus CP-insect GroEL interaction conspicuous in yeast cells also occurs in insect hemolymph. We do not know whether the monomer-to-monomer interaction in yeast reflects a physical interaction in the vector's hemolymph between virus capsid and GroEL tetradecamer or whether this interaction is the key step to successful virus passage through the hemocoel. Impairment of CP-GroEL interaction leading to a destruction of virus in the hemolymph might explain how geminivirus mutants such as Abutilon mosaic virus [65] and TYLCV-Sar [32] lose their whitefly transmissibility and why the whitefly Trialeurodes vaporariorum can acquire but not transmit geminiviruses [49, 66, 67].
LexA„.
LacZ
TRPl
GroEI
AD
1
i
GroEL
GroEL
CP TYLCV
CP TYLCV
BD
Expression of LEU2 LacZ/CP/GroEL
B0
LEU2
AD
LacZ
Expression of |3-galactosidase LacZ/CP/GroEL
LacZ / GroEL ]
LacZ/CP
LacZ/CP/GroEL Fig, 3 Binding of Israel strain of tomato leaf curl virus (TYLCV-Is) coat protein (CP) to Bemisia tabaci GroEL in the yeast two-hybrid system. The GroEL structural gene was fused in-frame to the B42 transcriptional activation domain (AD) in the yeast expression vector pB42AD. The virus CP gene was fused in-frame to the LexA binding domain (BD) in the yeast expression vector pLexABD. The two plasmids were co-introduced into yeast cells containing the LacZ reporter gene in plasmid pAop-LacZ. Each plasmid had a nutritional selective marker gene, TRPl (tryptophan), HIS3 (histidine), and URA3 (uracyl). As a result of interaction between CP and GroEL, the AD and BD became functional, bound to the LEU2 and LacZ promoter and induced the expression of these genes. The yeast cells containing the three plasmids grew in the absence of leucine and stained blue in a colony-lift assay for p-galactosidase activity. Yeast cells with only the GroEL plasmid or the CP plasmid did not grow in the absence of leucine and did not stain blue in the Pgalactosidase assay.
This Page Intentionally Left Blank
1.
TOMATO YELLOW LEAF CURL VIRUS
' ' 1 ' 22 25
2 8 ^
Time after acquisition (days) Fig, 4 Life-long association of Israel strain of tomato leaf curl vims (TYLCV-Is) Bemisia tabacL Females that emerged during a 24-hour period were caged with infected tomato plants for 48 hours. Insects were then reared on eggplant, a TYLCV nonhost. Groups of whiteflies were collected at the time indicated. From each group, 10 insects were analyzed for TYLCV DNA by hybridization with a TYLCV-Is DNA probe (DNA), 10 others were analyzed for TYLCV coat protein (CP) by Western blot using an antibody raised against TYLCV virions, and 20 more were caged with tomato test plants for a 48-hour inoculation feeding period (one insect per plant) to test for TYLCV transmission. The results are presented as percentage of initial values. Whereas viral DNA was associated with the insects during their entire adult life, TYLCV CP was no longer detectable alfter 2 weeks. The ability of test whiteflies to transmit the virus decreased progressively with time but did not disappear.
iV. Deleterious Effects of Virus on Whiteflies A. Whitefly Long-term Retention of Viral DNA and Coat Protein During feeding on infected tomato plants, whiteflies accumulate large amounts of TYLCV The number of virions plateaus at about 600 million per whitefly after about 12 hours of feeding [6S]. Similar numbers have been reported for TYLCVSar [69] and for SLCV [67]. These findings suggest that B. tabaci possesses a saturable number of begomovirus receptors. If all virions follow a circulative path through the vector, virus acquired during a 24-hour AAP on TYLCV-infected tomato ought to vacate the insect's body after 24 hours of feeding on a nonhost plant. This, however, is not the case. It appears that only a small fraction of the acquired virions follow the complete circulative transmission pathway. Whereas some of the virus is excreted in honeydew [49], most remains associated with a whitefly for much longer than the approximate 8 hours needed for acquired virus to be transmitted. We researched the fate of viral DNA and CP in 1-day-old female whiteflies after a 48-hour AAP on TYLCV-infected tomato and subsequent rearing on a TYLCV nonhost, eggplant (Fig. 4). Viral DNA was conspicuous during the insect's 4-week-long adult life. On the other han4 CP was no longer detectable
10
HENRYK CZOSNEK ET AL.
Summer
f 1 a 0
Urn gI 10 15 20 25 30 35 40
Winter
10 15 20 25 30 35 40 Time after acquisition (days) Fig, 5 High mortality rates of viruliferous whiteflies. Females that emerged during a 24-hour period were caged with tomato yellow leaf curl virus (TYLCV)-infected tomato plants for 48 hours. Insects were then reared on eggplant (20 insects per plant). Mortality rates of viruliferous insects (dark gray bars), expressed as the percentage of the insect population that died within a 24-hour period, were compared with those of nonviruliferous control populations of the same age reared under similar conditions (light gray bars). The experiments were performed during August-September 1996 and January-February 1997. Viruliferous whiteflies started to die earlier and did so at a higher rate than nonviruliferous controls.
approximately 14 days post-AAP. The ability of the aging insects to inoculate tomato plants with virus acquired earlier steadily decreased from 100% to 10-20% [17]. Another study revealed that TYLCV-Sar DNA remained detectable in whiteflies for up to 18 days post-AAP, much longer than suggested by insect inoculativity but not for the entire adult life [69]. B.
Effect of Tomato Leaf Curl Virus on Whitefly Longevity and Fecundity
We assessed the effect of long-term association of TYLCV with B. tabaci on whitefly longevity (Fig. 5). Whitefly females, 1 day postemergence, were caged with TYLCV-infected tomato plants for a 48-hour AAP and subsequently reared
1.
11
TOMATO YELLOW LEAF CURL VIRUS
40-
38.1
36.5 32.8
C/3
IDS) 30
o
-
28.0
Wi i^K'i
28.0
22.7 12.9
§
100Eggplant
Tomato
3 days-old insects
Eggplant
Tomato
11 days-old insects
Fig. 6 Decrease in fertility of viruliferous whiteflies. Female whiteflies that emerged during a 24hour period were divided into groups. The first group was caged with tomato yellow leaf curl virus (TYLCV)-infected tomato plants for a single 48-hour acquisition access feeding period (AAFP) and divided into two subgroups: the first subgroup was reared on eggplant plantlets, the second on tomato seedlings (one insect per plant). The second group of whiteflies was reared on eggplants for 8 days, caged with TYLCV-infected tomato plants for a single 48-hour AAFP, and divided into two subgroups as described above. The number of eggs laid during a period of 7 days by 20 3-day-old and 20 11-dayold insects was counted (dark gray bars) and compared with that laid by nonviruliferous insects of the same age (light gray bars). The mean number of eggs laid by viruliferous whiteflies was significantly lower than that laid by nonviruliferous insects, whether reared on eggplant or tomato.
on eggplant. The mortality rates of viruliferous populations were compared with those of same-age nonviruliferous populations reared under similar conditions, but only on eggplant. In both the summer and winter seasons of 1996-1997, viruliferous insects started to die earlier and at higher rates (up to day 30) than nonviruliferous controls. At the population level, the difference between viruliferous and nonviruliferous population life expectancies at the 50% mortality point was between 5 and 7 days [17]. We also investigated the effect of long-term whitefly-TYLCV association on the fertility of viruliferous insects, as measured by the mean number of eggs laid by single B. tabaci females. The number of eggs laid on eggplant and tomato by 3and 11-day-old viruliferous whiteflies over a 7-day period was compared with that laid by nonviruliferous insects of the same age reared under similar conditions (Fig. 6). The mean number of eggs laid by 3-day-old viruliferous whiteflies was significantly lower than that laid by nonviruliferous controls (22.7 versus 38.1 on eggplant; 28.0 versus 36.5 on tomato). Similar results were obtained with
12
HENRYK CZOSNEK ET AL.
11-day-old insects. The host plant, whether eggplant or tomato, did not significantly affect insect fecundity [17].
U A.
Sexual Transmission of Virus among Whitefiies
Tomato Yellow Leaf Curl Virus Transmission between Sexes
Because TYLCV-Is had several characteristics of an insect virus, we researched the possibility of horizontal virus transmission, from insect to insect, without the mediation of a TYLCV-infected plant. Transmission between sexes was tested by caging 20 viruliferous males with 20 nonviruliferous females. The insects were fed through a membrane on a 15% sucrose solution. Courtship and copulation were observed through the translucent membrane with a binocular stereoscope. After 48 hour on the artificial diet, the DNA from all insects was subjected to PCR using TYLCV-specific primers [42]. Viral DNA was amplified from all the viruliferous males, confirming the reliability of the PCR detection method. Surprisingly, viral DNA was also amplified from 10 of 18 surviving females (Fig. 7A). A similar experiment was conducted with 20 viruliferous females and 20 nonviruliferous males. In this case viral DNA was detected in 5 of the 18 surviving males (Fig. 7B). In identical experiments, CP of TYLCV was detected by immunocapture PCR in similar numbers of test whitefiies. In all the cases studied, there was no significant linear correlation between the length of the caging period (4, 8, 24, or 48 hours) and the rates of TYLCV transmission between sexes. These results indicate that TYLCV is transmitted from viruliferous males to nonviruliferous females and vice versa, likely in the form of encapsidated virions [50, 89]. The question of whether TYLCV is transmissible among whitefiies of the same sex was also investigated. Twenty viruliferous females and twenty nonviruliferous females were placed together in a membrane-feeding cage. In a similar experiment, 20 viruliferous males were mixed with 20 nonviruliferous ones. Viruliferous insects were marked with a tiny blue dot on the dorsum of the thorax. After 48 hours, all whitefiies (alive and dead) were analyzed by PCR. In either case (all males or all females), viral DNA was detected in all 20 of the dotted viruliferous whitefiies but in none of the nonviruliferous insects. Therefore, TYLCV is not transmissible between insects of the same sex (Fig. 8). These experiments also show that nonviruliferous whitefiies can not acquire TYLCV from feeding medium contaminated by viruliferous insects. To confirm that TYLCV is transmitted during sexual contact between partners, ten opposite-sex couples (one viruliferous male and one nonviruliferous female) were enclosed in ten separate cages. After 24 hours, the presence of viral DNA was assessed by PCR. Six of the ten females contained TYLCV DNA. In the reciprocal experiment, three of the ten males caged with viruliferous females contained TYLCV DNA. When similar experiments were conducted with five pairs
1.
13
TOMATO YELLOW LEAF CURL VIRUS
F caged with M* P M* 1 2 3 4
5
6 7 8 9 10 11 1213 1 4 15 16 17 18
M caged with F'' 1
2
3 4
5 6 7 8
9
10 11 12 13 14 15 16 17 18
Fig. 7 Transmission of an Israel strain of tomato leaf curl virus (TYLCV-Is) from viruliferous Bemisia tabaci males to females and from viruliferous females to males. Twenty viruliferous males were caged with twenty nonviruliferous females and vice versa. After 48 hours, the DNA prepared from each surviving insect was subjected to PCR by using TYLCV-Is DNA-specific primers [71]. The products were subjected to agarose gel electrophoresis and stained. (A) Analysis of surviving 18 females (1 to 18) caged with viruliferous males. (B) Analysis of surviving 18 males caged with viruliferous females. (P), plasmid pTYH20.7 (contains a dimeric copy of the TYLCV-Is genome [87]; (M*) and (F*), viruliferous male and female, respectively; (M) and (F), nonviruliferous male and female. (Arrow) ~410 bp amplified viral DNA fragment. Nine females (A) and five males (B) acquired DNA while caged with viruliferous insects of the other sex.
of males and five pairs of females (one of each same-sex pair viruliferous and the other not), none of the male or female nonviruliferous partners contained detectable viral DNA. An analysis of data from these experiments indicates that TYLCV-Is can only be transmitted between whiteflies by contact between opposite-sex partners, probably during copulation. B.
Virus Is Serially Transmissible among Sexual Partners
If TYLCV-Is causes disease in both whiteflies and plants, then it should be transmissible from infected donor (vector) whitefly to noninfected recipient whitefly, which in turn becomes an infected vector that transmits in-line to another healthy recipient, and so on. We, therefore, designed an experiment to test for such serial transmission by placing 100 viruliferous males and 100 nonviruliferous females together in a membrane-feeding cage. After 24 hours on the artificial diet, 60 of the surviving females were collected and caged with 100 nonviruliferous males for an additional 24-hour feeding period and then analyzed for TYLCV DNA. Of the surviving males, 60 individuals were collected and caged with 100 nonviruliferous females for an additional 24-hour feeding period and then analyzed for viral
14
HENRYK CZOSNEK ET AL.
Non-viruliferous females (F)
Viruliferousfemales (F*) 12 3 4 5 6 7
9 10 11 12 13 14 15 16 17 18 19 20
A Non-viruliferous males (M)
M* 1 2
3
2 3 4 5
>l
7 8
9 10 1112 13 14
15 16 17 18 19
Viruliferous males (M*)
^1 F* ]
B
6
2 3 4 5 6
7 8 9 10 11 12 13 14 15 16 17 18 19
Fig. 8 No transmission of an Israel strain of tomato leaf curl virus (TYLCV-Is) among whiteflies of the same sex. Viruliferous insects were marked with a tiny blue dot on the dorsal side of the thorax. (A) Twenty nonviruliferous females (F) were caged with twenty viruliferous females (F*). (B) Nineteen nonviruliferous males (M) were caged with nineteen viruliferous males (M*). After 48 hours, the DNA prepared from each insect was subjected to PCR using TYLCV-Is DNA-specific primers. The products were subjected to agarose gel electrophoresis and stained. (Arrow) ~410 bp amplified viral DNA fragment. Although we could amplify TYLCV DNA from all the viruliferous Bemisia tabaci, none of the nonviruliferous whiteflies acquired detectable viral DNA from insects of the same sex.
DNA. The experiment was continued in this manner for four passages, starting from the initial viruHferous males. A reciprocal experiment was conducted in a similar manner, starting with 100 viruliferous females caged with 100 nonviruliferous males, etc. The results, summarized in Fig. 9, show that the initial viruliferous males passed the virus to 83% of the potential recipient females. These females then passed the virus to 39% of the males, which in turn passed it to 33%)
1.
15
TOMATO YELLOW LEAF CURL VIRUS
M^F-^M
2
3
4
Fig. 9 Serial transmission of Israel strain of tomato leaf curl vims (TYLCV-Is) from viruliferous Bemisia tabaci to insects of the other sex. One hundred viruliferous males (M) were caged with one hundred nonviruliferous females (F). After 24 hours, 60 females were collected and caged with 100 nonviruliferous males for an additional 24 hours and then analyzed for the presence of viral DNA by PCR. Sixty males were collected and caged with one hundred nonviruliferous females for 24 hours and then analyzed for the presence of viral DNA. The experiment was continued until four passages were completed, starting from the initial viruliferous males. The reciprocal experiment was conducted in a similar manner, starting with 100 viruliferous females caged with 100 nonviruliferous males. Bars indicate the percentage of viruliferous whiteflies in the insect populations, males (M) and females (F). TYLCV-Is was transmitted in a sexual manner for at least four passages.
of females, which finally passed it to 11% of the potential male recipients. In the reciprocal experiment, the beginning viruliferous females passed the virus to 67% of the males, which passed it to 39% of the females, which in turn passed it to 22% of the males, which finally passed the virus to 22% of the females. These successful serial passages of virus from whitefly to whitefly strongly suggest that TYLCV-Is also infects and mutliplies in its whitefly vector. C.
Sexual Transmission Spreads Virus in Whitefly Populations
Since TYLCV can likely be acquired by sexual contact, the virus can spread in an insect population without the presence of infected plants. We researched the con-
16
HENRYK CZOSNEK ET AL.
100-
M F
M F
dayO
day 3
M F
M F
80
'B 60 o
40 20-1
day 5
day 7
Fig. 10 Accumulation of viruliferous whiteflies with time in an insect population. Three separated populations, each consisting of three couples of viruliferous whiteflies and 120 nonviruliferous insects (about twice as many females as males) were reared on cotton plants, nonhost of an Israel strain of tomato leaf curl vims (TYLCV-Is). Insects were collected randomly after 3 days from population #1, after 5 days from population #2, and after 7 days from population #3. Males and females were separated. The presence of TYLCV-Is DNA in each individual was assessed by PCR. Bars indicate the percentage of viruliferous whiteflies in the insect populations, males (M) and females (F). The viruliferous population, males and females alike, increased as a result of sex-related transmission of TYLCV
tribution of this type of transmission to the number of viruliferous insects in defined whitefly populations. Three identical whitefly populations were established. Each population contained three viruliferous males and three viruliferous females together with 120 nonviruliferous insects (the male to female ratio was about 1:2). The three populations were reared separately on three cotton plants. The insects of the first population were collected after 3 days, those of the second after 5 days, and those of the third after 7 days (Fig. 10). Analyses of all insects by PCR indicated that after 3 days 21% of the males and 33% of the females contained viral DNA. After 5 days, these values increased to 50%) of the males and 47% of the females. After 7 days, 69% of the males and 51% of the females contained viral DNA. These results show that TYLCV spreads with time among insect populations; within 7 days the percentage of viruliferous males increased from about 1% at the start to 69% in the 7-day populations. Similarly, the percentage of viruliferous females increased from about 4% to 51%). These data further confirm the pathogenic nature of TYLCV-Is in its whitefly vector. D.
Whiteflies That Acquire Virus Sexually Can Transmit Virus to Tomato
We also determined whether whiteflies that have acquired TYLCV-Is from a sexual partner are able to infect tomato test plants. Twenty viruliferous females were placed with twenty nonviruliferous males in a membrane-feeding cage. After two days on the artificial diet, the males were collected and caged with tomato test
1.
17
TOMATO YELLOW LEAF CURL VIRUS
Plants infected by males caged with viruliferous females 1 2
3
4
5 6 7
9 10 11 12 13 14
15 16 17 18 19 20
Plants infected by females caged with viruliferous males 1
2
3
4
5
6
7
9 10
11 12 13
Fig. 11 Infection of tomato plants by Bemisia tabaci caged with viruliferous whiteflies of the other sex. (A) Twenty viruliferous females were mixed with twenty nonviruliferous males; after 48 hours, the females were collected and caged with tomato test plants, one insect per plant, for a 48-hour inoculation feeding period. (B) The reciprocal experiment was conducted. Plants were analyzed by Southern blot hybridization after 5 weeks, with use of an Israel strain of tomato leaf curl virus (TYLCV-Is) DNA probe [88]. Insects that acquired TYLCV-Is from viruliferous whiteflies of the opposite sex were able to infect tomato test plants.
plants, one insect per plant, for a 48-hour inoculation feeding period (IFP). Five weeks later, 5 of the 20 plants showed typical TYLCV-Is disease symptoms and contained viral DNA forms typical of infected tomato plants (Fig. 11 A). A reciprocal experiment was conducted, in which 20 viruliferous males mixed with 20 nonviruliferous females. Two of thirteen plants caged with surviving females became infected (Fig. IIB). Moreover, immunocapture PCR and Western blot immunodetection indicated that the symptomatic plants contained encapsidated virions. These results demonstrate that TYLCV-Is is transmitted among sexual partners in the form of an infectious encapsidated virion. We also were able to show that sex-related transmission increases the capacity of a whitefly population to infect tomato test plants. Two hundred nonviruliferous whiteflies from a stock colony (containing about twice as many females as males) were placed in a membrane-feeding cage with six opposite-sex couples of viruliferous whiteflies. After 48 hours, the surviving insects were collected and randomly divided into groups of three whiteflies each. Each group was caged with a healthy tomato test plant (58 plants altogether) for a 72-hour IFP. Four weeks thereafter, infection was determined by Southern blot hybridization. Given an optimal transmission scenario in which 12 viruliferous whiteflies are distributed
18
HENRYK CZOSNEK ET AL.
.sP^ 60 h 40
20
12/58
>
C^"
.^^
22/58
30
6/30
11/30
mi UL 20
10
I
II
Fig. 12 Contribution of sex-related transmission of an Israel strain of tomato leaf curl virus (TYLCV-Is) to the ability of Bemisia tabaci populations to inoculate plants with TYLCV Experiment I: Two hundred nonviruliferous whiteflies were caged with six couples of viruliferous insects. After 48 hours, groups of three insects were collected randomly; each group was caged for 72 hours with one of the 58 tomato plants tested. Experiment II: Two hundred nonviruliferous whiteflies were caged with three couples of viruliferous whiteflies; after 48 hours, each group of five insects was caged with one of the 30 test tomato plants. Infection was assessed by Southern blot hybridization after 4 weeks. In both experiments, the number of plants infected was higher than the number expected if sex-related transmission did not occur.
among 12 three-insect groups and each one is able to infect a tomato plant, a maximum of 12 tomato plants should have been infected. In fact, 22 plants were infected. In a similar experiment, three opposite-sex couples of viruliferous whiteflies were caged with 200 nonviruliferous ones. After the 48-hour IFP, the surviving whiteflies were divided into groups of five insects each, and each group was caged with one of 30 tomato test plants. Eleven plants were infected, instead of the expected six (Fig. 12).
W. Concluding Remarks Bemisia tabaci acquires and transmits begomoviruses in a circulative manner. Parameters of acquisition, circulation, transmission, and retention have been extensively investigated for a number of begomoviruses from both the Old and the New World. From these studies it appears that despite some variability, which is likely due to experimental procedures, these parameters are similar for all begomoviruses and therefore are intrinsic proprieties of the whitefly vector. During the
1.
TOMATO YELLOW LEAF CURL VIRUS
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8- to 12-hour latent period, ingested virus circulates in the insect along a path seemingly shared by all begomoviruses. Even the velocity of translocation in various vector and organ systems seems shared by all begomoviruses, as determined for viruses as different as TYLCV-Is and SLCV Begomoviruses acquired by B. tabaci during a short AAP remain associated with the insect for several weeks. Some, like TYLCV-Is, can be found in the vector during its entire life-span [17]. Others, such as TYLCV-Sar [69], vanish several days before the insect dies. In all cases, the virus acquired during a 24-hour AAP is present in the insect for much longer than the time needed for all virus to be ejected during the latent period. Instead, whiteflies are able to infect test plants for several days or even weeks. Moreover, viral DNA persists in the insects longer than does infectivity [17, 69]. These observations suggest that at least some of the acquired virions are able to remain in the vectors as infective units within the circulative pathway or to reenter the pathway for several weeks after acquisition. The question of the fate of the ingested virus is intriguing. It seems that a large fraction of the TYLCV-Is virions acquired by B. tabaci from infected tomato plants during a 48-hour AAP soon leaves the transmission pathway. Whereas viral DNA remains detectable during the insect's life-span, coat protein is not detectable after 12 days. The capacity of viruliferous insects to transmit the virus progressively decreases with time without disappearing entirely. There is no alternative but to propose that the virus invades insect organs and tissues, where it is stored in insect cells. We do not know whether virus transmitted by 3- to 4-week-old whiteflies consists only virus acquired by these insects 3 to 4 weeks before. Persistence of the viral genome and in some cases, accumulation of viral DNA suggests a certain level of replication [70]. Invasion of the vector's reproductive system by TYLCV-Is and its effect on vector fecundity have been demonstrated. Long-time association of TYLCV-Is with B. tabaci is accompanied by about a 40% decrease in the mean number of eggs laid. These negative effects are expressed several days after acquisition, as if the virus first has to invade the reproductive system and somehow cause developing eggs to be aborted [17]. Indeed, eggs maturing in the ovaries of viruliferous whiteflies contain viral DNA detectable by PCR. Viral DNA was similarly detected in some adult insects that developed from these eggs [71]. The way in which TYLCV penetrates the whitefly reproductive system is unknown. We presume, but we do not have direct proof, that TYLCV-Is invades tissues others than those of the reproductive system, reducing the life expectancy of the insect population by about 20% [17]. Pesic-van Esbroeck et al. [72] reported, in abstract form, immunolocalizing squash leaf curl virus in organs of the reproductive, digestive, and excretory systems. They further reported that vector exposure to virus is associated with gross as well as ultrastructural abnormalities of the affected organs [72]. Unfortunately, a full report has not followed. TYLCV-Is is the first geminivirus shown to be spread among whiteflies in a sex-related manner. The virus can be transmitted from viruliferous males to
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females, and vice versa, but not between insects of the same sex. Virus spread can occur when insects are reared in groups or in couples on an artificial diet or on a TYLCV nonhost plant. The recipient insects are able to efficiently infect tomato test plants, indicating that virions so acquired are infectious and transmissible. Insect-to-insect transmission results in increasing numbers of whiteflies being able to infect tomato test plants. Taken collectively, and unless other some unknown mechanism of virus transmission exists, our results can only be explained by sexual transmission of TYLCV-Is. We do not know how TYLCV-Is is transmitted among sexual partners or if the insect's reproductive organs are infected. The fact that virus can be found in the hemolymph of recipient males or females, albeit not earlier than approximately 4 hours after males and females have been caged together [50, 89], suggests several possible modes of transfer. If sperm cells or seminal fluid of viruliferous male insects contain TYLCV, which is by no means proven, virus might be so transmitted to the female during copulation. Sperm migrates to the spermatheca, where it can be used or not by the female to fertilize eggs [73]. Eggs could be infected at fertilization and give rise to progeny containing TYLCV, as we indeed observed [89]. Alternatively, virion-containing sperm could be injected into the hemocoel [74]. The latter mechanism might explain the presence of virus in the hemolymph of recipient female whiteflies after copulation with viruliferous males. Similarly, it is not clear how viruliferous females contaminate males with virus. Virus is also found in recipient male hemolymph, which suggests contamination through the body fluids. During copulation, hemolymph of male and female might mix, thereby favoring the passage of TYLCV from one individual to the other. Indeed, whiteflies have an open circulating system, with hemolymph in the body cavity or hemocoel flowing over various organs, bathing them directly and providing them with vital nutrients [75]. Virus reaching the hemolymph during sexual activity would have to translocate to the salivary glands in order to be inoculated to tomato plants. In the hemocoel, sexually acquired virus would face the same perils and need to adopt the same survival strategies as do virions acquired via ingestion from infected plants. To understand the virus-vector interactions we have described, one needs to address the questions of when and how this virus-vector relationship started and how it has evolved to modem times. Until geminiviral DNA is identified in fossilized whiteflies, only indirect evidence indicates that geminivirus-whitefly interactions date far back in geologic time. Several fossil whitefly species, though not B. tabaci, have been found in amber [76]. Two of them, Bernaea neocomica and Heidea cretacia, are in 120 to 140 million-year-old amber from Lebanon [77, 78]. Fossil geminiviral DNA sequences related to a modem Old World begomovims have been found as a cluster of approximately 25 multiple direct repeats integrated into a single chromosomal locus of Nicotiana tabacum; they have also been found in three related Nicotiana species, all in the section Tomentosae, but not in nine other more distantly rQlatQd Nicotiana species [79, 80]. These findings indicate that these viral
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elements descend from a unique geminivirus integration event that occurred in a common ancestor of the Tomentosae species [81]. Molecular analyses of geminivirus and insect marker genes [20, 29] point to geographically separated virus-insect combinations stemming from long-term interaction and co-adaptation. While feeding on infected plants, whiteflies ingest very large amounts of virus, which remain associated with the insect for at least several days. Although this aspect of the begomovirus biology has not been thoroughly addressed, it seems that most begomoviruses have no dramatic deleterious effects on their whitefly vector. We do not know whether the ancestral begomoviruses had a similar quasineutral relationship. The pathogen-like behavior of TYLCV-Is suggests that this may have not been the case. A long-lasting virus-vector relationship of this magnitude implies that both partners have developed coevolutionary mechanisms, which ensure on one hand the survival and efficient transmission of the virus and on the other hand the protection of the insect host from possible deleterious effects of the virus. Both mechanism can be recognized in many aspects of today's begomovirus-5. tabaci combinations. In the case of geminivirus-5. tabaci pairings, it is likely that the virus is the partner that has adapted to the insect to maximize its ability to be transmitted. The begomovirus needs the insect to spread to host plants, where it can replicate and spread. Therefore, native virus might have evolved toward a better adaptation of its capsid to putative receptors and proteins of the insects of the same region. As a result, begomoviruses from a given region tend to be closely antigenically related, although they may infect different plants [82]. Optimization of virus transmission efficiency might have been the driving force behind capsid adaptation. Ancestor whiteflies, on the other hand, may have perceived the presence of such large amounts of virus as a potential threat from an alien invader. It is possible that primitive geminivirus-like particles had a deleterious potential far greater than that presented by modem TYLCV-Is. In the absence of an effective immune system, how, if at all, does the insect respond to the deleterious presence of the virus? It is possible that the evolutionary trend of the geminivirus-vector partnership was neutralization of any viral function that might negatively impact the insect and its progeny, such as long-term sojourn, replication, invasion of tissues, and transovarial transmission. Mechanisms evolved to ensure the smooth transit of ingested virus through the insect body and its rapid expulsion (transmission to plants). Some features of geminivirus-5. tabaci interaction may be interpreted in the light of these hypothetical evolutionary processes. Two insect strategies might be implemented, exclusively or simultaneously. The first would be to destroy incoming virions, the second to facilitate rapid transit of virus until it is safely expelled. Evidence is available to support both strategies. Whitefly extracts contain proteolytic enzymes that degrade phloem proteins in vitro, suggesting that these enzymes act in the gut [83]. Nonetheless, virions are not degraded in the insect digestive tract, at least not to the point of vanishing. Either the geminivirus architecture itself is not prone to degradation or cellular
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components may protect virions from destruction in the whitefly gut. Furthermore, proteins having antiviral properties are reportedly synthesized following acquisition of TYLCV-Is [84, 85]. It is not known whether these proteins constitute a broad response to any potential invader or a virion-specific response. From the gut, the virions penetrate the gut wall to the hemolymph en route to the salivary gland: this is the longest and most hazardous phase of the circulative path. Successful passage of virions is ensured by a GroEL-like chaperonin-synthesized endosymbiotic bacterium in the mycetome of B. tabaci. The same viral survival strategy has been described for an unrelated virus, aphid-transmitted potato leaf roll virus [52]. The whitefly GroEL homologue is synthesized in the cytoplasm of coccoid endosymbionts that are morphologically similar to aphid primary endosymbionts [61, 86]. The structural and biological properties of the B. tabaci GroEL homologue are strikingly similar to those of the GroEL homologue in M. persicae [52]. The latter suggests a conserved mechanism underlying circulative transmission of plant viruses, which may date back to the early days of insect-bacteria symbiosis [59]. Ancestor insects may have taken advantage of endosymbiotic proteins to facilitate virion circulation and expulsion. Apparently, this live-and-let live strategy has not that been adopted by other whitefly species. For example, the whitefly Trialeurodes vaporariorum is able to acquire but not transmit geminiviruses [49, 66, 67]. Once ingested during feeding, virus moves through the digestive tract but is unable to traverse the gut epithelial cells to the hemolymph and subsequently the salivary glands [49]. Virus is destroyed in the digestive system or excreted in the honey dew. We do not know the effects, if any, of begomoviruses on the longevity and fertility of T. vaporariorum. Infection of plant hosts by begomoviruses is still not fully understood, although the function of geminivirus genes has been thoroughly investigated for the last decade. We are beginning to understand how these viruses replicate and spread in plants. However, we still do not know how the virus interacts with unknown host factors to induce disease symptoms. We also do not know how geminivirus replication is repressed in resistant hosts or immune nonhost plants, or how resistance genes function. The way begomoviruses interact with their insect vectors is even less understood. It is abundantly clear that whiteflies are not mere go-betweens in the transmission process. A begomovirus such as TYLCV-Is shares many features with insect viruses. Other begomoviruses may have retained or gained some pathogenic features during their coevolution with B. tabaci that have yet to be discovered.
Acknowledgments This study was supported by grant 95-168 from The United States-Israel Binational Science Foundation (BSF), by grant IS-2566-95R from the United States-Israel Binational Agricultural Research and Development Fund (BARD), and by grants from the Chief Scientist of the Ministry of Agriculture Israel.
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80. Ashby, M.K., Warry, A., Bejarano, E.R., Kashoggi, A., Burrell, M., and Lichtenstein, C.P. (1997). Analysis of multiple copies of geminiviral DNA in the genome of closely related Nicotiana species suggest a unique integration event. Plant Mol. Biol. 35, 313-321. 81. Gerstel, D.U., and Sisson, VA. (1992). Tobacco. In "Evolution of Crop Plants" (J. Smartt and N.W. Simmonds, eds.). Longman Scientific and Technical, London. 82. Harrison, B.D., and Robinson, D.J. (1999). Natural genomic and antigenic variation in whiteflytransmitted geminiviruses (begomoviruses). Annu. Rev. Phytopathol. 37, 369-398. 83. Salvucci, M.E., Rosell, R.C., and Brown, J.K. (1998). Uptake and metaboHsm of leaf proteins by the silverleaf whitefly. ^rc/i. Insect Biochem. Physiol. 39, 155-165. 84. Cohen, S., and Marco, S. (1970). Periodic occurrence of an anti-TMV factor in the body of whiteflies carrying the tomato yellow leaf curl virus (TYLCV). Virology 40, 363-368. 85. Marco, S., Cohen, S., Harpaz, I., and Birk, Y. (1972). In vivo suppression of plant virus transmissibility by an anti-TMV factor occurring in an inoculative vector's body. Virology 7, 761-766. 86. Costa, H.S., Westcot, D.M., Ullman, D.E. and Johnson, M.W. (1993). Ultrastructure of the endosymbionts of the whitefly, Bemisia tabaci and Trialeurodes vaporariorum. Protoplasma 176, 106-115. 87. Navot, N., Pichersky, E., Zeidan, M., Zamir, D., and Czosnek, H. (1991). Tomato yellow leaf curl virus: A whitefly-transmitted geminivirus with a single genomic component. Virology 185, 151-161. 88. Ber, R., Navot, N., Zamir, D., Antignus, Y, Cohen, S., and Czosnek, H. (1990). Infection of tomato by the tomato yellow leaf curl virus: Susceptibility to infection, symptom development and accumulation of viral DNA. ^rcA. Virol. 112, 169-180. 89. Ghanim, M., and Czosnek, H. (2000). Tomato yellow leaf curl geminivirus (TYLCV-Is) is transmitted among whiteflies {Bemisia tabaci) in a sex-related manner. J. Virology 74, 4738^745, 2000.
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CHAPTER 2
Possible Etiology of Eriophyid Mite-Borne Pathogens Associated with Double Membrane-Bound Particles KYUNG-SOO KIM KYUNG-KU ABN ROSE C. GERGERICH
SuNG-BooN KIM
/. Introduction A large group of economically important plant diseases occurring worldwide, some with and some without known etiologies, has been reported to be associated with phytophagous mites [1-4]. Among the four families of phytophagous mites, the one that contains proven vectors of plant pathogenic agents, Eriophyidae, is divided into five subfamilies [2]. Genera with species reported to be vectors of plant pathogens, however, occur only in three of the subfamilies: Cecidophyosis and Colomerus in the Cecidophyinae, Eriophyes (or Aceria) and Phytopsis in the Eriophyinae, and Abacarus, Aculus, Calacrus, and Phyllocoptes in the Phyllocoptinae [2, 3]. These mites are known to be vectors of viruses but have not been demonstrated to transmit other plant pathogens such as bacteria, fungi, or phytoplasmas. At present, at least 20 plant diseases have been reported to be caused by viruses or viruslike agents transmitted by eriophyid mites. However, only a small number of these diseases have a confirmed viral etiology; many have uncertain etiologies and remain identified as viruslike diseases [ 2 ^ ] . Since most eriophyid miteassociated diseases, including those of unknown etiology, produce viruslike symptoms (mosaic, chlorosis, mottling, vein clearing, stunting, etc.), investigators have assumed that diseases such as wheat spot mosaic, pigeon pea sterility
Viras-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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KYUNG-SOO KIM ET AL.
mosaic, and High Plains are caused by viruses [2, 3]. A number of such assumptions were eventually proved true. Black currant reversion and cherry mottle leaf diseases, assumed to be virus diseases for many years, have been characterized sufficiently to confirm their viral etiology [5-7]. Eriophyid mite-associated diseases have generally been divided into two groups based on whether the host plants are monocotyledonous or dicotyledonous [2, 4]. In this chapter, we add an extra group solely on the basis of ultrastructural cytopathology, those diseases consistently having large double membrane-bound particles in the cytoplasm of cells in diseased plants. We will briefly review the eriophyid mite-associated diseases, especially those having double membrane-bound particles (DMPs) with a quasi-spherical, viruslike structure and diameters of 100-200 nm. Since the first report of DMP-associated wheat spot mosaic in the late 1960s [8, 9], the number of diseases in this group has been growing, including the economically important rose rosette disease [10, 11] and High Plains disease [12, 13]. Although no definite properties of these DMPs have been confirmed, some progress has been made in terms of their identity. Emphasis will be placed on the ultrastructure of DMPs and associated inclusions revealed in the most recently discovered diseases such as thistle mosaic [14, 15] and High Plains [13, 15], both of which affect herbaceous hosts. In this chapter we hope to establish, based on cytopathological and immunogold labeling studies, that these DMPs are indeed viral in nature and may represent a new group of plant viruses.
//. Groups of Eriophyid Mite-Associated Diseases A.
Diseases Caused or Possibly Caused by Rymoviruses
Eriophyid mite-associated diseases can be divided into three groups. As shown in Table I, the largest group of eriophyid mite-borne plant pathogens that have been confirmed to be viruses contains 10 viruses that infect monocots, including wellcharacterized viruses such as wheat streak mosaic [16-18], agropyron mosaic [19-21], and ryegrass mosaic [22, 23]. The viruses all have flexuous rod-shaped particles measuring about 700 nm in length, and most of them are serologically related to each other [3]. Also, most are easily transmitted by mechanical means and are known to induce cylindrical or "pinwheel" inclusions [3], an important taxonomic criterion for potyvirus classification [24, 25]. Thus, these viruses have been considered and referred to in the literature as potyviruses [2, 3,26], although there are no serological relationships to the aphid-borne, "true" potyviruses [27]. Recently, it was proposed that eriophyid mite-borne, flexuous, rod-shaped viruses be placed in the genus Rymovirus in the family Potyviridae, with ryegrass mosaic virus as the type virus [27].
Table I Three Groups of Plant Diseases Associated with Eriophyid Mites as Vectors of Known and Unknown Etiology Disease Group 1 Wheat streak mosaic (WSMV) Ryegrass mosaic (RGMV) Agropyron mosaic (AgMV) Hordeum mosaic (HoMV) Oat necrotic mottle (ONMV) Brome streak mosaic Onion mite-borne latent Shallot mite-borne latent Spartina mottle Garlic mosaic
Vector^
Virus group
Rymovims A. tosichella (formerly A. tulipae) (Potjrviridae)
Key references Oldfield [93], Slykhuis [94], Nault and Styer [95], Brakke [18] Mulligan [22], Slykhuis and Paliwal [23] Slykhuis [19,21]
unidentified mite A. tosichella
Rymovims (Potyviridae) Rymovims (Potyviridae) Rymovims Slykhuis [96], Slykhuis and Bell [97] (Pot3rviridae) Gill [98-100] Rymovims (Potyviridae) possible Rymovims Milicice^fl/. [101,102] possible Rymovims VanDijke^fl/. [103]
A. tosichella
possible Rymovims VanDijke^fl/. [103]
unidentified mite A. tosichella
possible Rymovims Jones [104] possible Rymovims Ahmed and Benigko [105]
E. inaequalis
Closterovims
E. incidiosus
Closterovims
C. ribis
possible Closterovims or Nepovims
A. tosichella
DMP^
Wheat spot chlorosis (WSpC) Pigeon pea sterility Yellow ringspot ofredbud Rose rosette
A. tosichella
DMP
E. cajani unidentified mite
DMP DMP
P. fructiphilus
DMP
Fig mosaic
E.flcus
DMP
Thistle mosaic High Plains
unknown A. tosichella
DMP DMP
Group 2 Cherry mottle leaf {CMoYSf) Peach mosaic (PMV) Black currant reversion Group 3 Wheat spot mosaic (WSpM)
A. hystrix E. hystrix unidentified mite unidentified mite
James and Mukerji [5], James [6], Oldfield [93] Oldfield e^«/. [106], GispQYtetal. [33], Creamer e^fl/.[32] Amos etal. [107], Roberts and Jones [34], LQmmQtty et al. [7] Slykhuis [42], Hiruki etal. [60], Chen and Hiruki [91], Bradfute and Nault [8] Nault and Styer [17], Bradfute et al. [9] Hiruki [3] Kim and Martin [49], Ahnetal. [15] Gergerich e^fl/.[10], Kim and Gergerich [11] Appiano [37], Appiano et al. [44], Ahnetal. [15] Ahnetal. [15] Jensen and Lane [40], Jensen et al. [76], Ahnetal. [13]
« Genus abbreviations: A. = Aceria (Eriophyes), E. -- Eriophyes, C. = Calacarus, P. = Phyllocoptes. ^ DMP = double membrane-bound particles.
31
32
B.
KYUNG-SOO KIM ET AL.
Virus or Viruslike Diseases of Woody Dicots
A small group of economically important diseases of woody fruit trees is known to be associated with eriophyid mite vectors [2, 3]. Black currant reversion [7, 28, 29], peach mosaic [30, 31], and cherry mottle leaf diseases comprise this group. They may also represent the earliest plant diseases demonstrated to be mite-associated. The association was recognized as early as the 1920s and 1930s. Progress in establishing the etiologies of these diseases, however, has been slow. Until recently, these diseases were listed as having unknown etiology [2, 3]. Recently, however, viruses with closterovirus-like properties were identified as the causal agents of cherry mottle leaf [5, 6], peach mosaic [32, 33] and black currant reversion [34] diseases. More recently, a new virus with 27-nm isometric particles and affinities with nepoviruses was isolated from plants affected with a severe form of black currant reversion [7]. This is the first isometric virus isolated from plants with an eriophyid mite-associated disease. However, no conclusive evidence that this virus causes black currant reversion disease has been presented. C. Diseases Associated with Double Membrane-Bound Particles The last group of diseases caused by known or suspected eriophyid mite-borne agents is categorized based on the ultrastructural cytopathology of diseased plant cells, particularly the presence of DMPs in the cytoplasm. In the past, DMPs were referred to as double membrane-bound bodies (DMBs). "Bodies" might be interpreted as connoting cellular components such as lipid and/or protein bodies or organelles such as microbodies. In this article, therefore, we prefer the term particles. Also, there is strong evidence that DMPs are viral in nature. The number of diseases in this group, which was only a few until the early 1990s, has grown (Table 1). Unlike the two groups mentioned above, the diseases in this group affect different types of plants, ranging from trees and other dicots to grasses and other monocots such as wheat and com. Some of the diseases associated with DMPs are economically important, such as fig mosaic [35-37], rose rosette [38, 39], and High Plains disease. The latter disease damages wheat and com crops, causing yield losses up to 80% in some cases [12, 40]. The causal agents of the diseases in this group are not sap-transmissible but are readily transmissible by grafting. Symptomatology includes foliar symptoms such as mosaic, chlorotic spots or streaks, and general chlorosis and mottling, all of which are commonly associated with known vims infections. The diseases in this group are not associated with the presence of specific ftingi, bacteria, or mycoplasma-like organisms, which ftxrther suggests that the causal agents are viral in nature. No typical vims or viruslike particles have been located in cells of diseased plants with any of the diseases. However, diseases of this group are consistently associated with DMPs. The nature of DMPs is still unknown despite efforts by several groups of investigators to characterize them.
2.
ERIOPHYID MiTE-BoRNE PATHOGENS
1.
33
WHEAT SPOT MOSAIC DISEASE
Wheat spot mosaic disease (WSpMD) is the first mite-related disease to be associated with DMPs in infected cells [8, 9, 41]. The DMPs of WSpMD are ovoid, approximately 100-200 nm in diameter, and present only in the cytoplasm of parenchyma, phloem, and epidermal cells. Some DMPs are elongated and medially constricted, which suggests binary fission. The internal constituents of DMPs consist of dispersed fibrils, possibly representing nucleic acid [8, 9]. However, neither ribosomes (characteristic of bacteria, mycoplasma-like organisms, and the psittocosis group of organisms) nor electron-dense central core regions (characteristic of many known viruses) were observed in the DMPs of wheat spot mosaic-diseased cells [8]. One of the most prominent cytopathic features of DMP-containing cells is the proliferation of membrane systems, in particular the rough endoplasmic reticulum to which DMPs are often physically connected [3]. WSpMD was first detected in Alberta, Canada in 1952 when Aceria (Eriophyes) tosichella Keifer (formerly Aceria tulipipae Keifer) was recognized as a vector of wheat streak mosaic virus (WSMV) [16, 42]. In the field, it was noted that diseased plants exhibited severe symptom expression and were usually doubly infected with WSMV [2, 42]. The double infection appeared synergistic with respect to symptom expression [2, 3,42]. The eriophyid mite vector, A. tosichella, transmits both WSMV and the agent of WSpMD, and the host range of WSpMD agent is also similar to that of WSMV [2]. It should be noted, however, that WSMV is sap-transmissible, whereas the WSpMD agent is vector-dependent for transmission [2]. 2.
WHEAT SPOT CHLOROSIS DISEASE
In 1970, almost 20 years after the report of WSpMD in Canada, an eriophyid mite-borne agent was recovered from diseased wheat and corn in Ohio [17]. The host range and symptomatology of the agent were similar to those of WSpMD [2, 42]. Thin-section electron microscopy of infected leaves of wheat, maize, and barley revealed the presence in the c3^oplasm of DMPs structurally similar to those associated with WSpMD [8, 9]. It was suggested that the pathogen of wheat spot chlorosis disease (WSCD) is an isolate of WSpMD [2]. 3.
FIG MOSAIC DISEASE
Fig mosaic disease (FMD), a classic eriophyid mite (Eriophyes ficusj-sissociated disease, was first reported in California in the early 1930s [43]. The disease is widespread and occurs naturally in most fig-producing areas [2, 3, 37, 44, 45]. Symptoms of FMD include a mosaic ranging from varied chlorotic spots to light green blotches, chlorotic mottling, and occasional leaf malformation. Association of DMPs with FMD was first reported from diseased fig plants in the United
34
KYUNG-SOO KIM ET AL.
States [9], followed by reports from Libya and Yugoslavia [46] and Italy and England [37, 44, 45]. The DMPs of different isolates of FMD are structurally indistinguishable from each other, with the exception of an English isolate. The latter is characterized by two types of DMPs, ovoid and cylindrical, which occur together in the cytoplasm [47]. One of the most significant cytopathic effects oberved in FMD-affected cells are nonmembranous tubular inclusions associated with the DMPs, called "tubular matrices" by Appiano and co-workers [37, 44, 48]. The DMPs and tubular matrices are specifically associated with both the endoplasmic reticulum and Golgi apparatus, suggesting a relationship between the two in DMP formation [44]. Tubular inclusions associated with DMPs, similar to those in European isolates of fig mosaic, were also found in a California isolate (Fig. la and b) [15]. 4.
REDBUD YELLOW RINGSPOT DISEASE
Redbud yellow ringspot disease (RYRD) is the first of four eriophyid mite and DMP-associated diseases discovered in northwest Arkansas and studied by our laboratory in the late 1970s [49]. Redbud, Cercis canadensis L., was noted with severe symptoms suggestive of virus infection, including bright yellow ringspots, oakleaf patterns, and necrotic spots in the older leaves. Trials for sap transmission to redbud and other herbaceous hosts failed. Thin-section electron microscopy, however, revealed DMPs containing fibrillar or granular material or both. The DMPs occurred randomly in the cytoplasm, often near peroxisomes, in leaf cells of yellow ringspot diseased redbud (Fig. la). A cytochemical study, however, failed to detect catalase in the DMPs [49]. Despite extensive efforts to locate DMP-associated inclusions, such as the tubular inclusions associated with DMPs of FMD (Fig. lb), no structural features suggestive of viral infection were evident. Large numbers of unidentified mites were collected from the yellow ringspot-diseased trees. These mites were transferred to healthy seedlings and after several months the typical symptoms of the disease were noted on these seedlings (unpublished data). Redbud trees with RYRD symptoms are very common in northwest Arkansas. Smaller leaves, dead twigs, and smaller branches are associated with diseased trees. It should be mentioned that the original three symptomatic redbud trees (one ftilly grown and two young seedlings with only a few branches) were all dead within a few years after sampling. 5.
ROSE ROSETTE DISEASE
Rose rosette disease (RRD), also referred to as witches' broom of rose, was first reported from Manitoba, Canada in the early 1940s [50, 51]. Since then the disease has been reported in the western and midwestern United States and is
2.
35
ERJOPHYID MITE-BORNE PATHOGENS
I
F;V...
M
la
~
' . ' r ^ C ' - ' ^ v - - - - , : , ' * ^ . . . . .;•
•O
;•. •• • :
,• •.v.- .• ••:. .;/:>••• ••yh'H'••
.•
^ ' ' V •••••"' O ; .^•^* * ..**
•:^^^:#^
^'i^'-.^^
F/g^. / Double membrane-bound particles (DMPs) (arrowheads) associated with redbud yellow ringspot, fig mosaic, rose rosette, and thistle mosaic diseases, (a) Four DMPs containing fibrils are shown close to a large microbody (M) in a leaf mesophyll cell of redbud leaf infected with yellow ringspot disease. Bar = 100 nm. (b) A large mass of tubular inclusion (TI) surrounded by DMPs (arrowheads) in fig mosaic-diseased leaf cells. Bar =100 nm. (c) DMPs similar to those in (a) and (b) in a mesophyll cell of a multiflora rose infected with rose rosette. Bar = 100 nm. (d) A large amorphous inclusion (AI) body is encircling a mass of tubular inclusion (TI). Bar = 250 nm. (e) A low magnification view of an epidermal cell of a thistle leaf with thistle mosaic disease, showing DMPs (arrowheads) throughout the cytoplasm. The DMPs in the square are associated with an amorphous inclusion body. Bar = 1,000 nm. (f) A higher magnification of the squared area in (e), showing the details of DMPs (arrowheads) and associated viroplasm-like amorphous inclusion (AI). Bar =100 nm. N = nucleus, G = Golgi body.
36
KYUNG-SOO KIM ET AL.
spreading rapidly eastward [3, 4]. The RRD causative agent attacks cultivated roses as well as native and introduced wild rose species. In fact, multiflora rose, Rosa multiflora, introduced from East Asia and now a serious pest on nontilled lands in the central United States, is severely affected by RRD [39]. Therefore, RRD has been considered as a biological control agent for this weed species [39, 52, 53]. Characteristic symptoms of RRD include misshapen, stunted leaves with bright red pigmentation. The proliferation of the short secondary shoots leads to a witches' broom or rosette appearance. The pathogen is transmitted from diseased to healthy roses by the eriophyid mite Phyllocoptes fructiphilus Keifer [38, 54]. Based on successful graft transmission from diseased to healthy roses [38, 54] as well as on involvement of a specific eriophyid mite vector, viral etiology has been suggested but not proved. In the early 1980s, RRD was widespread in Arkansas, Oklahoma, and Missouri in multiflora rose as well as in commercial roses [10, 55]. Thin-section electron microscopy of both stunted leaves with red pigmentation and mottled leaves revealed the consistent presence of DMPs (Fig. Ic). These DMPs occurred singly or as groups of several particles in the cytoplasm associated with proliferated endoplasmic reticulum [10, 11]. However, no tubular inclusions such as those associated with DMPs of fig mosaic disease were evident in cells of RRDinfected tissue. The studies of RRD were expanded by sampling RRD specimens from the neighboring states of Oklahoma, Kansas, and Missouri; without exception, DMPs were found in every specimen studied, confirming the association of RRD with DMPs [11]. Mechanical transmission using sap from symptomatic leaves to healthy roses and other herbaceous hosts failed. 6.
PIGEON PEA STERILITY MOSAIC DISEASE
Pigeon pea sterility mosaic disease (PPSMD) was first reported from India in the early 1930s and has been recognized as one of the most important constraints of pigeon pea production in India [56]. The eriophyid mite Aceria cajani Channa Basavanna has been reported by several investigators as a vector of PPSM agent [57, 58]. Symptoms include chlorosis, mosaic mottling, stunting, and partial or complete sterility [4, 57]. A viral etiology has been proposed, mainly because the agent is transmitted by an eriophyid mite (A. cajani) and by grafting [56, 58]. However, no virus or viruslike particles have been reported associated with the disease. Hiruki [3] has reported the presence of DMPs measuring 100 to 200 nm in diameter in the cytoplasm of leaf parenchyma cells of diseased plants (based on unpublished data without a supporting electron micrograph). Since Dr. Hiruki has published a number of papers on the subject of DMPs linked with eriophyid mite-associated diseases [3, 41, 59, 60], we assume that the DMPs he observed in PPSM-diseased plants are similar to those of other DMP-associated diseases.
2.
ERIOPHYID MiTE-BoRNE PATHOGENS
7.
37
THISTLE MOSAIC DISEASE
Thistle mosaic disease (ThMD) is one of the latest DMP-associated diseases discovered and studied by our laboratory [14, 15]. Field thistle, Cirsium discolor (Mull) Spreng, exhibiting typical viruslike symptoms was noticed in a roadside ditch in northwest Arkansas. The symptoms consisted of chlorotic spots of irregular size and shape intermingled with green areas, resulting in a mosaic pattern. Older leaves showed a more intense degree of chlorosis, exhibiting a yellowish pigmentation. Yellow vein clearing and puckering at the margin of the leaves were also common. The consistent presence of DMPs, similar to those associated with other diseases such as WSpMD, FMD, RRD, and RYRD, was noted by electron microscopy in symptomatic leaf specimens sampled numerous times in the past several years. Unlike other DMP-associated diseases, the DMPs in symptomatic thistle were numerous and densely scattered throughout the cytoplasm in many cells. In addition, DMPs were associated with inclusion bodies resembling those induced by other known viruses and interpreted as viroplasms (Fig. le and f) [15]. The viroplasmic-like structures associated with DMPs in ThMD-affected thistle provide additional cytopathological evidence that DMPs may represent structural entities of viruses that cause such diseases. There are two DMP-associated inclusions in thistle mosaic, an electron-dense amorphous inclusion (Fig. If) and an intertwined tubular inclusion (Fig. Id) [15]. Amorphous inclusions occur either as a great number of small round patches about the size of DMPs (Fig. 2a) or as large masses (Fig. 2b), which are often present together in the same area with the small patches (Fig. 2a). In either case, the inclusions are accompanied with greatly proliferated smooth membranes apparently derived from rough endoplasmic reticulum (Fig. 2a and b). Careful scrutiny revealed that the round patches are mixed with partially double membrane-bound and fiilly double membrane-bound particles (Fig. 2a), suggesting that the patches (non-membrane-bound) are the precursors of DMPs. In addition, the patches and DMPs are directly associated with proliferated membranous vesicles or tubules (Fig. 2a), suggesting that they are the source of the double membranes of the particles. In addition, the proliferated smooth membranes are at some point continuous with rough endoplasmic reticulum, indicating a relationship between the two (Fig. 2a). The amorphous inclusions often occur as large masses associated with proliferated rough endoplasmic reticulum, smooth membranes, and DMPs. The enclosure of small portions of amorphous material by double membranes (Fig. 2c) suggests that this represents a step in the process of DMP assembly. The other type of inclusion in thistle mosaic consists of thin, intertwined, tubular structures appearing as tubular aggregates surrounded by DMPs (Fig. lb). Inclusions of the two types, amorphous and tubular, occur either independently or closely associated at the same sites. Often, a large amorphous inclusion contains a compact aggregate of tubular inclusions in its center (Fig. Id)
38
KYUNG-SOO KIM ET AL.
Fig. 2 Amorphous inclusions associated with double membrane-bound particles (DMPs) in cells infected with thistle mosaic, (a) Epidermal cell showing amorphous inclusions (AI) that are scattered as small round patches (arrows) throughout the cytoplasm. DMPs (arrows) and greatly proliferated membranous vesicles and tubules (Me) are associated with the inclusion patches. W = cell wall, Ch = chloroplast. Bar = 1,000 nm. (a, insert) Higher magnification of an area in (a) showing the details of the small patches of amorphous inclusion (arrows), DMPs (arrowheads), partially double membrane-bound particles (X) and associated membranes (Me). A segment of rough endoplasmic reticulum (ER) is continuous with a tubular membrane (double arrowheads). Bar = 100 nm. (b) Large masses of amorphous inclusion bodies (AI) associated with DMPs (arrowheads) and proliferated endoplasmic reticulum (ER). Bar = 500 nm. (c) A DMP (arrow) seems in the process of maturation or budding, as shown by segments of double membranes partially enclosing amorphous inclusion material (AI). Bar = 100 nm.
2.
ERIOPHYID MiTE-BoRNE PATHOGENS
39
Plant viruses often induce intracellular inclusions that are indicators of virus infection. Most of these virus-induced inclusions, including viroplasms characteristic of particular or related viruses or both, have been used as valuable tools in virus identification and taxonomy [25, 61-65]. In many cases, virus infection can be recognized by the presence of virus-specific inclusions rather than by the presence of virus particles. Good examples of characteristic inclusions are cylindrical (or pinwheel) inclusions of potyviruses [61], virus-containing tubular structures of como- and nepoviruses [64, 66, 67], amorphous inclusions of tospoviruses, and phytoreoviruses [64, 68, 69], and viroplasms of caulimoviruses [70, 71] and animal or insect poxviruses [72]. Virus-induced inclusions usually consist of virus particles, virus-related materials (amorphous, fibrillar, granular, filamentous, or other proteinaceous structures with conspicuous morphologies) or ordinary cellular constituents that have an abnormal appearance (e.g., proliferated cell membrane systems, increased ribosomes, and vesiculated mitochondria, Golgi apparatuses, chloroplasts, and glyoxysomes). These occur either singly or, more often, in various proportions [62-64]. The amorphous inclusions associated with thistle mosaic are structurally very similar to those induced by tospoviruses, caulimoviruses, and phytoreoviruses [64, 68, 69]. Furthermore, the DMPs and associated amorphous inclusions (Fig. le and f) closely resemble immature particles and viroplasms of animal and insect poxviruses [72, 73]. 8.
HIGH PLAINS DISEASE
High Plains disease (HPD) is the disease we have most recently shown to be associated with DMPs [13, 74]. A new viruslike disease affecting maize and wheat was identified in several locations in the High Plains region of central western United States including the Texas panhandle, western Kansas, northern Colorado, and central Idaho [12, 40, 75]. The causal agent was not mechanically transmitted but was transmitted by the "wheat curl" eriophyid mite, Aceria tosichella Keifer (formally A. tulipae), which is also the vector of WSMV [16]. Initial attempts to identify the causal agent as a possible virus through transmission and serological tests revealed the presence of WSMV in field-collected samples [12, 40, 76]. Polyacrylamide gel electrophoresis of proteins in partially purified preparations from symptomatic maize leaf tissues revealed a distinct 32kDa protein band, in addition to the 44-kDa coat protein band of WSMV [12,40], strengthening the argument that a second pathogen was present. Since the size of 32-kDa protein is in the size range of coat proteins of tenuiviruses [77, 78], it was suspected that the additional pathogen might be a tenuivirus [12, 40]. Purified preparations of the 32-kDa protein contained threadlike structures measuring 5 to 8 nm in diameter (Fig. 3e) [13] and resembling purified tenuivirus particles [77-79]. Thin-section electron microscopy of infected maize and wheat leaf specimens, however, revealed the consistent presence of DMPs structurally indistin-
Fig. 3 Cytopathic effects of High Plains disease (HPD) in corn and wheat, (a) Double membrane-bound particles (DMPs) and wheat streak mosaic virus (WSMV) inclusions in an HPDaffected wheat leaf cell. A large number of DMPs form clustered aggregates (arrowheads) throughout the cytoplasm, together with WSMV particles (W) and cylindrical inclusions that appear as pinwheels (P) and bundles (B). Mt = mitochondrion. Bar = 500 nm. (b) DMPs (arrowheads) and associated electron-dense amorphous inclusions (arrows) in the cytoplasm of com leaf cells. Proliferated rough endoplasmic reticulum (ER) is closely associated with the DMPs and inclusions. Mb = microbody. Bar = 300 nm. (c) DMPs (arrowheads) and associated amorphous inclusions (arrows) in the cytoplasm of a wheat cell affected with HPD, inmiunogold-labeled with disease-specific 32-kDa protein antiserum in situ. Since the cell was not osmicated during fixation, membranes of DMPs and other cellular organelles, such as the nucleus (N), mitochondria (Mt), and microbody (Mb), are not well preserved. Bar = 300 nm. (d) HPD-affected com leaf cells containing DMPs and flexuous rods of WSMV and immunogold-treated with WSMV-specific antisemm. Note that the gold particles specifically labeled the flexuous WSMV particles (arrows) but not the DMPs (arrowheads). Bar = 200 nm. (e) Threadlike stmctures (arrowheads), often circular or twisted or both, in the partially purified preparation from HPD-affected com. (The threads were stained with 2% uranyl acetate.) Bar = 100 nm. (f) Gold-labeled threadlike stmctures (arrows) following immunogold treatment with HPD-specific 32kDa protein antiserum. Bar =100 nm.
40
2.
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41
guishable from those associated with other eriophyid mite and DMP-associated diseases (Fig. 3a and b) discussed in this chapter, but not the tenuivirus-like structures or intracellular inclusion bodies common to infections by most tenuiviruses [80-82]. Furthermore, DMPs of HPD are also associated with proliferated endoplasmic reticulum and amorphous inclusions similar to those associated with DMPs of thistle mosaic (Fig. 3b) It should be noted that many leaf cells of severely symptomatic maize and wheat that contain DMPs also contain cylindrical or pinwheel inclusions and bundles of flexuous rod-shaped particles indicative of potyvirus infection (Fig. 3a). This suggests that these cells are doubly infected with DMPs and a potyvirus previously identified as WSMV [12, 83]. It was also noted that cells containing both DMPs and cylindrical inclusions exhibit more DMPs than those cells without cylindrical inclusions, often forming a number of clustered DMP aggregates (Fig. 3a). In cells without cylindrical inclusions the DMPs are usually scattered randomly in the cytoplasm (Fig. 3b) as in the case of cells affected by wheat spot mosaic, redbud yellow ringspot, or rose rosette disease [9, 11, 49]. Among DMP-associated diseases, it was noticed that wheat spot mosaic disease (WSpMD) is similar to HPD in symptomatology and host range [13]. The major foliar symptoms of HPD, chlorotic spots or white flecks, are also present in the same hosts infected with WSpMD. While chlorotic spots are the major symptoms of WSpMD, these spots in HPD are mixed with other symptoms, such as general chlorosis in a mosaic or stripe pattern, reddening, necrosis, and stunting [12,40, 83]. Because of the more severe and unusual symptom expression and the presence of an HPD-specific 32-kDa protein in symptomatic leaf extracts, it was thought that HPD was caused by a potentially new virus tentatively named High Plains virus [12, 76, 83]. The preliminary report that four to six species of doublestranded RNA are associated with the 32-kDa protein strengthens the case that the protein is indeed viral in origin [12, 76]. However, the infectivity of the fraction containing the protein and the double-stranded RNA has not been demonstrated. The severe symptomatology of HPD could reflect the synergistic effects of mixed virus infection rather than a new virus [13]. Severe diseases caused by mixed infections are not uncommon in many crops, including maize, in the field. Corn lethal necrosis, a well-known disease caused by a mixed infection with maize chlorotic mottle (MCMV) and either maize dwarf mosaic or WSMV is a good example [84]. Either virus alone results in a mild disease but when they coinfect corn, the disease is severe, and the titer of MCMV can be increased as much as fivefold by the synergistic interaction [85]. Cowpea stunt is also a very important disease caused by a mixed infection by two distinct viruses, cucumber mosaic cucumovirus and blackeye cowpea mosaic potyvirus [86]. Symptoms of cowpea stunt are extremely severe owing to the synergistic effects of the viruses compared with symptoms on cowpea infected singly with either of these two viruses. In cowpea stunt-affected leaves, the titer of cucumber mosaic virus is many times higher than that in leaves infected with cucumber mosaic virus alone.
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Many HPD-affected plants, either collected from the field or exposed to eriophyid mites collected from HPD-affected plants, were reported to also be infected with WSMV [12, 40, 87]. The wheat curl mite that transmits HPD agent is also known to transmit WSMV [18, 42]. Based on these facts and on the presence of both potyvirus-characteristic cylindrical inclusions and DMPs in most cells of HPD-affected plants, it is hypothesized that HPD is a result of a mixed infection with WSMV and the agent represented by DMPs [13]. Highly populated DMPs in cells that contain cylindrical inclusions may reflect such a synergism. It is understandable, however, that some of the plants with HPD symptoms in the field are infected singly with either of the two pathogens [87]. The foliar symptoms of chlorotic spots and the transmission of the disease agent by the same mite vector in both WSpMD and HPD, as well as the occurrence of structurally indistinguishable DMPs in both diseases, suggest that the DMPs associated with these two diseases are similar if not identical. In order to determine where the HPD-specific 32-kDa protein is located in HPD-affected cells and how this protein relates to DMPs, immunogold labeling studies were carried out using the antiserum prepared against purified 32-kDa protein [13]. The study revealed that the immunogold particles specifically labeled the DMPs and associated amorphous inclusions of HPD-affected cells (Fig. 3c). When the threadlike 5- to 8-nm structures in purified preparations containing 32-kDa protein were immunogold-labeled using the same antiserum, gold particles specifically labeled them (Fig. 3e). When HPD-affected com leaf cells containing DMPs and flexuous rods of WSMV particles were treated with WSMV antiserum, gold particles specifically labeled the flexuous rods (Fig. 3d) but not the DMPs or their associated inclusion bodies. Therfore, the DMPs and their associated inclusion bodies are not serologically related to WSMV These studies indicate that the DMPs, amorphous inclusions, and threadlike structures are made up, at least in part, of the disease-specific 32-kDa protein. It is postulated that the threadlike structures represent tightly packed structural components, such as nucleocapsids, which are released from ruptured DMPs during the purification process. Attempts to label the DMPs in cells of plants infected with rose rosette and thistle mosaic by using antiserum to the 32-kDa protein of HPD were unsuccessful.
///. Concluding Remarks The double membrane-bound, viruslike particles described in this chapter are unique when compared with any known plant viruses. They not only have a unique morphology but also show no structural similarities to known plant viruses. The only virus group that may show some degree of similarity to DMPs is that of the tomato spotted wilt tospoviruses (TSWV). Tospoviruses, viruses
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with enveloped particles, are transmitted by thrips [65, 88]. The particles of TSWV are, however, much smaller in size (85 nm in diameter) and much more homogeneous in shape than DMPs. The single-membrane envelope of TSWV particles is highly glycosylated and bears spiked surface structures [64]. In addition, most TSWV particles, unlike DMPs, always occur as clustered aggregates in C3^oplasmic membranous cisternae derived from rough endoplasmic reticulum [68]. However, double-enveloped particles, somewhat similar to DMPs, especially those associated with amorphous inclusions, have been reported to occur in cells in early stages of infection with TSWV [89]. Recently, it has been reported that double-enveloped TSWV particles occur more commonly together with single-enveloped mature particles in certain isolates [68, 69]. These particles are formed by budding singly from cisternal membranes of smooth endoplasmic reticulum, thus acquiring an extra membrane, or from flattened cisternal margins of Golgi bodies. These double-enveloped particles are interpreted as immature [89] or defective forms of virions [69]. It is unlikely that DMPs and TSWV particles are identical morphological entities, but the amorphous inclusions associated with DMPs in thistle mosaic and High Plains diseases are very similar to the amorphous or granular, dense-staining, viroplasm-like structures associated with TSWV particles. It has been suggested that TSWV particles arise directly from condensing clumps of viroplasmic material into nearby proliferated membranes, resulting in the formation of a budding configuration [64, 68, 89]. In fact, this proposed amorphous inclusions-viroplasm connection is further strengthened by immunospecific gold labeling studies. These studies indicate that the amorphous inclusions induced by impatiens necrotic spot tospovirus consist of viral nucleocapsid [68, 69]. It should also be mentioned that the DMPs and associated amorphous inclusions in thistle mosaic, as shown in Fig. le and f and Fig. 2b and c, closely resemble immature particles and viroplasms of the poxvirus group of animal viruses with double-stranded DNA genomes [72, 73]. Viroplasms of poxviruses also consist of viral nucleoprotein and are referred to as viral "factories" [90]. Poxviruses have not been reported to occur in plants. Structural similarities between these viroplasms, including those of phytoreoviruses, and amorphous inclusions associated with thistle mosaic DMPs [15] would certainly suggest that the latter are viroplasms. Some DMP-associated diseases share common cytopathic effects, which suggests that they are related to one another. Proliferation of intracellular membrane systems, especially of endoplasmic reticulum, is a prominent cytopathic feature in WSpMD [3, 41, 91], FMD [37, 38, 44, 48], ThMD [15], and HPD [13]. The association of tubular inclusions with DMPs is a consistent cytopathic feature of FMD isolates from different countries, including Italy, England, and the United States [15, 44], and of ThMD as well [15]. It is apparent that these tubular inclusions are not membranous since they do not exhibit a characteristic membrane configuration (lipid bilayer). Preliminary studies indicate the presence of nucleic acid and protein in tubular inclusions, suggesting that these structures might be involved in
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DMP production and assembly [44]. In ThMD, the tubular inclusions are structurally indistinguishable from those occuring together with amorphous inclusion bodies in FMD [15]. In fact, a large mass of amorphous inclusions often contains tightly packed tubular aggregates in its interior, suggesting that the compositional nature of the two is similar and that the latter is derived from the former. The association of a 32-kDa protein with HPD is generally accepted, and its antiserum is widely utilized as a diagnostic tool for the disease. The 32-kDa protein has also provided evidence that a "new virus," other than WSMV, is involved in causing HPD. Immunospecific labeling studies using the 32-kDa protein antiserum clearly demonstrate that the antigen of the new virus is located in DMPs and associated amorphous inclusions [13]. The question that arises is: Are DMPs virus particles? Unfortunately, we do not have conclusive evidence that DMPs represent virus, primarily because of our inability to purify the DMPs from diseased plants for further characterization. Numerous attempts using various purification protocols to purify DMPs from HPD or any other DMP-associated disease were unsuccessful. It appears that then* double membrane-bound structure makes it difficult to separate DMPs from host-cell membrane-bound organelles during purification (unpublished data). The presence of double-stranded RNAs (dsRNAs), a diagnostic feature of RNA virus infection in plants [92], has also been tested for a number of DMP-associated diseases. Although isolation of several species of dsRNA from HPD has been claimed [76], no dsRNA has been successfully isolated from any symptomatic yellow ringspot of redbud, rose rosette, thistle mosaic, or fig mosaic samples (unpublished data). This suggests that the DMPs might be viruses with DNA genomes or that the dsRNA isolation method might not be appropriate for these hosts, which contain unusually high levels of interfering materials such as polysaccharides and tannins. In summary, DMPs associated with eight eriophyid mite-borne diseases reviewed in this chapter are unique in morphology. Some of them occur with cytopathological inclusions considered to be viroplasms; that is, sites of virus replication and assembly. Immunospecific gold labeling of HPD-specific 32-kDa protein (apparently encoded by a "new virus" associated with HPD) on DMPs and associated amorphous inclusions support the hypothesis that the inclusions are indeed viroplasms. Based on these findings and other common characteristics of viral diseases, such as symptomatology and graft and mite vector transmissibility, it is theorized that the DMPs are viral in nature and are the causal agents of the diseases discussed here. The DMPs may represent the virions of a new group of plant viruses.
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84. Niblett, C.L., and Claflin, L.E. (1978). Com lethal necrosis—a new virus disease of corn in Kansas. Plant Dis. Rep. 62, 15-19. 85. Goldberg, K.B., and Brakke, M.K. (1987). Concentration of maize chlorotic mottle virus increased in mixed infections with maize dwarf mosaic virus, strain B. Phytopathology 71, 162-167. 86. Anderson, E.J., Kline, A.S., Kim, K.S., Goeke, S.C, and Albritton, C.W. (1994). Identification of cowpea stunt disease in south central Arkansas. Ark. Farm Res. 43, 14-15. 87. Mahmood, T., Hein, G.L., and Jensen, S.G. (1998). Mixed infection of hard red winter wheat with High Plains virus and wheat streak mosaic virus from wheat curl mites in Nebraska. Plant Dis. S2, 311-315. 88. Ananthakrishnan, T.N. (1980). Thrips. In "Vectors of Plant Pathogens" (K.F. Harris and K. Maramorosch, eds.), pp. 149-164. Academic Press, New York. 89. Milne, R.G. (1970). An electron microscope study of tomato spotted wilt virus in sections of infected cells and in negative stain preparations. J. Gen. Virol. 6,267-276. 90. Murphy, F.A. (1996). Virus taxonomy. In "Fields Vkology" (B.N. Field, D.M. Knife, and D.M. Howley, eds.), 3rd ed pp. 15-57. Lippincott-Raven Publishers, Philadelphia. 91. Chen, M.H., and Hiruki, C. (1990). The ultrastructure of double membrane-bound bodies and endoplasmic reticulum in serial sections of wheat spot mosaic-affected wheat plants. Proc. XII Int. Congr Electron Microsc. 3, 694. 92. Valverde, R.A., Nameth, S.T., and Jordan, R.L. (1990). Analysis of double-stranded RNA for plant virus diagnosis. Plant Dis. 74,255-258. 93. Oldfield, G.N. (1970). Mite transmission of plant viruses. Annu. Rev. Entomol. 15,343-380. 94. Slykhuis, J.T. (1953). The relation of Aceria tulipae (K) to streak mosaic and other chlorotic symptoms of wheat. Phytopathology 43,484^85. 95. Nauh, L.R., and Styer, W.E. (1969). The dispersal of Aceria tulipae and three other grass-infesting eriophyid mites in Ohio. Ann. Entomol. Soc. Am. 62, 1446-1455. 96. Slykhuis, J.T. (1982). Hordeum mosaic. In "Compendium of Barley Diseases" (D.E. Mathre, ed.), p. 52. Am. Phytopathol. Soc. Press, St. Paul, MN. 97. Slykhuis, J.T., and Bell, W. (1966). Differentiation of agropyron mosaic, wheat streak mosaic, and a hitherto unrecognized hordeum mosaic virus in Canada. Can. J. Bot. 44, 1191-1208. 98. Gill, C.C. (1971). Purification of oat necrotic mottle virus with silver nitrate as a clarifying agent. J. Gen. Virol. 12, 259-270. 99. Gill, C.C. (1976). "Oat necrotic mottle virus." C.M.I./A.A.B. Descriptions of Plant Viruses. No. 169. 100. Gill, C.C. (1980). Some properties of the protein and nucleic acid of oat necrotic mottle virus. Can. J. Plant Pathol 2, 86-89. 101. Milicic, D., Kujundzic, M., Wrischer, M., and Plavsic, B. (1980). A potyvirus isolated from Bromus mellis. Acta Bot. Croatica 39, 27—32. 102. Milicic, D., Manula, D., and Plazibat, M. (1982). Some properties of brome streak mosaic virus. Acta Bot. Croatica 41, 7-12. 103. VanDijk, P., Verbeck, M., and Bos, L. (1991). Mite-borne virus isolates from cultivated Allium species and their classification into two new rymoviruses. Netherlands J. Plant Pathol. 97, 381-399. 104. Jones, P (1980). Leaf mottling of Spartina species caused by a newly recognized virus, spartma mottle virus. Ann. Appl. Biol. 94, 77-81. 105. Ahmed, K.M., and Benigko, D.A. (1985). Virus-vector relationship in mosaic disease of garlic. Ind. Phytopathol. 38, 121-125.
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106. Oldfield, G.N., Creamer, R., Gispert, C , Osorio, R, Rodriguez, R., and Perring, T.M. (1995). Incidence and distribution of peach mosaic and its vector, Eriophyes insidious (AcariiEriophyidae) in Mexico. Plant Dis. 79, 186-189. 107. Amos, I , Haltton, R.G., Knight, R.C., and Massee, A.M. (1927). "Experiments in the Transmission of Reversion of Black Currants." Ann. Rep. East Mailing Res. Sta., Kent., 1925, II Suppl., pp.126-150.
CHAPTER 3
An Anatomical Perspective of Tospovirus Transmission T. NAGATA D. PETERS
/. Introduction Plant cell walls are a barrier to infection. To overcome this barrier, some plant viruses depend on vectors to move from infected plant to healthy plant. Viruses that are spread mechanically infect plants through small wounds in epidermal cells. Insects are the most important vectors. Several members of the Aphididae, Aleyrodidae, and Thripidae can efficiently transmit several plant viruses. The evolution of plant virus-vector relationships has resulted in a wealth of different transmission mechanisms, resulting in characteristic relationships specific for each taxonomic group of viruses. These relationships, vector biology, and cropping systems are the main determinants of plant virus spread and the kinetics thereof. In this chapter we describe what is known about the kinetics of tospovirus transmission and the relationship between these viruses and their thrips vectors. Members of the Tospovirus genus are the only plant-infecting viruses in the Bunyaviridae; members of the other four genera infect animal hosts. The viruses of three of the latter genera utilize bloodsucking arthropods such as mosquitoes, phlebotomine flies and ticks as vectors, whereas the tospoviruses are transmitted by phytophagous thrips [1]. The biology of bunyavirus vectors dictates that only certain developmental stages are able to spread viruses. Acquisition and transmission of mosquito-borne bunyaviruses are restricted to adults because the larvae live in water and do not feed on virus-infected hosts. Tick-borne bunyaviruses can be acquired by larvae or nymphs and transmitted over successive instars. An even Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
5 1
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more complex situation exists for tospoviruses. Only larvae acquire the virus, whereas larvae at the end of the larval development and adults transmit. Two developmental instars, the prepupa and pupa, do not feed and hence can neither acquire nor transmit virus. We will also analyze the transmission kinetics and molecular biology of virus transmission by thrips. Transmission kinetics will elucidate the transmission competence of individual thrips. Molecular investigations will elucidate the mechanisms governing virus entry into cells, the role of viral membrane glycoproteins in virus acquisition, virus replication, and movement through thrips.
//.
Tospovirus Morphology and Composition
Tomato spotted wilt virus (TSWV) is named the type member of the genus Tospovirus, which contains 12 other species. The structure, genome, and coding strategies of tospoviruses are characteristic of other genera of Bunyaviridae. The particles are roughly spherical and 80-110 nm in diameter. They enclose three nucleocapsids in a lipid membrane acquired from the host during maturation in the Golgi complex [2]. Two glycoproteins (Gl and G2) are anchored on this membrane. The genome consists of three RNA segments (L, M, and S), encapsidated by the nucleocapsid (N) protein forming the nucleocapsids. The L RNA segment, which has a negative polarity, encodes the RNA-dependent RNA polymerase associated with the nucleocapsids. The M and S RNA segments are ambisense. The M segment encodes the precursor for the Gl, G2, and movement proteins, whereas the S RNA codes for N protein and a nonstructural protein (NSs). The NSs protein and the movement protein, neither of which is assembled in virus particles, can be detected in infected plant and thrips cells [3-6]. The NSs occurs in paracrystalline inclusions [5]; the movement protein is associated with plasmodesmata of infected plants [6].
///. A.
Ttirips Vectors and Tospovirus Transmission
Thrips
Pittman [7] discovered that Thrips tabaci transmits TSWV It was believed for decades that this species was the main vector of TSWV, although other vector species were known [8]. However, the vector status of T tabaci has been challenged in recent years [9] and appears more complex than previously surmised [10, 11]. The cosmopolitan T tabaci seems the main vector of this virus in tobacco in Eastern Europe, whereas Frankliniella schultzei is considered the main vector in tobacco in the United States [12, 13]. Studies in Poland revealed that populations
3.
AN ANATOMICAL PERSPECTIVE OF TospoviRus TRANSMISSION Table I
53
Recognized Thrips Species Transmitting Tospoviruses
Vector species Franklinella bispinosa Efusca E intonsa E occidentalis E schultzei "darkform " E schultzei "lightform " Thrips palmi T setosus T tabaci
Tospovirus species transmitted TSWV INSyTSWV TSWV CSNV, GRSV, INSV, TCSV, TSWV CNSV, TSWV, GRSV, TCSV TSWV, TCSV GBNV, TSWV TSWV lYSV, TSWV
^ CSNV = chrysanthemum stem necrosis virus; GBNV = groundnut but necrosis virus; GRSV = groundnut ringspot virus; ESTSV = impatiens necrotic spot virus; lYSV = iris yellow spot virus; TCSV = tomato chlorotic spot virus; TSWV = tomato spotted wilt virus.
on tobacco crops differed from those infesting other plant species in their abihty to transmit TSWV This abihty could be correlated with the presence of males in the populations on tobacco and their absence on other plant species [10]. The thrips were taxonomically identical to T. tabaci. Zawirska [10] considered those on tobacco as T. tabaci subsp. tabaci [14] and those on other hosts as T. tabaci subsp. communis [15]. The communis type can be distinguished from the tabaci type by the presence of a comb on the dorsum of the ninth abdominal tergite. The incompetence of T. tabaci as a vector of TSWV has been explained by incompatibilities among thrips populations and the TSWV isolates used [16]. However, laboratory studies revealed that the presence or absence of males in the populations might have affected transmission efficiency. Three thelytokous populations (populations with only females) failed to transmit TSWV [11], whereas arrhenotokous populations isolated from bean and leek were poor transmitters [11, 17]. Thrips from populations isolated from tobacco appeared to be efficient vectors of TSWV with transmission rates between 50 and 60% [12, 18]. It is evident from these results that more attention must be given to the vector status of T. tabaci to explain such discrepancies in transmission of TSWV by this species. Since the discovery of T. tabaci as a tospovirus vector, seven other vector species have been identified (Table I). Frankliniella occidentalis, after its worldwide expansion, is presently thought to be the most important vector [9, 11, 19]. The second identified tospovirus species, impatiens necrotic spot virus (INSV), surfaced during this expansion [20, 21]. Frankliniella occidentalis is well known in the horticultural and floral industries for the spreading of TSWV and INSV [19]. It seems to play a limited role in the spread of tospoviruses in large field crops such as tobacco, sweet pepper, and tomato.
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The vector status of some other species is subject to discussion. One of them, Thrips flavus, has been reported as a vector of watermelon bud necrosis virus, a virus probably identical to groundnut bud necrosis virus [22]. Its identity as a vector needs further confirmation because this species can be readily confused with Thrips palmi [23]. The vector status of Scirtothrips dorsalis has also been questioned, because it also can be easily misidentified as T. palmi [23]. Moreover, S. dorsalis does not belong to the genera Thrips and Frankliniella, which include all other tospovirus vectors. The claimed transmission of zucchini lethal chlorosis virus by Frankliniella zucchini, a recently described new thrips species, has to be confirmed in as much as no results of transmission experiments accompanied the report [24]. B.
Kinetics of Tospovirus Transmission
Knowledge of the kinetics of tospovirus transmission by thrips has progressed considerably in the last decade. The observation of Bald and Samuel [25] and Linford [26] that TSWV could only be transmitted by adults when the virus was acquired by larvae was known for decades. Transmission of the virus by secondary-stage larvae was suggested by Sakimura [8] in 1963 but demonstrated only in the last decade. Approximately 60 to 80% of larvae transmitted the virus before pupation, when newborn Frankliniella occidentalis larvae acquired the Brazilian isolate BR-01 and transmitted as adults [27]. Extrapolating the latter to field situations, most thrips emerging on infected plants become viruliferous and can transmit before pupation. However, this ability is not important to tospovirus spread since larvae are wingless. Plant-to-plant virus spread can only occur when healthy plants touch the plants on which the larvae emerged. The main spread will be by infected, winged, adult thrips. Minimum latent periods, varying between 4 and 18 days, were reported for TSWV in studies using F occidentalis, F schultzei, and T tabaci [8]. Less variable data were found for F occidentalis in the transmission of TSWV and INSV in serial transfer experiments [27]. Mean latent periods (LP50) of approximately 8, 5, and 4 days were recorded at 20°, 24°, and 27°C for F occidentalis larvae fed as newborn larvae for 24 hours on TSWV-infected plants. Similar temperaturedependent values were found for INSV These LP50S were slightly longer for adults that transmitted after but not before pupation, excluding the prepupal and pupal period from these data. The minimum acquisition access periods (AAPj^in) foi" T tabaci [8] and F occidentalis [28] larvae fed on infected Datura stramonium plants are 15 and 5 minutes, respectively. An average of 8.2% of F occidentalis thrips transmitted virus after the 15-minute AAP; hence, it can be concluded that the actual AAPmin is even shorter. The successful acquisition of virus during these brief AAPs strongly suggests that virus is ingested from superficial plant cells and that small amounts of ingested virus suffice to initiate an infection in thrips. The AAP after which the
3.
AN ANATOMICAL PERSPECTIVE OF TOSPOVIRUS TRANSMISSION
5 5
maximal numbers of thrips were able to transmit was 21 hours; the mean AAP (AAP50) was approximately 1 hour. Similarly brief periods characterize the minimum inoculation feeding period (IFP). Given an IF? of 5 minutes 6% of viruliferous E occidentalis thrips transmitted virus to petunia and 17% to D. stramonium leaf disks. The transmission rate also depends on the host to be infected. Mean IFPs, of 1 or 2 hours were found by feeding viruliferous thrips on petunia or D. stramonium leaf disks, respectively [28]. The percentage of thrips that become viruliferous decreases rapidly with the age of the larvae when virus is acquired [29, 30]. The ability of one population to acquire virus declined so rapidly that its second-stage larvae did not become viruliferous [30]. Ingestion of virus by adults does not lead to transmission. This failure does not mean that the midgut epithelial cells do not become infected. A weak midgut infection was occasionally found in adult E occidentalis [29]. More severe infections in the midgut were found in Thrips setosus adults after feeding for 2 hours on infected source material [31]. These adults also did not transmit. Viruliferous adults may remain inoculative for life [32]. The virus is transmitted at a constant rate by thrips during this stage, although this rate is slightly lower in the first 1 or 2 days after emergence of the adults [17, 32]. Some authors reported that transmission ability is lost long before thrips die [8]. Survival, developmental time, and reproduction of E occidentalis are not significantly affected by TSWV replication [32]. However, negative effects were found when thrips were exposed to TSWV-infected tobacco plants during their whole larval development [33] or to INSV-infected Lobelia plants [34]. These results suggest that infected plants may have deleterious effects on vector physiology, providing an evolutionary advantage against thrips as compared with uninfected plants. Since the midgut and salivary glands can be extensively infected, some physiological stress is likely to occur. Phytophagous thrips that transmit tospoviruses often cause considerably feeding damage to plants. Females seem to cause more damage than males [35]. They are larger than males and produce about four eggs per day; hence, they need more food than males. Damage is mainly caused by feeding activities that drain groups of cells. Since it is difficult to reconcile infection site development with drained cells, inoculation may occur other than during food uptake. Three processes, stylet penetration, salivation, and food ingestion, have thus far been distinguished by electrically recording the feeding behavior of i^ occidentalis [36]. Virus may be introduced either during cell penetration or when saliva is excreted. Viruliferous males are reported to exhibit a higher transmission rate than females, which suggests that their feeding behavior may differ from that of females [35, 37]. This higher rate can tentatively be explained on the basis of either more frequent stylet penetrations (probings) or more salivation excretions at different locations. It is well established that populations of various plant virus vectors show differences in their efficiency to transmit viruses. Thirteen populations of E occi-
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T. NAGATA AND D. PETERS
dentalis varied considerably in their competence (20 to 89%) to transmit TSWV [38]. No populations were found in which all individuals were able to transmit. The differences in vector competence could not be correlated to any biological or genetic parameter.
IV. A.
Thrips as Tospovirus Hosts
Replication of Tospoviruses in Thrips
Considering the morphological similarity of TSWV to bunyaviruses, Milne and Francki [39] suggested that TSWV might replicate in thrips. The first claim that TSWV multiplied in its vector was made in 1987 by Cho et al [40]. They showed that the virus titer decreased between 4 and 10 days after acquisition but increased after 19 days. Conclusive evidence was obtained by Ullman et al. [3] and Wijkamp et al [4]. The former authors measured the titer of the nucleocapsid protein by enzyme-linked immunosorbent assay (ELISA) and analyzed the accumulation of both this protein and the NSs protein by Western blot analysis. The virus titer rapidly decreased in the first hours after a 2-hour AAP, followed by a sharp increase at 6 hours post-AAP. The virus titer exceeded the initial quantity acquired and reached maximum values just before pupation and after adult emergence. The lower virus titers during the pupal stages result from sloughing of the midgut epithelium along with the infecting virus [29]. The decrease in virus titer during ecdysis from first to second larval instar was more difficult to explain. The NSs protein can be readily detected in infected thrips by Western blotting [4]. Large amounts of this protein occur in the salivary glands, as shown by light microscopy [4], and in paracrystalline structures in the midgut epithelial cells of viruliferous thrips, as shown by electron microscopy [3]. Synthesis of this protein and its accumulation in fibrous paracrystalline crystals provide strong evidence of TSWV replication in thrips, since NSs is produced after transcription of its mRNA from the viral complementary S RNA strand [41]. Replication of TSWV in thrips received ftirther confirmation in 1997 when titers of both the N and NSs proteins were shown to increase over time in TSWV-inoculated E occidentalis cultured in vitro [42]. B.
Midgut Infection by Tospoviruses
Since thrips are infected by and can subsequently transmit tospoviruses, it is evident that the virus moves through the thrips body in a stepwise fashion. A simplified pathway of TSWV circulation in thrips can be envisioned in several steps. The first step, entry of the virus into midgut epithelial cells, may lead to infection in these cells. Following replication, the virus must escape from the midgut either
3.
AN ANATOMICAL PERSPECTIVE OF TOSPOVIRUS TRANSMISSION
5 7
to the salivary glands or to other organs and tissues in the hemocoel that become infected. Entry of virus into salivary gland cells results in the infection, movement of virus into the salivary ducts, and its transmission in virus-laden saliva secreted during feeding. Infection of thrips by tospoviruses presumably starts with the attachment of virus particles to the surface of midgut epithelial cells. Receptor-mediated binding is the most frequently encountered process in cell recognition by membranebound viruses. The presence of the glycoprotein envelope in TSWV particles appears essential to infection of midgut cells. Primary infections can not be established by a morphologically deficient isolate of TSWV when it is ingested by thrips from infected plants [43]. This deficient isolate also lacks the ability to infect E occidentalis cells in tissue culture. It appears, therefore, that either one or both glycoproteins associated with the envelope are required for successful virus entry into midgut cells. Furthermore, the existence of a specific motif in one of these proteins seems likely. The amino acid sequence argine-glycine-aspartic acid in the G2 protein might play a role in the attachment of virus to thrips cells; this is a motif known to be involved in the attachment of foot-and-mouth disease virus to cells [44]. Efforts are underway to identify possible cellular receptors for TSWV in thrips. Bandla et ah [45] reported the detection and isolation from midgut cells of a 50kDa protein that has affinity for TSWV glycoproteins in an overlay assay. This protein was absent in other thrips tissues. In the same study, separate thrips organs were ground, gel-separated, blotted onto a membrane, and overlaid with a purified virus preparation. The 50-kDa protein had an affinity for virus particles or their glycoproteins, as shown by immunolabeling that targeted bound virus particles. Another thrips protein of 94 kDa may also play a role in the infection process. This protein was detected throughout the whole body in all developmental stages of E occidentalis and T. tabaci [46] and was consistently present in continuous cell cultures of 7^ occidentalis (T Nagata, unpublished data). However, this protein could not be detected in the midguts of these species. The 94-kDa protein was also present in adults of Parthenothrips dracenae, a nonvector thrips species, but absent in the aphid Myzus persicae. These results suggest that this particular protein does not play a role in virus entry into the midgut. Its functioning in the transport of virus through the body can not be ruled out, however, since G2 protein appeared to be the viral determinant in the reaction with this 94-kDa protein. The plasmalemma of the midgut epithelial cells represents the first barrier to be passed by the virus after binding to the cells. A specific binding of the virus is a crucial prelude for penetration of the minimal number of virus particles needed to infect the cell. Two strategies have been proposed to explain the entry of other arthropod-borne viruses (arboviruses) into mesenteron cells of mosquitoes. The viruses may fiise with the membrane, with subsequent direct release of the nucleocapsids into the cytoplasm or alternatively, the virus may enter the cells by endocytosis, a process in which the complete particles are engulfed in an endosome by the
58
T. NAGATA AND D. PETERS
plasmalemma. The nucleocapsids will then be released from these vesicles into the cytoplasm by a mildly low pH-mediated step. Most members of the Bunyaviridae are known to use this latter strategy to enter and infect animal cells. The strategy of TSWV entry and the site at which infection is initiated has yet to be elucidated. The gut environment may strongly influence virus attachment and entry into the midgut. This environment actively digests ingested material, a process that may be detrimental to the virus. Protection of the virus by simultaneously ingested fresh plant material in the midgut lumen might explain compared to the posterior regions of the gut why the virus can successfully infect the midgut [29]. This preferential infection site might also be explained by a restricted presence of the viral receptor on the midgut cells only. The midgut environment is not inevitably detrimental to the virus. Structural changes in viral envelope proteins, induced by a low pH in the midgut, have been shown to initiate membrane fusion and formation of endosomes for La Crosse virus and California encephalitis virus, both bunyaviruses [47]. C.
Significance of Midgut Infection
The alimentary canal of thrips consists of foregut, midgut, and hindgut. The midgut is divided by two loops forming three regions, designated 1, 2 and 3 [48, 49] (Fig. 1). Two types of muscle tissue surround the midgut. One type lines the midgut in a parallel and the other in a circular fashion. The first signs of virus infection are observed within 12 to 24 hours after acquisition in the epithelial cells of the first midgut region (Fig. 2A, see color insert). The infection expands to the midgut muscle cells of this region and slowly to the muscle cells of the second region and finally the third region. The longitudinal muscles are infected first, followed by the circular muscles that overlie the longitudinal ones. This route of infection through the visceral muscle may occur via cell-to-cell movement of mature virus particles. Such particles were observed in the cytoplasm of visceral muscle cells in second-instar larvae. No mature particles could be discerned in these cells in adults [29]. Aggressive cell renewal occurs in many tissues during pupation. Gut epithelial cells are sloughed, including virus-infected ones. No signs of reinfection of adult midgut epithelium cells have been observed by light and electron microscopy after emergence from the pupal stage. The foregut, which does not succumb to infection in larvae, appears to be infected after pupation (Fig. 2B, see color insert). The degree of the midgut infection depends qualitatively and quantitatively on the length of the AAP and the age of the larva acquiring the virus. The midguts of newborn larvae given an AAP of 16 hours become more widely infected than do those of similar larvae fed for 3 hours. The age of the thrips when virus is ingested has an even more pronounced effect on the degree of midgut infection (Table II). The capacity to transmit is also positively correlated with the duration of the AAPs and the degree to which the midgut becomes infected [29, 30].
Fig. 2 Laser scanning microscope images of tomato spotted wilt virus infected digestive system of Frankliniella occidentalis using FITC-labeled nucleocapsid protein antibodies. Intestinal tract and salivary glands of larvae 72 hours postacquisition (A) and of transmitting adult thrips (B). Notice in A that most of the midgut epithelium is infected and the muscle cells beyond the infected epithelial cells are already infected. In Fig. B, the muscle infection in Mgl and Mg2 can clearly be observed. A completely infected salivary gland lobe and partially infected lobe are shown in C. An infected ligament connects the infected midgut with an infected salivary gland (D). Mgl = anterior midgut; Mg2 = intermediate midgut; SG = salivary glands; Fg = foregut; Lg = ligament.
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3.
59
A N ANATOMICAL PERSPECTIVE OF TOSPOVIRUS TRANSMISSION
i^^^Blft
^BB^Bl
^^^iliiiiiiii
iftHlRS
lliiiigli HiKsMiiBiii
Fig. 1 Composite drawing of the alimentary tract and associated organs of the thrips Hercinothrips femoralis. Reprinted (as modified by T. Nagata) through the courtesy of Dr. H. Moritz
Not all thrips will transmit, although the midgut of all individuals is infected when they acquire virus as newborn larvae. Thus, not all thrips with midgut infections become transmitters, and the percentage of thrips tospvirus-positive in ELISA is always higher than the percentage of transmitters [27, 35, 37]. These observations show that other factors also play a role in converting thrips into transmitters. These factors, which have still to be elucidated, may either prevent infection of the salivary glands or prevent virus from reaching infectious levels in the glands or salivary ducts. Existence of thresholds has been reported for several arboviruses transmitted by different mosquito species [50], The threshold to infect the mesenteron seems to be dose-dependent. Existence of a dose-dependent threshold could also be demonstrated for thrips by using defective interfering (DI) isolates characterized by lower
60
T. NAGATA AND D. PETERS
Table II The Relationships among the Age at Which Thrips Acquire Virus, the Transmission Rate, and the Degree of Midgut Infection [29]
Age when fed on virus source 0- to 4-h-old larvae (1 st instar) 72- to 76-h-old larvae (2nd instar) 1-to 2-day-old adults
Percentage of adults transmitting 47 12 0
Percentage of thrips with infected midguts 100 35 2
numbers of replicating units in inocula. Nagata et al [43] showed that transmission efficiency corresponded with the degree to which the midgut becomes infected. The transmission of one isolate, infecting the midgut to the same degree as wild-type virus but at a lower rate, was not considerably impeded [43]. Using a nontransmissible DI isolate resulted in a limited infection of the midgut with respect to the number of infected sites as well as the severity of infection. The failure of some T. tabaci populations to transmit TSWV appeared not to be limited by a midgut infection barrier but by a low rate of virus replication in the epithelial cells. Infection in these thrips was almost eliminated by pupation, demonstrating that the infection did not expand to other tissues but remained restricted to the midgut epithelium. The competence of these thrips populations to replicate TSWV could also be demonstrated after inoculating cell cultures of these thrips populations [42]. As shown, midgut infection of R occidentalis does not always result in infection of the salivary glands, which explains the failure to transmit the virus [29, 43]. Histological studies using E occidentalis revealed that both lobes of the glands are severely infected in most transmitters (Fig. 2C). These lobes were only infected in a few nontransmitters. Salivary gland infection is usually restricted to a limited area in one of the lobes or completely absent in the large majority of nontransmitters [29]. No infection could be discerned in the salivary glands of any adult of nontransmitting T. tabaci populations [51]. These histological studies showed that the virus can severely infect the salivary glands before pupation when E occidentalis larvae ingest virus at an early stage of their life. Ingesting virus later in larval development results in limited infection of the salivary glands. These results indicate that the salivary glands have to be infected before pupation if a thrips is to become a transmitter. Thrips that do not transmit as larvae but do as adults may have become viruliferous before pupation, the period between becoming viruliferous and pupation being too brief for larval transmission to occur. D.
Infection of Salivary Glands
Dissemination of TSWV or its movement from the midgut to the salivary glands is the most baffling process in the infection of thrips. The mechanism
3.
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61
by which the virus invades and infects this organ is still not well understood. Movement of arboviruses in mosquitoes has been proposed to occur by release of virus from the mesenteron into the hemocoel and subsequent infection of the salivary glands [50]. Alternatively, the occurrence of large amounts of virus in neural tissue suggests a neural pathway for virus dissemination [52-55]. The results of electron microscopic studies do not support either of these pathways as possible routes for tospovirus passage through thrips [9, 29]. No clear evidence has been obtained for the presence of virus particles or viral protein aggregates in the hemocoel. A low virus concentration in the hemocoel might hinder particle observation by electron microscopy, as was concluded from studies with several other circulative plant viruses in their vector. The failure to find virus particles in the hemocoel does not exclude it as a possible route for TSWV circulation in the vector. Microinjection of TSWV into the hemocoel of more than 600 adult thrips did not result in salivary gland infection or render the recipient thrips inoculative. However, this failure could be attributable to technical difficulties encountered during microinjection (Nagata, unpublished data) and not necessarily to the inability of the hemocoel to serve as a viral passageway to the salivary gland system. In a recent study, another possible pathway for TSWV dissemination from midgut to the salivary glands has been suggested [29]. Viral antigens were detected by immunofluorescence in thin, threadlike structures or ligaments connecting the midgut with the salivary glands (Fig. 2D). These signs of infection are readily detectable in the glands before any infection of the gland cells is apparent. The first infections in the salivary glands were observed in those gland cells at the point of contact between the ligaments and the glands. The function and tissue type of these ligaments are still unknown. The canal of the tubular salivary glands was proposed as a third possible pathway for virus transport from gut to salivary system [49]. This organ is considered homologous to the accessory salivary glands of other insects. As proposed, this tubular organ ends in the midgut lumen, providing an avenue for virus migration to the salivary glands. However, infections have not been detected in these glands [29], and furthermore, their distal ends appear closed (i.e., not open to the midgut lumen) [56]. The presence of complete virus particles in the salivary glands has been unequivocally demonstrated [4]. They are present both in saliva vesicles and ducts. The presence of virus particles in saliva vesicles is indicative of intracellular maturation; their occurrence in ducts suggests maturation of virus at the apical plasmalemma of gland cells. TSWV may thus use two maturation strategies in the salivary glands, as has been suggested for Rift Valley fever virus, which may mature both at the cell surface and in cisternae of the Golgi complex [57]. Maturation of the tospovirus particles at saliva vesicle membranes and their subsequent release with the saliva into the ducts is more likely and requires only one strategy, namely, maturation at the membranes of the Golgi complex.
62
E.
T. NAGATA AND D. PETERS
Function of Viral Proteins in Infected Thrips
Infection of the midgut, the midgut muscle cells, and the foregut, ligament and salivary glands could be elucidated by histological studies using light microscopy. Infection of these tissues has mainly been demonstrated by using fluorescent-labeled antibodies to the N protein [29]. These studies revealed the development of infection with time but were not able to clarify the infection pathway. Our limited knowledge of the viral pathway in thrips is mainly due to the limited success of electron microscopic studies in detecting virus particles in infected tissues. In addition to N protein, accumulation of the NSs protein has been convincingly demonstrated in midgut epithelial cells by electron microscopy [3] and in salivary gland cells by light microscopy [4]. This protein accumulates in large paracrystalline structures in both plant and insect cells. The role of this protein in plant and thrips infection or in viral replication is not known. Since NSs occurs in large amounts in the salivary gland, it has been suggested as playing a role in the transmission of virus to plants [4]. The manifest accumulation of N and NSs proteins in various tissues is the result of an active replication of the S RNA segment and translation of its mRNAs. Accumulation of complete particles has only been observed in the salivary glands of 7^ occidentalis [4]. Occasionally, a few particles were discerned in the apical part of midgut epithelial cells and visceral muscle cells [51]. The finding of a limited number of complete particles indicates that the synthesis of G proteins (or the maturation of complete particles) is severely impaired during infection of thrips cells. This conclusion may be supported by a study on the compartmentalization and intracellular transport of TSWV proteins in midgut epithelial cells [58]. Both glycoproteins were found in trace amounts in amorphous inclusions and at membrane structures thought to be part of the Golgi complex. However, complete particles were not apparent [58]. Other observations also point to a limited synthesis of the G proteins in thrips [59]. These proteins were not detectable by ELISA in single thrips that were positive for the N protein or in more concentrated thrips extracts (25 thrips pooled from an N-positive population). In a study on the expression and cellular manifestation of the movement protein, small amounts could be detected in the cytoplasm of midgut epithelium in E occidentalis larvae 5 days postacquisition and in amorphous inclusion bodies in the salivary glands of adults [60]. This protein plays a role in cell-to-cell movement of viruses in plants via plasmodesmata. There is circumstantial evidence that tospoviruses are transported in plants not as complete virus particles but as nucleocapsids [61]. Since membrane-bound viruses are released by budding or exocytosis at the plasmalemma of animal and insect cells, it is not expected that this protein plays a role in the spread of tospoviruses in thrips. However, this protein does bind to nucleocapsids in infected plant cells during
3.
AN ANATOMICAL PERSPECTIVE OF TOSPOVIRUS TRANSMISSION
63
their transport to plasmodesmata [6, 62]. This association of nucleocapsids with movement protein may prevent or suppress maturation of virus particles, as shown in plant cells. Expression of the movement protein in thrips may also suppress mature particle formation and explain the rarity of complete particles in midgut epithelial cells. As a consequence, TSWV may move between some thrips tissues in the form of nucleocapsids.
y. Concluding Remarks A role for various barriers (e.g., entry into and release from the midgut and salivary gland barriers) has been proposed to describe the infection of vectors by arboviruses and persistently transmitted plant viruses [49, 63, 64]. These same barriers exist in the infection of thrips by tospoviruses, even though the infection pathway is not yet fully elucidated. Infection of the midgut appears to depend on the age of thrips, the virus dose acquired, and the replication rate in the midgut. Midgut infection has to surpass a certain level to successfully infect the salivary glands and render the virus-exposed thrips a transmitter. Conversion of thrips to transmitters requires that the salivary glands be heavily infected before pupation. Thrips with mildy infected salivary glands prior to pupation do not transmit virus as adults. It is an enigma why an initial salivary gland infection in adults does not render them transmitters. This raises the question of whether the salivary glands lose their potential for tospovirus replication during pupation of the thrips or after adult emergence.
References 1. Mound, L.A. (1997). Biological diversity. In "Thrips as Crop Pests" (T. Lewis, ed.), pp. 197-215. CAB International, Wallingford, UK. 2. Kikkert, M., Van Lent, J., Storms, M., Bodegom, P., Kormelink, R., and Goldbach, R. (1999). Tomato spotted wilt virus particle morphogenesis in plant cells. J. Virol. 73, 2288-2297. 3. Ullman, D.E., German, T.L., Sherwood, J.L., Westcot, D.M., and Cantone, F.A. (1993). Tospovirus replication in insect vector cells: Immunoc3^ochemical evidence that the nonstructural protein encoded by the S RNA of tomato spotted wilt tospovirus is present in thrips vector cells. Phytopathology 83,456^63. 4. Wijkamp, L, Van Lent, J., Kormelink, R., Goldbach, R., and Peters, D. (1993). Multiplication of tomato spotted wilt virus in its vector, Frankliniella occidentalis. J. Gen. Virol. 74, 341-349. 5. Kitajima, E., de Avila, A.C., Resende, R. de O., Goldbach, R., and Peters, D. (1992). Comparative cytological and immunolabelling studies on different isolates of tomato spotted wilt virus. J. Submicrosc. Cytol. Pathol. 24, 1-14.
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6. Storms, M.M., Kormelink, R., Peters, D., Van Lent, J.W., and Goldbach, R. (1995). The nonstructural NSm protein of tomato spotted wilt virus induces tubular structures in plant and thrips cells. Virology 214,485-493. 7. Pittman, H.A. (1927). Spotted wilt of tomatoes. Preliminary note concerning the transmission of the spotted wilt of tomatoes by an insect vector (Thrips tabaci Lind.). J. Counc. Sci. Ind. Res. 1, 74-77. 8. Sakimura, S. (1963). The present status of thrips-borne viruses. In "Biological Transmission of Disease Agents" (K. Maramorosch, ed.), pp. 3 3 ^ 0 . Academic Press, New York. 9. UUman, D.E., Sherwood, J.L., and German, T.L. (1997). Thrips as vectors of plant pathogens. In "Thrips as Crop Pests" (T. Lewis, ed.), pp. 539-565. CAB International, Wallingford, U.K. 10. Zawirska, L (1976). Untersuchungen iiber zwei biologische Typen von Thrips tabaci Lind. (Thysanoptera, Thripidae) in der VR Polen. Arch. Phytopathol. Pflanzenschutz 12, 411^22. 11. Wijkamp, L, Almarza, N., Goldbach, R., and Peters, D. (1995). Distinct levels of specificity in thrips transmission of tospoviruses. Phytopathology 85, 1069-1074. 12. Chatzivassiliou, E.K., Katis, N.I., and Peters, D. (1998). Transmission of tomato spotted wilt virus (TSWV) by Thrips tabaci grown on tobacco and non-tobacco crops. "Proceedings of the 4th International Symposium on Tospoviruses and Thrips in Floral and Vegetable Crops," pp. 59-62. Wageningen, The Netherlands. 13. McPherson, R.M., Pappu, H.R., and Jones D.C. (1999). Occurrence of five thrips species in fluecured tobacco and impact on spotted wilt disease in Georgia. Plant Dis. 83,765-767. 14. Lindemann, K. (1888). Die schadlichsten Insekten des Tabaks. Byull. Mosk Sch. Ispyt. Prir. NS2, 10-7. 15. Uzel, H. (1895). "Monographic der Ordnung Thysanoptera." Koniggratz. 16. Paliwal, Y.C. (1956). Some characteristics of the thrips vector relationships of tomato spotted wih virus in Canada. Can. J. Bot. 54,402^05. 17. Chatzivassiliou, E.K., Nagata, T, Katis, N.I., and Peters, D. (1999). Transmission of tomato spotted wilt tospovirus by Thrips tabaci populations originating from leek. Plant Pathol 48, 700-706. 18. Karadjova, O., and Hristova, D. (1998). Transmission of tomato spotted wilt virus by a Bulgarian population of Frankliniella occidentalis Perg. and Thrips tabaci Lind. "Proceedings of the 4th International Symposium on Tospoviruses and Thrips in Floral and Vegetable Crops," pp. 95-96. Wageningen, The Netherlands. 19. Daughtrey, M.L., Jones, R.K., Moyer, J.W., Daub, M.E., and Baker, J.R. (1997). Tospoviruses strike the greenhouse industry: INSV has become a major pathogen on flower crops. Plant. Dis. 81,1220-1230. 20. Law, M.D., and Moyer, J.W. (1990). A tomato spotted wilt-like virus with a serologically distinct N protein. J. Gen Virol. 72, 2597-2601. 21. Avila, A.C. de. Huguenot, C , Resende, R. de O., Kitajima, E.W., Goldbach, R.W., and Peters, D. (1990). Characterization of a distinct isolate of tomato spotted wilt virus (TSWV) from Impatiens sp. in the Netherlands. J. Phytopathol. 134, 133-151. 22. Singh, S.J., and Krishnareddy, M. (1996). Watermelon bud necrosis: A new tospovirus disease. Acta Hort. 431, 6S-n. 23. Mound, L.A. (1996). The Thysanoptera vector species of tospoviruses. Acta Hort. 431, 298-307. 24. Nakahara, S., and Monteiro, R.C. (1999). Frankliniella zucchini (Thysanoptera: Thripidae), a new species and vector of tospovirus in Brazil. Proc. Entomol. Soc. Wash. 101, 290-294. 25. Bald, J.G., and Samuel, G. (1931). Investigation on "spotted wilt" of tomatoes. II. Austr Commonwealth Council Sci. Ind. Res. Bull. No 54.
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26. Linford, M.B. (1932). Transmission of the pineapple yellow-spot virus by Thrips tabaci. Phytopathology 22,301-324. 27. Wijkamp, I., and Peters, D. (1993). Determination of the median latent period of two tospoviruses in Frankliniella occidentalis using a novel leaf disk assay. Phytopathology 83,986-991. 28. Wijkamp, L, van de Wetering, R, Goldbach, R., and Peters, D. (1996). Transmission of tomato spotted wilt virus by Frankliniella occidentalis: Median acquisition and inoculation access period. Ann. Appl. Biol 129, 303-313. 29. Nagata, T, Inoue-Nagata, A.K., Smid, H., Goldbach, R., and Peters, D. (1999). Tissue tropism related to vector competence of Frankliniella occidentalis for tomato spotted wilt virus. J. Gen. F/ro/. 80,507-515. 30. Van de Wetering, E, Goldbach, R., and Peters, D. (1996). Tomato spotted wilt tospovirus ingestion by first instar larvae oi Frankliniella occidentalis is a prerequisite for transmission. Phytopathology %6, 900-905. 31. Ohnishi, X, Hosokawa, D., Fujisawa, I., and Tsuda, S. (1998). Tomato spotted wilt tospovirus movement into salivary glands during pupation of the thrips vector, Thrips setosus, is associated with the transmissibility. "Proceedings of the 4th International Symposium on Tospoviruses and Thrips in Floral and Vegetable Crops," pp. 51-53. Wageningen, The Netherlands. 32. Wijkamp, I., Goldbach, R., and Peters, D. (1996). Propagation of tomato spotted wilt virus in Frankliniella occidentalis does neither result in pathological effects nor in transovarial passage of the virus. Entomol. Exp. Appl. 81,285-292. 33. Robb, K.L. (1989). Analysis of Frankliniella occidentalis (Pergande) as a pest of floricultural crops in California greenhouses. Ph.D. Dissertation, University of California, Riverside. 34. DeAngelis, J.D., Sether, DM., and Rossignol, P.A. (1993). Survival, development, and reproduction in western flower thrips (Thysanoptera: Thripidae) exposed to impatiens necrotic spot virus. Environ. Entomol. 34, 1308-1312. 35. Van de Wetering, E, Hulshof, J., Posthuma, K., Harrewijn, P, Goldbach, R., and Peters, D (1998). Distinct feeding behaviour between sexes of Frankliniella occidentalis results in higher scar production and lower tospovirus transmission by females. Entomol. Exp. Appl. 88, 9-15. 36. Harrewijn, P., Tjallingii, W.F., and Mollema, C. (1996). Electrical recording of plant penetration by western flower thrips. Entomol. Exp. Appl. 79, 345-353. 37. Sakurai, T., Murai, T, Maeda, T, and Tsumuki, H. (1998). Sexual differences in transmission and accumulation of tomato spotted wilt virus in its vector Frankliniella occidentalis (Thysanoptera: Thripidae). Appl. Entomol. Zool. 33, 583-588. 38. Van de Wetering, E, van der Hoek, M., Goldbach, R., Mollema, C , and Peters, D. (1999). Variation in tospovirus transmission between populations of Frankliniella occidentalis (Thysanoptera: Thripidae). Bull. Entomol. Res. 89, 579-588. 39. Milne, R.G., and Francki, R.I.B. (1984). Should tomato spotted wilt vims be considered as a possible member of the family Bunyaviridae? Intervirology 22, 72-76. 40. Cho, J. J., Mitchell, W C , Hamasaki, R.T., and Gonsalves, D. (1987). Detection of tomato spotted wilt virus (TSWV) in individual thrips in an ELISA. Phytopathology 11, 895. 41. Kormelink, R., Kitajima, E.W, de Haan, P, Zuidema, D., Peters, D , and Goldbach, R. (1991). The nonstructural protein (NSs) encoded by the ambisense S RNA segment of tomato spotted wilt virus is associated with fibrous structures in infected plants. Virology 181,459^68. 42. Nagata, T, Storms, M.M.H., Goldbach, R., and Peters, D. (1997). Multiplication of tomato spotted wilt virus in primary cell cultures derived from two thrips species. Virus Res. 49, 59-66. 43. Nagata, T, Inoue-Nagata, A.K., Prins, M., Goldbach, R., and Peters, D. (2000). Impeded thrips transmission of defective tomato spotted wilt virus isolates. Phytopathology 90, 454-459.
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44. Fox, G., Parry, N., Bamett, P, McGinn, B., Rowlands, D., and Brown, E (1989). The cell attachment site of foot-and-mouth disease virus includes the amino acid sequence RGD (arginineglycine-aspartic acid). J. Gen. Virol. 70, 625-637. 45. Bandla, M.D., Campbell, L.R., Ullman, D.E., and Sherwood, J.L. (1998). Interaction of tomato spotted wilt tospovirus (TSWV) glycoproteins with a thrips midgut protein, a potential cellular receptor for TSWV Phytopathology 88, 98-104. 46. Kikkert, M., Meurs, C , Van de Wetering, R, Dorftniiller, S., Peters, D. Kormelink, R., and Goldbach, R. (1998). Binding of tomato spotted wilt virus to a 94-kDa thrips protein. Phytopathology 88, 63-69. 47. Hacker, J.K., and Hardy J.L. (1997). Adsorptive endocytosis of California encephalitis virus into mosquito and mammalian cells: A role for Gl. Virology 235,40^7. 48. Moritz, G. (1997). Structure, growth and development. In "Thrips as Crop Pests" (T. Lewis, ed.), pp. 15-63. CAB International, Wallingford, UK. 49. Ullman, D.E., Westcot, D.M., Hunter, W.B., and Mau, R.F.L. (1989). Internal anatomy and morphology of Frankliniella occidentalis (Pergande) (Thysanoptera: Thripidae) with special reference to interactions between thrips and tomato spotted wilt virus. Int. J. Insect Morphol Embryol. 18,289-310. 50. Hardy, J.L. (1988). Susceptibility and resistance of vector mosquitoes. In "The Arboviruses: Epidemic and Ecology" (T.P Monath, ed.), pp. 87-126. CRC Press, Boca Raton, FL. 51. Nagata, T. (1999). The factors determining vector competence and specificity for tomato spotted wilt virus transmission. In "Competence and Specificity of Thrips in the Transmission of Tomato Spotted Wilt Virus," Thesis, Wageningen Agricultural University, pp. 57-71. 52. Chamberlain, R.W. (1968). Arboviruses, the arthropod-borne animal viruses. Curr. Top. Microbiol. Immunol. 4, 38-58. 53. Linthicum, K.J., Piatt, K., Myint, K.S., Lerdthusnee, K., Innis, B.L., and Vaughn, D.W. (1996). Dengue 3 virus distribution in the mosquito Aedes aegypti: An immunocytochemical study. Med. Vet. Entomol. 10, 87-92. 54. Miles, J.A.R., Pillai, J.S., and Maguire, T. (1973). Multiplication of Whataroa virus in mosquitoes. J. Med Entomol. 10, 176-185. 55. Murphy, FA., Whitfield, S.G., Sudia, W.D., and Chamberlain, R.W. (1975). Interactions of vector with vertebrate pathogenic viruses. In "Invertebrate Immunity" (K. Maramorosch and R.E. Shope, eds), pp. 2 5 ^ 8 . Academic Press, New York. 56. Del Bene, G., Dallai, R., and Marchini, D. (1991). Ultrastructure of the midgut and the adhering tubular salivary glands oiFrankliniella occidentalis (Pergande) (Thysanoptera: Thripidae). Int. J. Insect Morphol. Embryol. 20, 15-24. 57. Anderson Jr, G.W., and Smith, J.F. (1987). Immunoelectron microscopy of Rift Valley fever viral morphogenesis in primary rat hepathocytes. Virology 161, 91-100. 58. Ullman, D.E., Westcot, D.M., Chenault, K.D., Sherwood, J.L., German, T.L., Bandla, M.D., Cantone, FA., and Duer, H. (1995). Compartmentalization, intracellular transport, and autophagy of tomato spotted wilt tospovirus proteins in infected thrips cells. Phytopathology 85, 644-654. 59. Roggero, P., Ogliara, P., Ramasso, E., Arzone, A., Tavella, L., and Alma, A. (1996). Detection by TAS-ELISA of tomato spotted wilt virus nucleocapsid and Gl glycoprotein in Frankliniella occidentalis. Acta Hort. 431, 333-340. 60. Storms, M.M.H. (1998). Expression of the movement protein of tomato spotted wilt virus in its vector Frankliniella occidentalis. In "The role of NSm during tomato spotted wilt virus infection". Thesis, Wageningen Agricultural University, pp. 87-95. 61. Resende, R. de O., de Haan, P, de Avila, A.C., Kitajima, E.W, Kormelink, H.R., Goldbach, R., and Peters. D. (1991). Generation of envelope and defective interfering RNA mutants of tomato spotted wilt virus by mechanical passage. J. Gen. Virol. 72, 2375-2383.
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62. Kormelink, R., Storms, M., van Lent, J., Peters, D., and Goldbach, R. (1994). Expression and subcellular location of the NSm protein of tomato spotted wilt virus (TSWV), a putative viral movement protein. Virology 200, 56-65. 63. Hardy, J.L., Houk, E.X, Kramer, L.D., and Reeves, W.C. (1983). Intrinsic factors affecting vector competence of mosquitoes for arboviruses. Annu. Rev. Entomol. 28,229-262. 64. Ammar, E-D., and Nault, L.R. (1985). Assembly and accumulation sites of maize mosaic virus in its planthopper vector. Intervirology 24, 3 3 ^ 1 .
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CHAPTER 4
Analysis of Circulative Transmission by Electrical Penetration Graphs W. FRED. TJALLINGII ERNESTO PRADO
/. Introduction Stylet penetration activities form the major causes for the acquisition and inoculation of plant viruses. Understanding the responsible activities is important in evaluating the possibilities of new plant varieties resistant either to the virus vectors or to the viruses themselves. One cannot predict whether plant resistance to the vector will reduce virus spread in the field or whether plant resistance to the virus will affect vector host specificity. The use of electrical systems for monitoring insect probing and feeding has elucidated the processes of plant penetration by insects, especially those of aphids [1,2]. The details both of insect activities and of plant reactions to these activities [3] make questions about virus transmission mechanisms of plant viruses more approachable. In this chapter, we will briefly introduce the direct current (DC) electrical penetration graph (EPG) method. This monitoring system provides detailed analyses of stylet penetration by aphids and the vector-plant interactions involved in the transmission of plant viruses [4-6]. As will become clear from this chapter and the one that follows, most of the research in this area has been carried out by the research groups of Fereres (Madrid, Spain), Powell (Silwood Park, U.K.) and Pirone (Lexington, KY), more or less in cooperation with our group in Wageningen, the Netherlands. In this chapter, we focus on viruses transmitted by aphids in a circulative way and report mainly on EPGs of aphid transmission of barley yellow dwarf virus (BYDV). For an overview of persistently transmitted circulative plant viruses, we refer the reader to recent reviews such as that by Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
59
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W. FRED. TJALLINGII AND ERNESTO PRADO
(voltage source)
E Ri V
electrode potentials input resistor system voltage (V = Vs+E+E)
Vi Vs
input voltage (EPG signal) voltage supply
Fig. 1 The electrical penetration graph (EPG) recording circuit. The output (amp) is connected to a recording device, consisting mainly of a computer, with screen display and hard disk data acquisition.
Nault [7]. The next chapter will report on viruses transmitted in a noncirculative manner by aphids.
//.
The Electrical Penetration Graph Technique
In the electrical penetration graph (EPG) setup, plant and aphid are incorporated in an electrical circuit (Fig. 1) by inserting one electrode into the soil and attaching another with conductive silver glue to the aphid's dorsum via a thin (20|im), flexible gold wire. In principle, the electrical resistance or conductivity changes due to aphid-plant interactions cause the voltage at the measuring point to vary. Such resistance changes are caused, for example, by the opening and closing of valves of the food and saliva pumps. This is the R component of the EPG, which is measured by the original "feeding monitor" introduced by McLean and Kinsey [1] and its successors, the so-called alternating current (AC) systems. In contrast, the more recent DC system records more than just conductivity fluctuations. It also records some voltages (electromotive forces) at the measuring point that are generated by the insect-plant combination, that is the emf component of the EPG. Such voltages are caused, for example, by the membrane potentials of punctured plant cells along the stylet track and in the phloem. We will confine considera-
4.
71
CiRcuLATivE T R A N S M I S S I O N ANALYSIS
OVERVIEW Ih
path
path
phloem
path
phloem
DETAILS phloem (sieve element)
Fig. 2 Electrical penetration graph (EPG) of aphid feeding behavior. Top trace: One hour of probing and nonprobing alternation. A probe is a stylet penetration period; np is nonprobing. First two probes contain pathway-phase (path) only. The third probe also includes a xylem phase and two phloem phases. Many potential drops occur in the path periods. The first of the two phloem phases is short with El only; in the second. El is followed by E2. The rectangular part includes the E1-E2 transient, depicted in detail in Fig. 4. Bottom traces: Details of each phase with waveform indications; pd, potential drop related to a brief intracellular puncture.
tions here to the DC system. A more complete comparison of the two systems is given elsewhere [8]. The EPG waveforms, a 1-hour example of which is shown in Figure 2, arise as soon as the stylets are inserted into the plant. Correlation studies revealed the main relationship of waveforms to aphid activities and the location of the stylet tips in the plant tissue [3,5,6, 9-11]. The EPG studies on virus transmission reported in this book contributed substantially to these correlations. Periods of probing (stylet penetration) alternate with periods of nonprobing (np), which normally represent walking and selection of a new probing site in free, nonwired aphids. During probing, three behavioral phases can be distinguished, each containing one or more waveforms: (1) the pathway phase, including waveforms A, B, C, potential drop (pd), and F (not shown in Fig. 2); (2) the xylem phase, waveform G; and (3) the phloem phase, waveforms El and E2. Waveforms are distinguished on the basis of amplitude, frequency, voltage level, and electrical origin (R or electromotive force [emf]). They were neutrally labeled, using alphabetic characters sequentially assigned in order of their occurrence or description. The main correlations of the major waveforms are listed in Table I. With respect to virus transmission, ingestion is important for the acquisition of virus by its vector, whereas salivation (or other
Table Z Main Features and Correlations of EPG Waveforms Characteristics Wave-form (main part)“
Correlations
Relativeb amplitude
Rep. rate
Volt. level
Electr. origin
A
100
5-10
e
r
epidermis contacts
B
75
0.2-0.3
e
R
epidermis and mesophyll
C
30
mixed
e
R
all tissues
Pd
na
i
emf
all living cells, I and I11 extracellular, I1 intracellular
i i i e i
emf emf emf emf emf emf R Wemf emf R
4
I4
11-1
11-2 11-3 Ele El E2 w P F Gw P
na na na na na na
5 5 na 0-60
11-20 3-9 8-25 2-1 2-1 6 9 0.54 11-19 4-9 4-9
1
i e e e
Plant tissue
unknown sieve elements sieve elements sieve elements all tissues xylem xylem
Aphid activity electrical o d o f fwith salivation, possibly sheath salivation many activities during stylet pathway stylet puncture
salivation unknown ingestion unknown salivation (watery?) salivation (passive) ingestion erratic mechanical stylet work active ingestion unknown
Remarks first wave-form, electrical stylet contacts waveforms overlap therefore, often lumped in EPG analyses as “stylet pathway” (C) 3 phases, ph. I1 with 3 parts
non-pers. virus inoculation non-pers. virus acquisition same activity as E l ? persistent virus inoculation persistent virus acquisition penetration “difficulties” “drinking”, not in all probes
For each pattern the characteristics are quantified in terms of amplitude (min. to m a . ) relative to waveform A (=loo%), repetition rate of peaks or waves (Hz), extracellular (e) or intracellular (i) voltage level, and main electrical origin; i.e., fluctuation of resistance (R) or electromotive force (emf). Waveform parts: p, peaks; w, waves. Relative amplitudes (A=100%) are given as not applicable (na) for emf-origin waveforms [8].
4.
73
CiRCULATivE TRANSMISSION ANALYSIS
2 (of 3) types of ingestion
4
4
1 : cell punctures in path ^ ^ (phloem feeding) ingestion.
4
4
phloem
probing
4 types of
salivation
pathway Phloem salivation
2 ^
cell punctures in path (phloem feeding) salivation.
Fig. 3 An electrical penetration graph (EPG) with its consecutive main waveforms related to ingestion and salivation. Two types of ingestion (areas included in upward arrows) and four types of saliva secretion (areas included in downward arrows) are shown. One ingestion-period waveform is not shown, namely, waveform G related to xylem sap ingestion.
excretion) provides the vehicle for virus inoculation to healthy plants. For the waveform correlations found so far, the relevant activities are periods of salivation and ingestion, as summarized in Figure 3. Since viruses mainly operate inside living cells, the intracellular ingestion and excretion periods are of special importance. For acquisition, the presence of transmissible virions is important, whereas for inoculation, it is crucial that virus deposition occurs in cells that live long enough for virus to be replicated and transferred to neighboring cells. Since aphids' stylets normally do not destroy cells during plant penetration, no virions leak out and the ingestion of intercellular fluids seems unrelated to virus acquisition. Similarly, it seems unlikely that virions can penetrate living plant cells without the help of a vector. With respect to vector behavior, only intracellular salivation during stylet punctures seems important for virus inoculation. Such punctures into epidermal and mesophyll cells as well as into phloem cells are reflected in EPG waveforms, which makes EPGs usefiil in studying the probing events responsible for virus transmission. Intracellular punctures do occur in nearly all cells along the stylet track. These typically 5 to 10-second punctures are reflected by pd waveforms [8] (see next chapter) in the EPG and occur during the pathway phase of probing (Fig. 2, pd in path). Furthermore, apart from the brief pd punctures, long intracellular punctures occur in phloem sieve elements, and these are characterized by waveforms El and E2. Prior to virus transmission experiments described here, the relationship of the E2 waveform to (passive) phloem sap
74
W. FRED. TJALLINGII AND ERNESTO PRADO
ingestion and concurrent saliva excretion was demonstrated electrophysiologically [2] by using radioisotopic markers [10], by transmission electron microscopy after styletectomy [3, 12-14], and by simultaneous EPG and honeydew clock recording [15, 16].
///. A.
Barley Yellow Dwarf Virus Transmissiory by Rhopalosiphon pad!
Phloem Relationships of Virus and Vector
Luteoviruses such as BYDV are restricted to the phloem, involving sieve elements, companion cells, and phloem parenchyma cells interconnected by plasmodesmata. Virus ingested with phloem sap passes to the digestive tract and crosses the hindgut epithelium to the hemocoel. From the hemocoel, it is carried by hemolymph to the accessory salivary glands and inoculated with saliva to a healthy plant [17,18]. Barley yellow dwarf virus can serve as a marker for studying aphid salivation into (inoculation) or sap ingestion from (acquisition) phloem cells. Earlier attempts to link probing events to BYDV transmission were made by Scheller and Shukle [ 19] using an AC system for behavioral monitoring. The AC system only distinguishes one phloem-related waveform. The two major phloem-related waveforms distinguished by the DC system are El and E2; both waveforms indicate that the stylet tip is in a sieve element. The El waveform is often, but not always, followed by E2 in the same sieve element; on the other hand, E2 is always preceded by an El of variable duration. In 1978, the El and E2 waveform [then E(pd)] were correlated with passive phloem sap ingestion and concurrent salivation [2]. A revision of waveforms labels and their relationships to probing behavior was proposed in 1990 [20]. Still later, the phloem location of El could be confirmed [3], but no behavioral correlation was made until this study on BYDV transmission. B.
Material and Methods Used 1.
APHIDS
The aphids came from a stock culture of nonviruliferous Rhopalosiphum padi (L.) reared on wheat in the greenhouse at 20 ± 2°C and an L16:D8 photoperiod. Apterous virginoparous adults were used 3 to 6 days after ecdysis. Newly molted adults were transferred in small, individual clip-on cages to B YDV-infected wheat for 48 to 72 hours. Seventeen aphids were given a 1-week inoculation feeding period (IFP) on uninfected wheat to test their ability to transmit virus. 2.
PLANTS
Wheat (Triticum aestivum L.) c.v. Okapi served as virus source plants, test plants, and aphid-rearing plants. Test seedlings, 10 to 12 days old (two-leaf stage) were grown in a greenhouse at 23 ± 2°C.
4.
75
CiRcuLATivE TRANSMISSION ANALYSIS EPG
E1
E2
20 sec
E1, inoculation / salivation into sieve element
Wm cell wall
^ S sheath saliva
I stylet
E2, acquisition / Ingestion from sieve element
I watery saliva FTri plant sap
plasmalemma
Fig. 4 Diagram of electronic penetration graph (EPG) waveform El transient to E2 (top, c.f. framed part in Fig.l) and the events at the stylet tips (bottom). During El (bottom left), saliva is secreted into the sieve element. The fluid-filled food canal and the closed cibarial valve (not shov^n) do not allow saliva to be ingested. During E2 (bottom right), the sieve element sap is forced (under high pressure) into the food canal. The secreted saliva, therefore, will not reach the plant but is mixed with the sap. The fusion between the two canals near the tip is an essential anatomical feature.
3.
VIRUS
The B YDV source plants for test aphids were obtained after three successive R. padi transmissions of virus from original field-collected BYDV-infected wheat from Wageningen. A triple antibody sandwich, indirect enzyme-linked immunosorbent assay (TAS-ELISA) [21] of these original plants showed them to be positive for the (PAV; i.e. Ripadi associated) isolate of BYDV Polyclonal antibody to PAV (MAFF, Rothamsted, U.K.) and monoclonal antibody WAU-A7 [22] served to trap and detect BYDV, respectively. Each leaf sample was tested twice. 4.
EPG RECORDING
After wiring the aphid and inserting the plant electrode in the soil of a potted source or test plant, signals were recorded on a personal computer hard disk and analyzed by STYLET 2.0 software [23]. The setup (Fig. 1) was placed in a Faraday cage to eliminate noise. For analysis, the stylet pathway phase (Fig. 3), waveforms A, B and C (Table 1), and the two sieve element waveforms. El and E2 (Figs. 3 and 4), were distinguished. Waveforms F and G occurred occasionally but appeared irrelevant (GLIM analysis, see section in.D.2). About 1 hour was needed to collect aphids
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W. FRED. TJALLINGII AND ERNESTO PRADO
from culture plants or BYDV source plants, wire them, and give them access to plants. On the basis of waveforms, four categories of aphids were distinguished: those that produced (1) pathway-phase waveforms only; (2) pathway-phase and El waveforms; (3) pathway phase. El, and E2 (<10 min) waveforms; and (4) pathwayphase, El, and E2 (>10 min) waveforms. All aphids that produced E2 waveforms also produced El and pathway-phase waveforms. Durations of pathway-phase, El, and E2 waveforms were determined from the EPGs for each aphid. C.
Transmission Experiments 1.
ACQUISITION EXPERIMENT
Nonviruliferous R. padi were given access to BYDV source plants (2 months postinoculation) and their EPGs recorded. Plant access periods were standardized to 2 hours, but a number of aphids were monitored longer, up to 8 hours, to obtain enough replicates of each waveform category, for example, some aphids needed more time to begin producing E2 waveforms. On the other hand, to get aphids in the "pathway-phase + E l " waveform category, aphids had to be lifted from their plants before E2 production while producing the El waveform. After recording EPGs, each individual test aphid was gently lifted off the virus source plant (its gold wire was severed near the body) and transferred to its own uninfected test seedling for 7-10 days in the greenhouse. (Individual plant cages were used to avoid contamination.) After this period, test plants were fumigated with dichorvos to kill the aphids, and the cages were removed. Fumigation was repeated every 10-12 days. ELISA tests of each test plant were conducted approximately 3 weeks later. In total, 117 aphids were recorded and subsequently tested for BYDV transmission. 2.
INOCULATION EXPERIMENT
In a preliminary experiment, more than 94% of aphids reared on BYDVinfected plants and transferred to healthy test plants for 1 week transmitted virus. Thus, presumably at least 94% of the viruliferous test aphids had acquired virus. Such viruliferous aphids (proven transmitters) were then given access to a healthy seedling each and their EPGs were recorded. After recording, the aphids were removed and the seedlings placed in the greenhouse, treated with dichorvos and, 3 weeks later, tested by ELISA for BYDV infection. D.
Results and Discussion 1.
DIRECT
DATA
a. Acquisition. Positive ELISA results in test plants showed that BYDV was acquired, with certainty, by only 1 out of 42 test aphids (Table II) when only
4.
77
CiRCULATivE TRANSMISSION ANALYSIS
Table II Numbers of Barley Yellow Dwarf Virus (B YDV) Infected Test Plants in Relation to Electrical Penetration Graph (EPG) Waveforms Recorded During Acquisition and Inoculation Probes by R. padi. Virus Acquisition Waveforms shown Pathway Pathways-El Pathway + E l + E 2 Pathway + El + E2>10min Total
Test plants
Infected plants (%)
42 22 29 24 117
1 (2.4) a« 1 (4.5) a b 7 (24.1) b 14 (58.3) c 23 (20)
Test plants
Infected plants (%)
Virus Inoculation Waveforms shown Pathway Pathway + El Pathway + E l + E 2 Pathway + El + E2>10 min Total
31 21 44 44 140
4 6 24 23 57
(12.9) (28.6) (54.5) (52.3) (41)
a« b c bc
^ Treatments followed by the same letter are not significantly different according to the chi-square test {P < 0.05).
pathway-phase waveform C was produced during source plant penetration (category 1). Additional El activity (category 2) did not increase virus acquisition; however, when E2 activity also occurred, acquisition increased to 24% for category 3 (E2<10 min) and 58% for category 4 (E2>10 min). As with E2, the total duration and the number of periods of waveform El varied considerably for aphids in categories 1,2, and 3. Total durations of El waveforms ranged from 19 seconds to 14.4 minutes and occurred in one to six separate periods of El shown per aphid. No separate category was made for long-duration El activity. b. Inoculation, In the inoculation experiment, 4 of 31 viruliferous aphids transmitted the virus after pathway-phase only activities (category 1) (Table II, lower part). A higher proportion, 28%) inoculated plants after producing pathwayphase and El waveforms (category 2). A further increase in inoculation to 54% occurred when E2 waveforms were also produced (categories 3 and 4). However, total E2 durations of 10 minutes or more (category 3) did not result in higher transmission success than did shorter ones (category 4). As in the acquisition experiments, the total duration of the El waveforms varied considerably in categories 2, 3, and 4.
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W. FRED. TJALLINGII AND ERNESTO PRADO
2.
STATISTICAL PROCESSING OF TRANSMISSION DATA
Although the direct data (Table 2) already show a relationship between E2 activity and BYDV acquisition, the inoculation data are somewhat confusing. Actually, both activities seem to be involved. Virus acquisition and inoculation do not depend solely on stylet-tip positioning in the plant or on the occurrence of certain aphid activities; they also depend on the concentration of virus in ingested sap [24] or in the excreted saliva, respectively. Therefore, transmission remains more a matter of probability than particular conditions only. For this reason, aphids that match the conditions and produce appropriate EPGs may not successfully acquire or inoculate BYDV Apart from the occurrences of certain conditions, that is, waveforms or activities (Table 2), our data also include the precise number of times each type of waveform occurred and its total duration. This made the data suitable for frirther statistical analysis by GLIM (Generalized Linear Interactive Modeling) analysis [25]. As a dependent variate, the negative or positive reading in the ELIS A test was used with a binomial error distribution. The link function, linking the scale of measurement (0-1) to the linear scale on which the covariates were supposed to act, was the logit. In this logistic regression, we mcluded the following covariates per aphid: probing time, pathway time, numbers and durations of El and E2 waveforms, number of recognized phloem sieve-element punctures (i.e., periods with El or E1+E2), and durations of waveforms F and G. Testing for significant contribution of covariates in GLIM models is accomplished through comparisons of nested models, resulting in changes of deviance, which have a chi square distribution (under HQ). It appears that only the transformed values of El and E2 duration contributed significantly to any combination or sequence in the model. Therefore, the resulting final model was: logit {p) = log {pl\-p) = |30+pl*log(ElJ+l) + p2*log(E2^+l), in which/? is the probability of virus acquisition or inoculation, log(El
4.
CiRCULATivE TRANSMISSION ANALYSIS
79
Table III Deviance in the Logistic Regression Model of Acquisition and Inoculation Data. Acquisition Variables Empty model Model + logEld + logE2d Model + logE2d + logEld
deviance Deviance^ />-value
Inoculation
Deviance
Deviance" /?-Value
191
116 -19.0* (<0.001) -0.1 n.s. (0.75)
-2.8 n.s. -6.6*
(0.09) (0.01)
The change in deviance shows the effect of adding E2 (log duration) to the model after El (top) versus adding El after E2 (bottom), respectively. ^ Significant (*) and not significant (n.s.) as determined by Chi-square analysis.
At least a number of the brief pd punctures have been demonstrated to ocour in phloem parenchyma and companion cells as well as sieve elements [3]. Moreover, these pd punctures form the main probing events that are responsible for the acquisition and inoculation of nonpersistently transmitted viruses in epidermal and mesophyll cells [6] (see chapter 5). Presumably, only exceptionally high concentrations of virus particles in punctured phloem parenchyma or companion cells would lead to effective BYDV acquisition, since the ingested sap samples would be minute. In general, the aphids without E2 waveforms failed to acquire BYDV in spite of the many phloem cell pd punctures that they must have made. The positive correlation of the duration of the acquisition access period (AAP) and acquisition of persistent viruses found by others [24] is in agreement with our results, although that correlation is less direct and accurate than our correlation of acquisition with E2 duration. Sieve element acceptance, that is, sustained phloem sap ingestion (E2>10 min), usually does not occur earlier than 3 to 4 hours after the start of plant access by R. padi on wheat. In some aphid-plant combinations, a period of 5 to 7 hours from plant access to sieve element acceptance is normal [23]. This is about the mean period to achieve BYDV acquisition, which is considerable longer than the minimum acquisition time required by certain individual aphids. b. Inoculation, Logistic regression shows that adding the duration of El to the model after E2 results in a significant increase in deviance, whereas adding E2 after El does not (Table 3). An increase in duration of El caused increasing inoculation success by aphids that produced any El (Table 2), whereas an increase of E2 did not. Thus, El is the waveform related to virus inoculation and apparently reflects saliva excretion from the accessory glands into a sieve element. Long periods (>10 min) of subsequent E2 did not increase inoculation, in spite of the fact that during E2 there is supposedly continuous salivation [2, 10]. Evidence that E2 duration cannot be related to BYDV inoculation may either mean that this saliva does not come from the accessory glands or, more likely, that saliva secreted during E2 waveforms does not reach the plant. It has been sug-
80
W. FRED. TJALLINGII AND ERNESTO PRADO
gested earlier [3] that the E2 saHva is immediately ingested with the phloem sap (Fig. 4, bottom right). At the tips of the maxillary stylets, the two separate stylet canals, the food canal and the saliva canal, form a single lumen [30]. When the homopteran cibarial valve [31-33] is closed, phloem sap cannot enter the food canal, and therefore the excreted saliva that contains the virus will be injected into the sieve element (Fig. 4, left side). When the cibarial valve opens, however, the phloem sap is forced into the food canal by the high pressure in the sieve elements, and the saliva excreted into the common lumen of the stylet canals is carried up the food canal by the sap. In the latter scenario, saliva is unable to reach the plant (Fig. 4, right side). After a 1-week inoculation feeding period (IFF), the transmission eff^iciency increased to 94.1%. Following short IFPs in the experiments, the maximum efficiency was 54%. Power et al [34] obtained 47% and 87% inoculation efficiency, using a 2- and a 6-hour IFF, respectively. Sylvester [17, 35] suggested inoculation efficiency could be improved by allowing viruliferous aphids to make several probes (discontinuous feeding). It can be now be concluded that it is not the IFF duration or the number of probes but, more pointedly, the duration of sieve-element salivation periods (El waveforms) during a probe(s) that is positively correlated to inoculation efficiency. Longer plant access implies longer and more El waveforms, at least when the aphid is not continually ingesting phloem sap (E2). Most aphids do not start phloem sap ingestion at the first occasion but rather show several phloem phases, each starting with El before continuous feeding (E2) [23]. The fact that a few aphids inoculated test plants after having shown only pathway-phase activities of probing (waveforms A, B, and C) suggests that in some cases saliva can be injected into sieve elements or other infectible cells prior to phloem-phase activities (E waveforms). As for the non-phloem-phase acquisition, the pd punctures in any infectible cells can be responsible. Furthermore, non-phloem-phase inoculation apparently has a somewhat higher probability than similar acquisition. The amounts of fluids in both cases must be minute (see chapter 5), but the inoculations suggest that in a few aphids the virus concentration in saliva was high enough for transmission. If true, this may indicate that the saliva excreted during pd punctures also contains accessory gland material.
IV, studies of Other Circulative Viruses In earlier AC system studies, Leonard and Holbrook [36] and Scheller and Schukle [19] found no higher efficiency of virus transmission with longer periods of their "phloem ingestion." In the AC systems they used [1, 37], the "PI" (phloem ingestion) waveforms are equivalent to our DC system El and E2 waveforms. The AC system allows no distinction between El and E2 waveforms within the PI waveform, since they are not sensitive enough to the emf components in the electrical
4.
CiRCULATivE TRANSMISSION ANALYSIS
81
signals (see section II). The PI waveforms from the AC systems should be referred to as phloem position or phloem puncture, not as phloem ingestion, which cannot accurately be derived from AC signals (Table 1) [38]). Realizing that E2 periods longer than 10 minutes or even 1 hour are not preceded by proportionally longer El periods, these earlier observations seem to be in agreement with our observations. The invisible (to the AC system) part of real phloem ingestion (E2) during PI waveforms does not contribute to the inoculation. In a recent EPG (DC) study by Jiang et al [39], it was demonstrated that inoculation of the circulative geminivirus tomato yellow leaf curl virus (TYLCV) by the whitefly Bemisia tabaci (Gennadius) shows many parallels with BYDV transmission. Also, TYLCV inoculation is positively correlated with the duration of the El equivalent in Bemisia EPGs, i.e., the E(pd)l waveforms [40]. This similarity in a completely different virus group supports the idea that salivation into the living sieve elements is the main way of inoculation for phloem-restricted plant viruses that are persistently transmitted by phloem-feeding homopterans in a circulative way. In contrast to aphids, whiteflies do not puncture companion and parenchyma cells (pd waveforms) extensively before inserting their stylets into sieve elements. Nevertheless, some inoculated test plants in their study occurred without phloemphase activities (E waveforms) (no salivation from pd punctures of infectible phloem cells occurred), as we suggest for aphids (see section III.D.2.b). Kimmins and Bosque-Perez [41] could correlate an L2 waveform to the inoculation of the circulative maize streak virus (MSV), a geminivirus, using EPGrecorded probing by an African cicadellid leafhopper. Similarly to our El waveforms preceding phloem ingestion waveforms (E2), their L2 waveforms preceded a honeydew-related, intracellular L3 waveform. Very likely this L3 is related to phloem ingestion, although they did not perform any confirmatory acquisition experiments. Whether all observed L2 waveforms are related to inoculation is not clear. If so, leafhopper phloem salivation lasts extremely long (30 min or more) in comparison to salivation by aphids (generally a few minutes). Why would leafhoppers need to inject saliva into a sieve element much longer than do aphids to suppress the plant's wound-healing reactions (a suggested function of this salivation)?
V, Vector Resistance in Plants Can plant resistance to aphids reduce virus spread? Resistance (antixenosis) may deter aphids from inserting their stylets beyond tissues containing the resistance factor, a chemical or physical barrier. Some EPG studies have demonstrated that these factors can be located in different tissues [42,43]. It seems, however, that the impact on virus spread can be negative or positive, depending on the tissue location and on the strength or effectiveness of the deterring resistance. Ideally, vector resist-
82
W. FRED. TJALLINGII AND ERNESTO PRADO
ance against a phloem-restricted virus should prevent both phloem activities, El and E2, that occur after the phloem is reached. Hence, a resistance factor should be present in prephloem tissues such as the epidermis, mesophyll, or peripheral vascular tissue, and it should be strong enough to cause rejection of the plant or at least stylet withdrawal. If the resistance factor is located in the phloem itself, it may prevent long ingestion periods (E2) and thus suppress virus acquisition. However, aphids on resistant plants may salivate in the phloem (El) for the same time as [42] or even longer than aphids on nonresistant plants. The latter would presumably enhance inoculation by viruliferous aphids and hence primary virus spread. Shukle et al [44] demonstrated that phloem-located resistance to aphids seemed to be correlated with poor inoculation in some Agropiron species and cultivars, but others showing equal or greater apparent phloem resistance were very efficiently inoculated. Their study, however, could not discriminate between the effects of vector resistance and possible incompatibility of plant host and virus. Also, their AC system could not distinguish between El and E2 waveforms. Whether phloem factors will be able to prevent or reduce phloem ingestion periods (E2) enough to prevent virus acquisition is questionable. If reduced, the E2 periods may be short but repeated more often on resistant plants. Reduced acquisition would favorably affect the secondary spread of the virus. Resistant wheat plants with high levels of DIMBOA, purported to occur in the peripheral cells of the vascular bundle [45, 46], seem good candidates for protection against BYDV spread. Givovich and Niemeyer [47], using an EPG-monitored, 6-hour IFP, showed reduced BYDV inoculation by R. padi of wheat with high DIMBOA levels due to delayed phloem-phase activities (E waveforms). Later field tests of the same wheat seemed to confirm the reduced transmission. It remains unclear, however, whether the reduced field transmission was due to reduced primary infection, secondary spread, or both or to reduced aphid fecundity on these high-DIMBOA cultivars.
W. Concluding Remarks In general, EPGs are very suitable for analyzing probing behavior, identifying behavioral events responsible for acquisition and inoculation, and localizing plant tissues or cells that are involved in transmission. Conversely, viruses have turned out to be excellent markers for use with the EPG analytical system. Still other studies [42, 43, 48, 49] have shown the efficacy of EPG studies in localizing host plant resistance factors. However, one needs other methods to predict the value of host plant resistance to virus vectors in the epidemiological sense. One problem of electrically recorded probing on resistant plants is that wired aphids might be less affected by the plant's resistance, owing to locomotive restrictions, than are free, nonwired aphids [50, 51]. A wired aphid cannot leave the plant after "rejection" and will start to probe again; as a result.
4.
CiRCULATivE TRANSMISSION ANALYSIS
83
host plant resistance is often underestimated in long-duration EPG experiments. Thus, the impact of resistance should always be tested in additional experiments with free aphids.
References 1. McLean, D.L., and Kinsey, M.G. (1965). Identification of electrically recorded curve patterns associated with aphid salivation and ingestion. Nature 205, 1130-1131. 2. Tjallingii, W.F. (1978). Electronic recording of penetrations behavior by aphids. Entomol. Exp. Appl. 24, 521-530. 3. Tjallingii, W.F., and Hogen Esch, Th. (1993). Fine structure of aphid stylet routes in plant tissue in correlation with EPG signals. Physiol. Entomol. 18, 317-328. 4. Powell, G. (1991). Cell membrane punctures during epidermal penetrations by aphids: Consequences for the transmission of two potyviruses. Ann. Appl. Biol. 119, 313-321. 5. Prado, E., and Tjallingii, W.F. (1994). Aphid activities during sieve element punctures. Entomol. Exp. Appl. 12, 151-165. 6. Martin, B., Collar, J.L., Tjallingii, W.F., and Fereres, A. (1997). Intracellular ingestion and salivation by aphids may cause acquisition and inoculation of non-persistently transmitted plant viruses. J. Gen. Virol. 78,2701-2705. 7. Nault, L.R. (1997). Arthropod transmission of plant viruses: A new synthesis. Ann. Entomol. Soc. ^m. 90,521-541. 8. Tjallingii, W.F. (2000). Comparison of ac- and dc-systems for electronic monitoring of stylet penetration activities by homopterans. In "Principles and Applications of Electronic Monitoring and Other Techniques in the Study of Homopteran Feeding Behavior" (G.P. Walker and E.A. Backus, eds.). Thomas Say Publishers, Entomol. Soc. Am. Lanham, MD., pp. 41-69. 9. Tjallingii, W.F. (1985). Membrane potentials as an indication for plant cell penetration by aphid stylets. Entomol. Exp. Appl. 38, 187-193. 10. Tjallingii, W.F., (1988). Electrical recording of stylet penetration activities. In "Aphids, Their Biology, Natural Enemies and Control" (A.K. Minks and P. Harrewijn, eds.), Vol.2A, pp.95-108. Elsevier, Amsterdam. 11. Spiller, N.J., Kimmins, KM., and Llewellyn, M. (1985). Fine structure of aphid stylet pathways and its use in host plant resistance studies. Entomol. Exp. Appl. 38,293-295. 12. Mentink, P.J.M., Kimmins, KM., Harrewijn, P., Dieleman, F.L., Tjallingii, WE, van Rheenen, B., and Eenink, A.H. (1984). Electrical penetration graphs combined with stylet cutting in a study of host plant resistance to aphids Entomol. Exp. Appl. 36,210-213. 13. Kimmins, KM., and Tjallingii, W.F. (1986). Ultrastructure of sieve element penetration by aphid stylets during electrical recording. Entomol. Exp. Appl. 39, 135-141. 14. Kimmins, KM. (1986). Ultrastructure of the stylet pathway of Brevicoryne brassicae in host plant tissue, Brassicae oleracea. Entomol. Exp. Appl. 41, 283-290. 15. Tjallingii, W.F. (1995). Regulation of phloem sap feeding by aphids. In "Regulatory Mechanisms in Insect Feeding" (R.F. Chapman and G. de Boer, eds.), pp. 190-209. Chapman and Hall, New York. 16. Prado, E., and Tjallingii, W.F. (1997). Effects of previous plant infestation on sieve element acceptance by two aphids. Entomol. Exp. Appl. 82,189-200. 17. Sylvester, E.S. (1980). Circulative and propagative virus transmission by aphids. Annu. Rev. Entomol. 25, 257-286.
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18. Gildow, F.E. (1991). Barley yellow dwarf vims transport through aphids. In "Aphid-Plant Interactions, Populations to Molecules" (D.C. Peters, L.A. Webster, and C.S. Chlouber, eds.), pp. 165-177. Oklahoma State University, Stillwater, OK. 19. Scheller, H.V, and Shukle, R.H. (1986). Feeding behavior and transmission of barley yellow dwarf virus by Sitobion avenae on oats. Entomol. Exp. Appl. 40, 189-195. 20. Tjallingii, W.F. (1990). Continuous recording of stylet penetration activities by aphids. In "Aphids-Plant Genotype Interactions" (R.K. Campbell and R.D. Eikenbary, eds.), pp. 89-99. Elsevier, Amsterdam, the Netherlands. 21. Converse, R.H. and Martin, R.R. (1990). ELISA methods for plants viruses. In "Serological Methods for Detection and Identification of Viral and Bacterial Plant Pathogens" (R. Hampton, E. Ball, and S. de Boer, eds.), pp. 179-196. American Phytopathological Press, St. Paul, MN. 22. Heuvel, J.RJ.M. van den, de Blank, CM., Goldbach, and Peters, D. (1990). A characterization of epitopes on potato leafroll virus coat protein. Arch. Virol. 115, 185-197. 23. Tjallingii, W.F., and Mayoral, A. (1992). Criteria for host acceptance by aphids. In "Proceedings of the 8th International Symposium on Insect-Plant Relationships" (S.B.J. Menken, J.H. Visser and P. Harrewijn, eds.), pp. 280-282. Kluwer, Dordrecht, the Netherlands. 24. Gray, S.M., Power, A.G., Smith, D.M., Seaman, A.J., and Altman, N.S. (1991). Aphid transmission of barley yellow dwarf virus: Acquisition access periods and virus concentration requirements. Phytopathology 81, 539-545. 25. Aitkin, M., Anderson, D , Francis B., and Hinde, J. (1990). Statistical modelling in GLIM. Oxford Statistical Sciences Series 4. Oxford Science Publications. Clarendon Press, Oxford, England. 26. Barker, H., and Harrison, B.D. (1986). Restricted distribution of potato leafroll virus antigen in resistant potato genotypes and its effect on transmission of the virus by aphids. Ann. Appl. Biol. 109, 595-604. 27. Derrick, P.M. and Barker, H. (1992). The restricted distribution of potato leafroll luteovirus antigen in potato plants with transgenic resistance resembles that in clones with one type of host gene-mediated resistance. Ann. Appl. Biol. 120,451^57. 28. Shepardson, S., Essau, K., and McCrum, R. (1980). Ultrastructure of potato leaf phloem infected with potato leafroll virus. Virology 105, 379-392. 29. Heuvel, J.RJ.M., van den, de Blank, CM., van Lent, J.W.M., and Peters, D (1989). In situ localization of potato leafroll virus in infected plants by inmunogold-silver staining. (Abst.) In "Electron Microscopy Applied in Plant Pathology." (Abstracts) Konstanz, Germany. 30. Forbes, A.R. (1969). The stylets of the green peach aphid, Myzus persicae (Homoptera: Aphididae). Can. Entomol. 101, 31-41. 31. Weber, H. (1928). Skelet, Musculatur, und Darm de schwarzen Blattlaus, Aphisfabae. Scop. Zool. (Stuttgart) 76, 1-120. 32. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Morphology of the sweet potato whitefly, Bemisia tabaci (Homoptera, Aleyrodidae) relative to virus transmission. Zoomorpholog7 116, 143-156. 33. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Anatomy of a virus vector. In "Bemisia: 1995 Taxonomy, Biology, Damage, Control and Management" (D. Gerling and R.T. Mayer, eds.), pp. 289-318. Intercept Limited, Andover, U.K. 34. Power, A.G., Seaman, A.J., and Gray, S.N. (1991). Aphid transmission of barley yellow dwarf virus: Inoculation access periods and epidemiological implications. Phytopathology 81, 545-548. 35. Sylvester, E.S. (1949). Transmission of sugar beet yellow net virus by the green peach aphid. Phytopathology'^9, 117-132. 36. Leonard, S.H., and Holbrook, F.R. (1978). Minimum acquisition and transmission times for potato leafroll virus by the green peach aphid. Ann. Entomol. Soc. Am. 71, 493^95.
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37. Brown, CM. and Holbrook, F.R. (1976). An improved electronic system for monitoring feeding of aphids. Am. Potato J. 53, A51-A62. 38. Reese, J.C., Tjallingii, W.F., van Helden, M., and Prado, E. (2000). Waveform comparisons among AC and DC systems for electronic monitoring of aphid feeding behavior. In "Principles and Applications of Electronic Monitoring and Other Techniques in the Study of Homopteran Feeding Behavior" (G.P. Walker and E.A. Backus, eds.), Thomas Say Publishers, Entomol. Soc. Am. Lanham, MD. (In press). 39. Jiang, Y.X., Bias C. de., Barrios, L. and Fereres, A. (2000). Correlation between whitefly feeding behavior and TYLCV transmission. Ann. Entomol. Soc. Am. 93, 573-579. 40. Jiang, Y.X., Lei, H., Collar, J.L., Martin, B, Muniz, M., and Fereres, A. (1999). Probing and feeding behavior of two distinct biotypes of Bemisia tabaci (Homoptera: Aleyrodidae) on tomato plants. J. Econ. Entomol. 92, 357-366. 41. Kimmins, F.M., and Bosque-Perez, N.A. (1996). Electrical penetration graphs from Cicadulina spp. and the inoculation of a persistent virus into maize. Entomol Exp. Appl 80, A6-A9. 42. Helden, M. van, and Tjallingii, WF. (1993). Tissue localisation of lettuce resistance to the aphid Nasonovia ribisnigri using electrical penetration graphs. Entomol Exp. Appl 68,269-278. 43. Gabrys, B., Tjallingii, W.F., and Beek, T.A. van (1997). Analysis of EPG recorded probing by cabbage aphid on host plant parts with different glucosinolate contents. J. Chem. Ecol 23, 1661-1673. 44. Shukle, R.H., Lampe, D.J., Lister, R.M., and Foster, J.E. (1987). Aphid feeding behavior: Relationship to barley yellow dwarf virus resistance in Agropyron species. Phytopathology 11, 725-729. 45. Argandona, VH., Zufiiga, G.E., and Corcuera, L.J. (1987). Distribution of gramine and hydroxamic acids in barley and wheat leaves. Phytochemistry 26,1917-1918. 46. Massardo, F, Zufiiga, G.E., Perez, L.M., and Corcuera, L.J. (1994). Effects of hydroxamic acids on electron transport and their cellular location in com. Phytochemistry 35, 873-876. 47. Givovich, A., and, Niemeyer, N.M. (1991). Hydroxamic acids affecting barley yellow dwarf virus transmission by the aphid Rhopalosiphum padL Entomol Exp. Appl 59, 79-85. 48. Caillaud, CM., Pierre, J.S., Chaubet, B., and Pietro, J.P di (1995). Analysis of wheat resistance to the cereal aphid Sitobion avenae using electrical penetration graphs and flow charts combined with correspondence analysis. Entomol. Exp. Appl 75, 9-18. 49. Caillaud, CM., Pietro, J.P. di, Chaubet, B., and Pierre, J.S. (1995). Application of discriminant analysis to electrical penetration graphs of the aphid Sitobion avenae feeding on resistant and susceptible wheat. J. Appl Entomol 119,2, 103-106. 50. Tjallingii, WF (1986). Wire effects on aphids during electrical recording of stylet penetration. Entomol Exp. Appl 40, 89-98. 51. Lei, H., Tjallingii, WE, and van Lenteren J.C (1997). Effect of tethering during EPG recorded probing by aduhs of the greenhouse whitefly. J. Appl Entomol 111, 211-217.
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CHAPTER 5
Analysis of Noncirculative Transmission by Electrical Penetration Graphs ALBERTO FERERES JOSE LUIS COLLAR
/. Introduction In recent years there has been a considerable effort to elucidate and understand the transmission mechanisms and relationships between plant viruses and their insect vectors. Since the first efforts of Watson and Roberts [1] in 1939 to provide a system to classify insect-transmitted plant viruses, several authors have provided new classifications based on new findings and relationships discovered in the past 40 years [2-6]. Several techniques have provided new insights in the mechanisms of transmission and the interactions between viruses, vectors, and their host plants. These techniques include the use of artificial membranes (Parafilm) for the study of helper factors involved in the transmission process [7, 8] and visual observations of stylet penetrations [9, 10]. Electron microscopy techniques have been used extensively to localize virus particles in their insect vectors [11-14]. Transmission electron microscopy combined with immunogold labeling has provided information on the specific sites where helper components [15]) or virions [16] are attached and retained inside their insect vectors. The use of cDNA technology has also helped elucidate the mechanisms of plant virus transmission by their insect vectors [17-19]. One technique that has become indispensible in transmission mechanism studies is electronic monitoring of insect probing and feeding behavior, also called the electrical penetration graph (EPG) technique (also see chapter 4). The EPG technique, developed in the early 1960s by McLean and Kinsey [20], was first applied to aphids and to insect-plant interactions. During the last 10 Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
§7
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years, this technique has been appHed to behavioral studies of insect vectors during plant virus transmission. This in part has been possible because of the development of an improved electronic device based on direct current, the DC amplifier [21, 22]. Using the newer DC system, researchers can correlate recorded waveform patterns with specific stylet penetration and feeding activities of the insect. The DC system also distinguishes between intra- and extracellular stylet-tip positioning in plants, according to the potential level of the recorded signal [23]. This chapter deals with the analysis of electrically recorded probing and feeding activities of insect vectors associated with noncirculative transmission of plant viruses. Noncirculative transmission is a term that was first used by Harris [24] to describe aphid-transmitted viruses that are not able to circulate through the hemocoel and salivary system of their vector. This type of virus-vector interaction has only been described for insects in the order Homoptera.
//. Noncirculative Transmission: Properties and Vector Participants The main characteristics of noncirculative transmission of plant viruses could be surmnarized as follows: 1. There is no detectable latent period. Insects can inoculate virus to a healthy plant immediately after acquiring virus from an infected source plant. 2. Viruhferous insects are no longer inoculative following ecdysis. The virus is retained on the insect's cuticle and therefore lost during molting. 3. There is no evidence that noncirculative viruses are present in the hemocoel or the salivary system of their vectors. Also, the vector cannot transmit virus injected into the hemocoel. Noncirculative viruses have been subdivided into two main groups according to their persistence in vectors, nonpersistent [1] and semipersistent [25]. However, some authors prefer to refer to the nonpersistent viruses as stylet-borne {sensu Kennedy et al [2] carried at the tips of the stylets), and to semipersistent ones as foregut-bome {sensu Nault and Ammar [4], carried on the cuticular linings of the anterior alimentary canal). Others choose to refer to both nonpersistent and semipersistent noncirculative viruses as cuticula-bome {sensu Harris et al [5, 6], carried anterior to the foregut on the cuticular lining of the feeding-apparatus lumen). The differences between nonpersistent and semipersistent viruses as described by Harris [26] (summarized in Table I) are as follows: 1. Preacquisition starvation effect: Nonpersistent viruses are much more efficiently transmitted if vectors are subjected to a 1- to 2-hour starvation period prior to virus acquisition from an infected source plant. This effect is absent in the semipersistent type of transmission.
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ANALYSIS OF NONCIRCULATIVE TRANSMISSION BY ELECTRICAL PENETRATION GRAPHS
Table I
89
Discriminating Properties of Nonpersistent and Semipersistent Types of Transmission
Property Preacquisition starvation effect Acquisition and inoculation thresholds Optimal acquisition access period Retention of inoculativity by feeding aphids Acquisition and inoculation tissues
Nonpersistent Present Seconds Short (seconds) Minutes Epidermis
Semipersistent Absent Minutes Long (hours) Hours to days Phloem
Modified from Harris [26].
2. Acquisition and inoculation periods: The nonpersistent viruses are optimally transmitted when vectors make superficial brief (seconds) probes, whereas semipersistent viruses require longer probes lasting several minutes. The nonpersistent viruses are present in most plant tissues, including the epidermis, whereas semipersistent ones are usually phloem-restricted and therefore require longer acquisition and inoculation feeding periods. 3. Optimal acquisition of nonpersistent viruses occurs after very brief probes: Acquisition access periods lasting several minutes decrease the subsequent inoculation potential of nonpersistent viruses and increase the inoculation potential of semipersistent viruses. 4. Vector inoculativity and prolonged feeding: Vector retention of nonpersistent viruses is brief as compared with that of semipersistent viruses. When a vector acquires a nonpersistent virus, its ability to inoculate plants with virus decreases quickly (within minutes) with prolonged feeding. For semipersistent viruses, prolonged feeding does not negatively affect inoculation potential. Some viruses exhibit characteristics of both nonpersistent and semipersistent transmission. This unusual bimodal transmission pattern has been described for two viruses, cauliflower mosaic virus (CaMV) [27] and pea seed-borne mosaic virus (PSMV) [28]. These viruses may be effectively acquired during brief (seconds) or prolonged (several hours) probes. Depending on the vector species selected, virus can be acquired more efficiently after short or long acquisition access periods. For example, Myzus persicae acquires CaMV more efficiently during short (5-min) acquisition periods, whereas Brevicoryne brassicae acquires much more efficiently after long (8-h) access periods [27, 29]. Chalfant and Chapman [27] suggested the term bimodal because the first peak in their bimodal acquisition curve was the optimum acquisition access time for nonpersistently transmitted virus and the second peak was the optimum for semipersistently transmitted viruses. However, the term bimodal is somehow misleading, because the optimum acquisition peaks of CaMV may vary and exhibit a bi- or multiphasic pattern depending on the species of vector used for the transmission experi-
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ments [30]. There are probably behavioral differences in the way that specific vectors penetrate a plant's tissues that allow particular vectors to acquire virus after either short or long penetration periods. Alternatively, different virus attachment sites of different vector species could also explain why some aphid species acquire this type of virus best after short or long acquisition access times [31]. Of the more than 700 plant viruses known, more than half (58%) are transmitted by insects belonging to the order Homoptera [32]. Almost all of the known nonpersistent viruses (208 of 211) are transmitted by aphids (Homoptera: Aphididae), whereas semipersistent viruses are transmitted by aphids, whiteflies (Homoptera: Aleyrodidae), leaftioppers (Homoptera: Cicadellidae) or mealybugs (Homoptera: Pseudococcidae). Aphids are specially adapted to transmit plant viruses. From a total of 228 aphid species recorded as virus vectors, 200 belong to the subfamily Aphidinae [33]. The Aphidinae includes three genera of very efficient virus vectors (Aphis, Myzus, and Macrosiphum). Most of the Aphidinae are associated with herbaceous plant species, whereas the rest of the aphid subfamilies have tree hosts. The fact that viruses of herbaceous species have been much more intensively studied than those of trees could explain why the Aphidinae have been recorded as being the most numerous group of virus vectors [33]. Noncirculative viruses transmitted by aphids include members of the following taxonomic genera: Potyvirus, Carlavirus, Caulimovirus, Cucumovirus, Alfamovirus, Fabavirus, Closterovirus, and Sequivirus [34]). The first six groups contain members transmitted either nonpersistently or bimodally, whereas closteroviruses and sequiviruses are transmitted semipersistently. Although aphids are the largest group of vectors of noncirculative viruses, insects in other taxonomic groups are vectors as well. Some Closterovirus members are transmitted by mealybugs and whiteflies. Some carlaviruses as well as a few members of the Potyviridae are also transmitted by whiteflies (e.g., sweet potato mild mottle virus). Leafhoppers are vectors of the semipersistent maize chlorotic dwarf virus (MCDV, genus Waikivirus; family Sequiviridae). Some beetles (Coleoptera: Chrysomelidae) are vectors of semipersistently transmitted sobemoviruses (e.g., southern bean mosaic virus).
///.
Electronic Analysis of Nonpersistent Transmission
In this section, we will review the existing literature on electrically recorded aphid-probing activities and address crucial questions related to the process of nonpersistent virus transmission in an effort to better explain and understand the peculiar and unique properties of these viruses. These exclusive properties of nonpersistently transmitted viruses are difficult to understand, and a satisfactory explanation for all of them has not yet been formulated. However, electrical monitoring of aphid activities in plants during virus transmission has provided some possible explanations for the characteristics unique to this type of transmission.
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Table II Transmission Efficiency (%) of Cucumber Mosaic Virus (CMV) and Potato Virus Y (PVY) by A. gossypii and M. persicae, Respectively, during Acquisition and Inoculation Probes after Artificial Interruption at Different Phases of an Intracellular Puncture
Acquisition probe Interruption after subphase II-1 II-2 II-3
CMNIAphis gossypiilmQlon 6.1 % 9.4 % 35.4 %
Inoculation probe Interruption after subphase II-1 II-2 II-3
Virus/vector/plant pathosystem WYIMyzus persicae/pQpper 8.1% 18.7% 62.5 %
(4/65)« b (6/64) b (23/65) a
(3/37)^ b (3/16) b (35/56) a
Virus/vector/plant pathosystem CMWIAphis gossypiilmQXon 45.3 % 58.3 % 45.0 %
WYIMyzus persicae/pepper
(24/53) a (28/48) a (27/60) a
50.0 % 38.5 % 33.3 %
(6/12) a (5/13)a (9/27) a
" Numbers in parenthesis indicate the no. of infected plants/total plants tested. Values followed by the same letter within a table cell are not significantly different according to a chi-square test (contingency table). Modified from Martin et al. [39].
A.
Elucidating Transmission Mechanisms
Nonpersistent vims transmission can be divided into two well differentiated steps: (1) a brief probe on the superficial tissue of an infected plant (virus uptake or acquisition); and (2) a viruliferous aphid landing and probing on the superficial tissue of a healthy plant (virus release or inoculation). 1.
How Do APHIDS ACQUIRE NONPERSISENT VIRUSES?
Aphids can acquire virus particles during single intracellular punctures on infected plants while searching for a suitable host plant [35]. Aphids ingest from plants not only virus particles but in many cases also a virus-encoded "helper" protein, which becomes attached to the stylets [12]. It has been proposed that the helper protein present in several groups of nonpersistent viruses acts as a bridge between the virions and the retention sites of the stylets [15, 16, 36]. Intracellular punctures responsible for nonpersistent virus transmission can be monitored and visualized as potential drops using the EPG technique [21]. This technique distinguishes three distinct subphases: II-l, II-2, and II-3 [37, 38]. Virus acquisition or uptake occurs during subphase II-3, as demonstrated by using artificial diets [37] and plants [39] (Table II). Electrophysiological recordings of cibarial pump muscle activity of Acyrthosiphum pisum [21] show a repetition rate similar to the wave of the II-3 subphase of Myzus persicae (mean, 10 Hz;
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range, 7-12 Hz [38]) and Aphis gossypii (mean, 10 Hz; range, 8-1 1 Hz [40]). This presumably indicates that active ingestion during this phase is likely. Also, some pulses or archlets are associated with subphase 11-3 (1-2 Hz), but these do not correlate well with cibarial pump muscle activity as previously proposed by Powell et al. [37]. The significance of the archlet activity has yet to be determined. During subphase 11-3, aphids may ingest sap from the punctured cell together with the virus particles and any helper protein present. The virus particles are then retained in the distal third part of the maxillary stylets [16] for some time until the aphid probes another plant. Hodges and McLean [41] found a significant correlation between bean yellow mosaic virus (BYMV) virus acquisition and a characteristic waveform produced by A. pisum during superficial probing. The waveform was identified with an AC system (see chapter 4) as a plateau period. They suggested that this waveform occurred when the stylets of the aphid were extended beyond the tip of the salivary sheath during which time no saliva was being secreted indicating that the stylets were exposed to the infected tissue, thus allowing for virus acquisition. No correlation was found however, between inoculation of BYMV and the occurrence of a plateau period. One possible explanation for these results could be that the plateau waveform represents part of the potential drop (pd) waveform of EPG recordings. Unfortunately, the AC system shows no details in the plateau waveform, which makes comparison with pd subphases (DC system) impossible. 2. WHATMAKESAN APHIDINOCULATE A NONPERSISTENT VIRUS? A viruliferous aphid is capable of inoculating a healthy plant very soon after probing (in less than 10 s). We have recently found that virus release from the stylets occurs at the very beginning of an intracellular puncture, subphase 11-1 [39] (Table 11), which is usually a very short, high-frequency (12-18 Hz) subphase of about 1 second duration. The mechanism of virus release by the aphid is not yet fully understood. Two different hypotheses have been proposed to explain the process of nonpersistent virus inoculation by aphids. a An Egestion Mechanism. Harris [3] proposed that aphids could egest previously ingested sap through the alimentary canal and thus drive out previously acquired virus particles. He proposed that the cibarial pump can work in both directions, for either sap ingestion or egestion. Evidence for the egestion hypothesis was obtained from observations (with a light microscope) of aphids feeding on artificial diets [7]. Harris [3] found that aphid ingestion of ink particles through Parafilm membranes occurs before any egestion takes place: "During ingestion, ink particles flow from the surrounding medium toward the sheath tip. . . . The stylets usually extend almost to the opening in the stylet sheath tip during ingestion, but they are occasionally withdrawn some distance into the sheath or rarely projected several micrometers past the tip. Ingestion stops, as indicated
5.
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93
/'N/«--'«^
EPG
pd phase:
I
pd 11-1, inoculation / intracellular salivation
^m
cell wall ^ ^ stylet
I sheath saliva
III
pd 11-3, acquisition / intracellular ingestion
I watery saliva [•:-.-.| cytoplasma —— plasmalemma
• virion
Fig. 1 Top half, typical potential drop (pd) produced by M. persicae soon after the beginning of a probe and its intracellular subphases (II-1, II-2, and II-3) correlated (bottom half) with newly hypothesized mechanism of nonpersistent virus inoculation (bottom left) and acquisition (bottom right). "Cross-sections" through stylet bundle tip showing common duct (fused maxillary and food canals) during intracellular puncture of an epidermal or mesophyll plant cell. (From Martin et al [39]).
by a cessation in the flow of ink particles, and is usually followed by further ingestion or by regurgitation." Kloft [42] also indicated that aphids previously fed on tritium-labeled plants were able to transfer a large amount of tritium to leaf discs during probes lasting 1 to 3 minutes. However, the latter experiments could not differentiate whether salivation or egestion inoculated tritium to the leaf disks. b. A Salivation Mechanism. Abundant salivation during aphid probing (visualized as a B waveform with use of a DC device) continues until puncturing of an epidermal or mesophyll cell. At that point, Martin et al. [39] believe that watery saliva excretion flushes the virus particles into the cytoplasm (Fig. 1). The hypothesis is based on the fact that aphids inoculate nonpersistent viruses before any ingestion can take place. The sequence of events in EPGs makes salivation a better candidate for inoculation than egestion, because egestion seems to occur after ingestion during prolonged aphid feeding on artificial systems [9]. Moreover, it seems that aphids need to ingest first to determine host suitability. No other gustatory chemoreceptors are present beside the ones in the region of the cibarial valve [43]. Why should an aphid egest any sap material before sampling a new host? Salivation, as opposed to egestion, is a continuous event, which occurs
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from the very beginning of plant penetration by an aphid. Presumbably, virions can be released by saliva excretion because the food and salivary canals fuse about 2-8 |Lim from the tips [44]. Therefore, it would seem that attached virus particles can be released equally well by saliva excretion. B.
Virus-Vector Specificity
The largest group of plant viruses, the nonpersistent viruses, are transmitted almost exclusively by aphids. This suggests the existence of one or more important morphological, physiological, or behavioral traits that separate aphids from the rest of the insect vectors of plant viruses. 1.
WHY DO APHIDS ALONE TRANSMIT NONPERSISTENT VIRUSES?
Transmission of nonpersistent viruses is restricted, almost exclusively, to very brief (5- to 10-s) intracellular punctures of epidermis or mesophyll [45, 46]. Such probes can be recorded and visualized as pds by using a DC system. Intracellular punctures by aphids are very brief and frequent, occurring on average at 1-minute intervals [47, 48] and lasting about 5 to 7 seconds. For other homopterans, intracellular punctures occur much less frequently than in aphids, although they can last longer. In the cassava mealybug, Phenacoccus manihoti, the average duration of an intracellular puncture is 20 seconds [49]. For the sweet potato whitefly, Bemisia tabaci, intracellular punctures occur very rarely; their average duration in tomato plants is approximately 5 seconds [50]. The foregoing probably relates to the fact that aphids do not have, as do other homopterans, chemoreceptors on the tarsi or on the tip of the labium, but have only mechanoreceptors [24, 51, 52]. The major gustatory organ in aphids is located just before the cibarial pump [43] on the epicibarium of the antecibarium {sensu Harris et al [6]), and sap ingestion through the food canal is necessary to "taste" a plant. There is direct evidence that aphids can sample the cell contents during intracellular punctures from the very beginning of plant penetration to discriminate between host and nonhost plants and to find the favored site on their host [53, 54]. In this process, aphids can acquire nonpersistent viruses very efficiently [37, 39]. Another particular property that many aphid species display during the last part of a long potential drop (pd-L) is a characteristic and specific subpattern structure within subphase II-3 containing several pulses or archlets (three or more at about 2-Hz frequency). These pd-Ls occur most frequently during the first brief superficial probes and have been associated with nonpersistent virus transmission of CMV by ^. gossypii [55]. Such pd-Ls with their characteristic subphase II-3 have never been recorded in other groups of insect vectors such as whiteflies [50, 56, 57], leafhoppers [58], or mealybugs [49]. It is likely that ingestion from punctured epidermal cells occurs only for aphids. Other homopterans do not need to sample cell contents before reaching the phloem, since they carry chemoreceptors
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ANALYSIS OF NONCIRCULATIVE TRANSMISSION BY ELECTRICAL PENETRATION GRAPHS
95
at the stylet tips. Actually, in the case ofB. tabaci whiteflies, the first intracellular puncture occurs about 2 hours after obtaining access to a tomato plant [50]. In contrast, the first intracellular puncture by aphids usually occurs within the first minute of access to a plant (often within 15 sec). 3.
WHY ARE SOME APHIDS MORE EFFICIENT VECTORS THAN OTHERS?
For a specific virus-plant combination, there exist both efficient and inefficient vectors. Powell et al [59] found a positive correlation between the frequency of intracellular punctures (recorded as pds) and potato virus Y (PVY) transmission efficiency when comparing the probing behavior of Brachycaudus helychrisi and Drepanosiphum platanoidis on tobacco plants. Chen et al. [55] found that the mean duration of a pd-L was significantly longer for melon plants that became infected with CMV than for ones that did not. This same research showed that the longest intracellular punctures were very significantly associated with a high transmission efficiency of CMV by A. gossypii. Collar and Fereres [60] carried out an experiment using a PVY-infected pepper plant as a virus source in which they recorded the first pd within the first probe for several aphid species. They found that the most efficient vectors of PVY (M persicae and A. gossypii) were producing pd-Ls (with a long II-3 subphase) in most of the cases. On the other hand, poor vectors of the virus (Sitobion avenae and Rhopalosiphum padi) only rarely produced pd-Ls. This suggested that aphid species that colonize pepper plants show a typical early sap sampling behavior, directed at host discrimination, whereas noncolonizers (cereal aphids) have a tendency to walk along the leaf and when forced to probe (due to the wiring), usually make very short pd's with no associated ingestion activity and therefore with very low chances for virus acquisition. Moreover, many S. avenae and R. padi individuals were discarded because they did not probe or did not produce a pd within 5 minutes. This may indicate that many individuals from these two species did not need to sample plant sap (via epicibarial chemoreceptors) to exclude the pepper as a host. Instead, olfactory receptors on the antennae or mechanoreceptors on tarsi, antennae, or stylet tips could be enough to identify the plant as not suitable for settling and probing. Furthermore, in recent research conducted in our laboratory (Fereres, unpublished results) we found that Rhopalosihum maidis, a major vector of maize dwarf mosaic virus [61], almost always produces pd-Ls (with a long II-3 subphase) during the first probe on maize test plants. In contrast, less efficient vectors such as R. padi and S. avenae produce pd-Ls at a much lower frequency. This same research showed that some EPG variables are dependent on the host plant on which the aphid feeds, such as preprobing time and probing time to the first pd, whereas others depend on the aphid species (e.g., duration of subphase II3 and relative occurrence of pd-Ls). All the foregoing suggest that efficient aphid vectors of nonpersistent viruses tend to produce pd-Ls with several archlets during their first probes.
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It should be noted as well that in addition to behavioral factors, the potyvirus helper component plays an essential role in aphid transmission efficiency and vector specificity by interacting differentially to specific binding sites in the food canal of different aphid species [62]. The existence of anatomical and physiological differences between aphid species could regulate their interaction with potyvirus helper components and explain the qualitative and quantitative differences in their ability to retain and release specific nonpersistent viruses. C. Preacquisition Starvation Effect Since Watson [63] first described the preacquisition starvation effect, several authors have proposed various explanations for the phenomenon. Some proposed the existence of a virus-inactivating substance, perhaps carried by aphid saliva [1] or present in plant sap and remaining in aphid stylets [64], that would be absent in aphids subjected to fasting before acquisition. The existence of plant components that interfere with the retention of virions in the aphid stylets was supported by Wang and Pirone [65], who reported that aphids fed on artificial diets did not show the preacquisition effect as did the ones maintained on plants. Other authors [24, 35, 66, 67] have suggested that behavioral differences may be responsible for the observed phenomenon. Harris [3] indicated that feeding could discourage aphids from their typical sap sampling behavior, consisting in brief probes and directed toward early host plant selection. Nonstarved aphids would show instead a phloem search behavior, less likely to result in virus acquisition. Interestingly, Bradley [68] reported that those few nonstarved aphids that produced brief probes transmitted as efficiently as starved ones. Collar and Fereres [60] (see Table III) found that individuals of M. persicae subjected to a preacquisition starvation period of 1 hour (efficient PVY vectors) were very likely to produce very long II-3 subphases with several archlets during their first pd or intracellular puncture, as opposed to aphids that were not starved (inefficient PVY vectors). Interestingly, a similar decrease in the duration of subphase II-3 (and in the number of II-3 pulses) was also observed for M. persicae and A. gossypii when previously fasted aphids were allowed to produce consecutive pds within the same probe [40, 60]. A possible explanation for this particular behavior is that starved aphids could have a greater need for sampling the plant (because of dehydration during starving) and that this need decreases as the probe proceeds and successive intracellular punctures are produced. However, other nonbehavioral explanations to the preacquisition starvation effect should not be excluded (as proposed by Wang and Pirone [65]). The described starvation effect does not seem to work for every aphid species. It has been reported that there is at least one aphid species, the cabbage aphid, B. brassicae, which does not exhibit this type of behavior (Chalfant and Chapman [27]). Instead, B. brassicae tends to feed continuously for at least 5 minutes just after starving, whereas M persicae makes several short probes lasting about 30 seconds.
5.
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Table III Comparison of Behavioral Variables Obtained by Electronic Monitoring of Starved and Nonstarved M. persicae on a Pepper Plant during their First Probe and First Potential Drop (Indicating an Intracellular Puncture) Behavioral variables
Starved aphids
Preprobing time, s Probing time to 1 st pd, s Duration of pd, s Duration of phase II-1, s Duration of phase II-2, s Duration of phase II-3, s No. II-3 pulses No. records without II-3 pulses/total
45.96 ±5.94 10.46 ±0.84 6.22 ±0.19 1.62 ±0.04 0.97 ± 0.03 3.63 ±0.18 5.74 ± 0.39 6/54
Nonstarved aphids
P
42.42 ± 7.60 13.04 ±1.61 4.92 ± 0.24 1.46 ±0.05 0.94 ±0.04 2.52 ±0.21 3.41 ±0.44 22/56
.095« .246^ <.001^ .023^ .144« < .OOP < .OOP < .OOP
^ according to Mann-Whitney t/test (non-Gaussian variables) ^ according to Student's ^test (Gaussian variables). ^ according to chi square test (contingency table), pd = potential drop. From Collar and Fereres [60].
D.
Optimal Acquisition Time and Retention of Inoculability
Nonpersistent viruses are generally present throughout an infected plant because the infection becomes systemic a few days after inoculation. However, aphids only acquire nonpersistent viruses during short superficial probes. When an aphid makes consecutive pds within a probe, the II-3 phase duration and number of archlets also decreases dramatically, as has been reported for M persicae [60] and A. gossypii [40], two of the most efficient vectors of nonpersistent viruses. Therefore, if subphase II-3, which seems to be the crucial step for virus acquisition, is not long enough (minimal time or virus concentration or both is needed), the probability of virus acquisition is notably reduced. The first pds or intracellular punctures after the first plant penetration by an aphid's stylets are the ones that show a longer II-3 subphase and are also the ones related to virus acquisition [40]. Conversely, aphids that probe on a plant long enough do not usually produce pds with long II-3 subphases. This has been shown in 4-hour EPG recordings on melon for both A. gossypii and M. persicae [69]. More precisely, for A. gossypii, 20% of all pd-Ls recorded occurred as the first intracellular puncture, 22% occurred as the second puncture, and only 8% occurred after the second puncture. Similarly, for M persicae, 75% of all pd-Ls occurred as the first intracellular puncture and 25% occurred as the second or third. It is possible that once an aphid tastes a plant after a few long intracellular punctures, ftirther cell punctures are very short, sap sampling is interrupted, and consequently the probability for virus acquisition is dramatically reduced. Once the plant is recognized as a host, brief tasting stops and the aphid starts searching for a sieve element in the phloem to obtain a continuous food source. This is consistent with the fact that M persicae
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is a more efficient vector of CaMV during short feeds (<5 min) than during long ones (8 h). However, B. brassicae transmits better during long feeds [27, 29]. Interestingly, B. brassicae is not affected by the preacquisition starvation effect [27], which correlates with the ability of this species to maintain or increase its transmission potential after long feeds. It is possible that B. brassicae is able to produce intracellular punctures with a long II-3 subphase after several hours of penetration into the plant, but experimental evidence for this is lacking. We only know that B. brassicae tends to spend more time in nonprobing activity and makes more probes than M persicae before extending the probe duration when feeding on wild and cultivated Brassica species [70]. Instead, another explanation could be that B. brassicae is able to accumulate and retain CaMV particles in its stylets after feeds lasting as long as 8 hours, as suggested by Markham et ah [30]. These authors found out that CaMV behaves more like a semipersistent virus and suggested that the differences between this virus and turnip mosaic virus (a typical nonpersistent virus) are likely to be due to differences in the interaction between the virus and the vector (e.g., different binding sites). It has been shown many times that once an aphid acquires a nonpersistent virus, its ability to retain and subsequently transmit the virus into a healthy plant decreases very soon. Collar et al [38] studied PVY acquisition by M persicae during 5-minute recordings. They observed that the time elapsed from the last pd or intracellular puncture produced by an aphid and the end of the probe was crucial for subsequent PVY transmission. Interestingly, longer probing times from the last pd to the end of the probe were correlated with lower transmission efficiencies. This suggests that virus inoculability begins to decrease immediately after virus acquisition if probing continues. It has been shown that the retention of inoculability for nonpersistent viruses may be greatly enhanced up to more than 30 hours when viruliferous aphids are not allowed to probe on solid surfaces [71-73]. This means that the viability of the transmissible virus inside the aphid's stylet is much longer than the 5 minutes used for the experiments described by Collar et al [38]. Therefore, the observed loss of inoculability found in such a short time interval must be attributed to some kind of stylet activity produced by the aphid during the stylet pathway (C waveform using EPG terminology), and not to a passive process. One explanation to the observed phenomenon is that the virus particles may be swept away from the aphid's retention sites as a result of some salivation or egestion events taking place after virus acquisition. Active salivation or egestion could explain why the virus is not retained by the aphid's mouthparts for a long period of time after acquisition. E.
Effect of Insecticides and Mineral Oils on Nonpersistent Transmission
The specific and peculiar properties of nonpersistent transmission (specially the very short acquisition and inoculation thresholds required, together with the absence of a latent period) make the spread of nonpersistent viruses very difficult
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to prevent. In most field situations an efficient control of the vector population by chemical means is not enough to achieve a real inhibition of disease [74-76]. In other cases, it has been reported that insecticides could provide some control against secondary virus spread by colonizing apterae [77-79]. Primary virus spread by incoming alatae is much more difficult to avoid [80, 81]. An ideal insecticide for efficient control of nonpersistent virus diseases should be able to kill the aphid vectors before they are able to accomplish the transmission process, which in some cases may last only a few minutes. Some authors have reported that certain pyrethroids provide a quick knockdown [82, 83], which could be used for the control of nonpersistent viruses [84, 85]. Some studies have used electronic monitoring techniques to study the effect of several insecticides or antifeeding substances on aphid probing behavior [86-93], but very few have also considered their effect on nonpersistent virus transmission. Collar et al [94] used the EPG technique to study the behavioral response of M persicae to three insecticides (cypermethrin, pirimicarb, and imidacloprid) during PVY acquisition from pepper plants (10-min recordings). Only the pyrethroid cypermethrin was able to affect aphid probing behavior in that short period of time. Cypermethrin induced an abrupt interruption of the probing activity after a 2.5-minute exposure to the treated plant and incapacitated the aphid vector with respect to subsequent virus inoculation. Nevertheless, if shorter acquisition access times were considered, M persicae individuals were able to efficiently acquire and inoculate PVY before cypermethrin took effect. Bradley et al [95] first discovered that oils could inhibit nonpersistent virus transmission by aphids. Since then, many laboratory and field studies have shown that mineral oil application to plants, alone or in combinations with insecticides, successfiilly prevented or reduced nonpersistent virus spread [76, 96-103]. The mechanism by which oil inhibits virus transmission of aphids has not been completely elucidated. Several [104, 105] indicate that mineral oil can interfere with the retention of the virus particles in the aphid mouthparts. Wang and Pirone [105] speculated that the formation of an oil hydrophobic layer in the food canal might be responsible for this interference with the retention of virions. Few authors have used electronic monitoring techniques to determine if behavioral changes induced by oil application could be also involved in the prevention of virus transmission. Simons et al [106] reported that the preprobing time shown by aphids was enhanced on oil-sprayed plants. Powell [107] also reported a significant delay in the initiation of penetration but observed that the percentage of aphids that produced intracellular punctures (recorded as pd's) was not affected by oil application to PVY-infected tobacco plants. However, Powell did not distinguish between short and long pd's (with a long II-3 subphase and its associated ingestion activity). Collar and Fereres [108] studied the probing behavior of M persicae individuals (via an EPG monitor) during 5-minute access periods on a PVY-infected pepper plant (Table IV). Oil application (Vektaphid) to a pepper plant significantly
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Table IV
ALBERTO FERERES AND JOSE LUIS COLLAR
Effect of Vektaphid Oil on M. persicae Probing Behavior on a Pepper Plant dxiring 5 Minutes
Behavioral variables Preprobing time, s Time to 1st potential drop (pd), s Number of potential drops (pds) Average duration of pds, s
Before oil application
After oil application
P^
14.4 ±2.4 27.2 ±5.1 2.6 ± 0.2 5.9 ±0.2
46.9 ± 8.9 76.2 ±11.5 1.5 ±0.2 5.6 ±0.5
<.01 .14 <.01 .12
" P according to Mann-Whitney U test (non-Gaussian variables).
increased preprobing time, as previously reported. Also, aphids probing oiltreated plants produced fewer pds, although their average mean duration was not affected. Interestingly, phase II-3 of the first pd produced was slightly shortened after oil treatment (data not shown). In a separate experiment, aphid behavior was recorded during a single penetration on a PVY source plant (before and after oil application). The probe was interrupted after the first pd was observed, and aphids were immediately transferred to single healthy pepper seedlings for assessment of subsequent PVY transmission. The same behavioral changes were observed, with longer preprobing times and slightly shorter II-3 subphases (Table V), as well as differences in transmission efficiency (45% for controls versus 10% for oil-treated plants). However, no correlation between aphid behavior and PVY transmission was found when comparing EPG recordings from aphids that transmitted PVY with those from individuals that did not. Interestingly, the number of aphids that produced a pd-L, with more than three pulses observed in subphase II3, was not affected by oil application (>95% in both cases). These results indicate that although some behavioral changes may be induced by oils, other nonbehavioral factors must be mainly responsible for its inhibitory effect on transmission, as suggested by Wang and Pirone [105].
\V. Electronic Analysis of Semipersistent Transmission Semipersistent cuticula-bome viruses have been called foregut-bome [4]. However, recent morphological studies of the homopteran feeding apparatus indicate that the latter term is a misnomer. Virions positioned posterior to the opening in the pharynx of the foregut are nonegestible and therefore nontransmissible [5, 6], Semipersistent viruses are transmitted by aphids, whiteflies, leafhoppers, and mealybugs. These viruses do not need a latent period in the vector to be transmitted, cannot be recovered from vector hemolymph, and cannot be transmitted after injection into the vector hemocoel. Infectivity is retained only a few days and is lost after ecdysis.
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Table V Effect of Vektaphid Oil on M. persicae Probing Behavior on a PVY-Infected Pepper Plant until the First Intracellular Puncture (Recorded as a Potential Drop) was Produced Behavioral variables Preprobing time, s Duration of pd, s Duration of phase II-1, s Duration of phase II-2, s Duration of phase II-3, s No. II-3 pulses No. records with >3 pulses/total PVY incidence on test plants
Before oil appHcation
After oil application
P
36.25 ± 9.34 9.70 ± 0.45 1.89±0.11 1.08 ±0.06 6.72 ± 0.44 9.95 ± 0.68 20/20 45.0% (9/20)
67.87 ±13.59 7.70 ±0.33 1.83 ±0.08 0.97 ± 0.03 4.89 ±0.35 7.05 ± 0.56 19/20 10.0% (2/20)
<.01* <.01* .42^ .13^ <.01^ <.01« .3K <.0K
" P according to Mann-Whitney (7 test (non-Gaussian variables) ^ P according to Students /-test (Gaussian variables) ^ P according to chi-square test (contingency table). pd = Potential drop. Modified ft-om Collar and Fereres [108].
Transmission of semipersistent viruses such as maize chlorotic dwarf virus (MCDV) by Graminella nigrifrons leafhoppers (Homoptera: Cicadellidae) has been studied by researchers [14, 26, 109-111]. Harris et al have proposed that MCDV is inoculated to plants by egestion [14, 26]. Harris [26] further suggested that the electron-dense matrix material observed in MCDV inclusions in plants [112] and leafhoppers [14, 26] and between virions and leafhopper cuticula [14] could be helper protein(s) by which virions bind to sites on the lining of the vector's ante- and postcibarium {sensu Harris et al. [5, 6]). Ammar and Nault [109], on the other hand, propose that leafhoppers inoculate MCDV to plants by extravasation. As coined by McLean and Kinsey [113], the term extravasation is based on the supposition that homopterans such as aphids and leafhoppers are anatomically incapable of egestion. However, both these homopterans routinely utilize egestion while feeding [9, 10]. Furthermore, more recent and more detailed anatomical studies indicate that the morphology of the homopteran feeding apparatus is ideally suited for egestion as well as ingestion [5, 6]. Egestion also explains leafhopper inoculation of rice tungro and tungro-like viruses, and even the bacterium responsible for Pierce's disease [10]. It is very unlikely that MCDV would be inoculated to plants by salivation since most of the MCDV-like particles found attached to the feeding apparatus of G. nigrifrons are in the antecibarium, postcibarium, or cibarial pump {sensu Harris et al [5, 6]) and the pharynx, through which ejected saliva does not pass. Also, as noted earlier, virions in the pharynx are nonegestible and therefore nontransmissible [5, 6]. Particles of MCDV have been occasionally found in the maxillary food canal but never in the salivary canal of viruliferous leafhoppers [14, 109]. Even if the inoculation of MCDV could occur during salivation, the probability of transmission of this phloem-restricted virus would be very low because most of
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the salivation produced by G, nigrifrons has been correlated with nonphloem tissues [114, 115]. The leafhopper x-waveform (phloem contact) is thought to be responsible for MCDV inoculation [116]. The limited research on the mechanisms of semipersistent virus transmission indicate that they are transmitted in an ingestion-egestion manner [3, 10]. The ingestion-salivation mechanism is unlikely in the case of semipersistent viruses. This opens the possibility that some viruses may be transmitted in a typical nonpersistent or a semipersistent manner depending on the type of behavior of the vector species involved. Short superficial probes result in acquisition of nonpersistent (or bimodal) viruses, whereas longer probes associated with phloem ingestion result in acquisition of semipersistent (or bimodal) viruses. An alternative explanation is that semipersistent and nonpersistent viruses have different retention sites in their respective vectors. Virions involved in transmission in a nonpersistent manner are retained in the distal third of the food canal [12], whereas semipersistent viruses are likely to be retained and released from the antecibarium and postcibarium (sensu Harris et al [5, 6]) Xylella fastidiosa, SL bacterium transmitted in a semipersistent manner by leafhoppers (Cicadellinae) and spittlebugs (Cercopidae) is lost after molting of the vector. This bacterium is transmitted by xylem feeding insects with very low vector specificity. In this case, the mechanism of transmission is thought to be egestion [10] from the antecibarium and postcibarium or extravasation from the foregut caused by a malfunction of the precibarial valve [116]. Leafhopper food and salivary canals do not fuse at the end of the maxillary stylets as they do in aphids. Therefore, salivation does not seem to be the mechanism involved in inoculation of plant pathogens that are retained in the feeding apparatus of leafhoppers and other vectors of the Auchenoryncha group.
V
Concluding Remarks
Electrical monitoring of stylet penetration activities has significantly elucidated our understanding of virus transmission mechanisms and relationships between plant viruses and their insect vectors. Studies conducted in the past 10 years have clarified important aspects and properties of nonpersistent virus transmission and made possible a new hypothesis on the mechanism of transmission. Most of these studies have concentrated on the EPG analysis of intracellular punctures during stylet penetration by aphid vectors. These intracellular punctures, recorded as pds, can be divided into distinct subphases, which have been associated with specific activities and transmission events. More precisely, acquisition occurs primarily during the last subphase (II-3) of intracellular stylet punctures, whereas inoculation is achieved during the first subphase (II-1). This sequence of events
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appears to make salivation a better candidate for explaining the inoculation of nonpersistent viruses than the previously proposed egestion hypothesis. Therefore, ingestion-salivation is a viable alternative to the widely accepted ingestion-egestion hypothesis. Existing information suggests that semipersistent viruses are likely to be transmitted in an ingestion-egestion fashion. However, the amount of data available on semipersistent virus transmission is very limited, and other mechanisms cannot be excluded. The EPG technique has also provided a satisfactory explanation for the preacquisition starvation effect of nonpersistent transmission. Aphids subjected to preacquisition starvation are more likely to produce long II-3 subphases than are unstarved ones. Increased durations of subphase II-3 are correlated with increases in transmission efficiency. Therefore, this could explain why starved aphids acquire nonpersistent viruses more efficiently, although nonbehavioral factors might also be involved. With respect to egestion, it will be very worthwhile to search for a specific waveform in order to study its contribution to transmission. If it occurs, it presumably occurs within the complex, of C (stylet-path) waveforms. It has also been found that when an aphid makes consecutive intracellular punctures within the same probe, the II-3 subphase duration and number of archlets of the punctures also decreases as time progresses. It seems that once an aphid "tastes" a plant during long intracellular punctures, further cell punctures are very short, sap sampling is interrupted, and consequently, the probability of virus acquisition is dramatically reduced. Also, for a given virus-plant pathosystem, efficient vectors are more likely to produce intracellular punctures with long II-3 subphases than are inefficient ones. Finally, another contribution of the electronic monitoring technique has been the clarification of the mode of action by which insecticides and mineral oils interfere with nonpersistent virus transmission. Some insecticides, such as cypermethrin, can affect aphid behavior and reduce the transmission efficiency of nonpersistent viruses. However, this reduction is only achieved when aphid exposure to the insecticide-treated plants is long enough. Although oil application to plants can induce some behavioral changes in aphid probing, nonbehavioral factors seem to be mainly responsible for its inhibitory effect on nonpersistent virus transmission.
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21. Tjallingii, W.E (1978). Electronic recording of penetration behaviour by aphids. Entomol Exp. Appl. 24, 521-530. 22. Tjallingii, W.F. (1988). Electrical recording of stylet penetration activities. In "Aphids: Their Biology, Natural Enemies and Control" (A.K. Minks and P. Harrewijn, eds.). Vol. 2A, pp. 95-108. Elsevier, Amsterdam. 23. Tjallingii, W.F. (1985). Membrane potential as an indication for plant cell penetration by aphid stylets. Entomol. Exp. Appl 38, 187-193. 24. Harris, K.F. (1983). Stemorrhynchus vectors of plant viruses: Virus-vector interactions and transmission mechanisms. Adv. Virus Res. 28,113-139. 25. Sylvester, E.S. (1956). Beet yellows virus transmission by the green peach aphid. J. Econ. Entomol. 49, 789-800. 26. Harris, K.F. (1990). Aphid transmission of plant viruses. In "Plant Viruses" (C.L. Mandahar, ed.). Vol. 2, pp. 177-204. CRC Press, Boca Raton, FL. 27. Chalfant, R.B., and Chapman, R.K. (1962). Transmission of cabbage viruses A and B by the cabbage aphid and the green peach aphid. J. Econ. Entomol. 55, 584-590. 28. Lim, WL., and Hagedom, D.J. (1977). Bimodal transmission of plant viruses. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 237-251. Academic Press, New York. 29. Bouchery, Y., Givord, L., and Monestiez, P. (1990). Comparison of short- and long-feed transmission of the cauliflower mosaic virus: Cabb-S strain and S 11 hybrid by two species of aphid: Myzus persicae (Sulzer) diwd Brevicoryne brassicae (L.). Res. Virol. 141,677-683. 30. Markham, P.G., Pinner, M.S., Raccah, B., and Hull, R. (1987). The acquisition of a caulimovirus by different aphid species: Comparison with a potyvirus. Ann. Appl. Biol. I l l , 571-587. 31. Pirone, T.P., and Harris, K.F. (1977). Nonpersistent transmission of plant viruses by aphids. Annu. Rev. Phytopathol 15, 55-73. 32. Francki, R.I.B., Fauquet, CM., Knudson, D.L., and Brown, F. (1991). Classification and nomenclature of viruses. Fifth Report of the International Committee on Taxonomy of Viruses. Arch. Virol Suppl. 2. 33. Eastop, VF. (1983). The biology of the principal aphid virus vectors. In "Plant Virus Epidemiology" (R.T. Plumb and J.M. Thresh, eds.), pp. 115-132. Blackwell Scientific Publications, Oxford. 34. Murphy, FA., Fauquet, CM., Bishop, D.H.L., Ghabrial, S.A., Jarvis, A.W., Martelli, G.P, Mayo, M.A., and Summers, M.D. (1995). "Virus Taxonomy: The Classification and Nomenclature of Viruses. The Sixth Report of the International Committee on Taxonomy of Viruses." SpringerVerlag, Vienna. 35. Lopez-Abella, D., Bradley, R.H.E., and Harris, K.F. (1988). Correlation between stylet paths made during superficial probing and the ability of aphids to transmit nonpersistent viruses. In "Advances in Disease Vector Research" (K.F. Harris, ed,). Vol. 5, pp. 251-285. Springer-Verlag, New York. 36. Blanc, S., Lopez-Moya, J.J., Wang, R.Y., Garcia-Lampasona, S., Thombury, D.W., and Pirone, T.P. (1997). A specific interaction between coat protein and helper component correlates with aphid transmission of a potyvirus. Virology 231, 141-147. 37. Powell, G., Pirone, T.P, and Hardie, J. (1995). Aphid stylet activities during potyvirus acquisition from plants and in vitro system that correlate with subsequent transmission. Eur. J. Plant Pathol 101,411-420. 38. Collar, XL., Avilla, C , and Fereres, A. (1997). New correlations between aphid stylet paths and nonpersistent virus transmission. Environ. Entomol 26, 537-544. 39. Martin, B., Collar, J.L., Tjallingii, W.E, and Fereres, A. (1997). Intracellular ingestion and salivation by aphids may cause acquisition and inoculation of non-persistently transmitted plant viruses. J. Gen. Virol 78,2701-2705.
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40. Martin, B. (1998). Evaluacion y caracterizacion de las resistencias en melon (Cucumis melo L.) a Aphis gossypii Glover y a la transmision no persistente de vims por este vector. Ph.D. Thesis. Universidad Politecnica de Madrid. 41. Hodges, L.R., and McLean, D.L. (1969). Correlation of transmission of bean yellow mosaic virus with salivation activity of Acrythosiphon pisum (Homoptera: Aphididae). Ann. Entomol. 5'oc.^m. 62, 1398-1401. 42. Kloft, W.J. (1977). Radiosotopes in aphid research. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 291-310. Academic Press, New York. 43. Wensler, R.J., and Filshie, B.K. (1969). Gustatory organs in the food canal of aphids. J. Morphol 129, 473^77. 44. Forbes, A.R. (1969). The morphology and fine structure of the gut of the green peach aphid, Myzus persicae (Sulzer) (Homoptera: Aphididae). Mem. Entomol. Soc. Can. 36, 1-74. 45. Lopez-Abella, D., and Bradley, R.H.E. (1969). Aphids may not acquire and transmit stylet-borne viruses when probing intercellularly. Virology 39, 338-342. 46. Powell, G. (1991). Cell membrane punctures during epidermal penetrations by aphids: Consequences for the transmission of two potyviruses. Ann. Appl. Biol. 119, 313-321. 47. Montllor, C.B., and Tjallingii, W.F. (1989). Stylet penetration by two aphid species on susceptible and resistant lettuce. Entomol. Exp. Appl 52, 103-111. 48. Hogen Esch, Th., and Tjallingii, W.F. (1992). Ultra structure and electrical recording of sieve element punctures by aphid stylets. In "Proceedings of the 8th International Symposium on InsectPlant Relationships" (S.B.J. Menken, J.H. Visser, and P. Harrewijn, eds.), pp. 283-285. Kluwer Academic Publishers, Dordrecht, The Netherlands. 49. Calatayud, PA., Rahbe, Y., Tjallingii, W.F, Tertuliano, M., and Le Ru, B. (1994). Electrically recorded feeding behaviour of cassava mealybug on host and non-host plants. Entomol. Exp. Appl. 12,221-234. 50. Jiang, Y.X., Lei, H., Collar, J.L., Martin, B., Mufiiz, M., and Fereres, A. (1999). Probing and feeding behavior of two distinct biotypes of Bemisia tabaci (Homoptera: Aleyrodidae) on tomato plants. J. Econ. Entomol. 92, 357-366. 51. Wensler, R.J. (1977). The fine structure of distal receptors on the labium of the aphid Brevicoryne brassicae L. (Homoptera). Cell Tissue Res. 181, 409^22. 52. Tjallingii, W.F. (1978). Mechanoreceptors of the aphid labium. Entomol. Exp. Appl. 24,531-537. 53. Watson, M.A., and Nixon, H.L. (1953). Studies on the feeding of Myzus persicae (Sulz.) on radioactive plants. Ann. Appl. Biol. 40, 537-545. 54. Hennig, E. (1968). Uber Beziehungen zwischen dem "Probieren" der schwarzen Bohnenlaus {Aphis fabae, Scop.) und dem Stofftransport bei Viciafaba L. Arch. Pflanzenschutz 4, 75-76. 55. Chen, J.Q., Martin, B., Rahbe, Y, and Fereres, A. (1997). Early intracellular punctures of two aphids on near-isogenic melon lines with and without the virus aphid transmission (Vat) resistance gene. Eur J. Plant Pathol. 103, 521-536. 56. Lei, H., Tjallingii, W.F., Van Lenteren, J.C, and Xu, R.M. (1996). Stylet penetration by larvae of the greenhouse whitefly on cucumber. Entomol. Exp. Appl. 79, 77-84. 57. Lei, H., Tjallingii, WE, and van Lenteren, J.C. (1998). Probing and feeding characteristics of the greenhouse whitefly in association with host-plant acceptance and whitefly strains. Entomol. Exp. Appl. 88, 73-80. 58. Rezaul-Karim, A.N.M, and Saxena, R.C. (1991). Feeding behavior of three Naphotettix species (Homoptera: Cicadellidae) on selected resistant and susceptible rice cultivars, wild rice, and graminaceaous weeds. J. Econ. Entomol. 84, 1208-1215. 59. Powell, G., Harrington, R., and Spiller, N.J. (1992). Stylet activities and potato virus Y vector efficiencies by the aphids Brachycaudus helichrysi and Drepanosiphum platanoidis. Entomol. Exp. Appl. 62,293-300.
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60. Collar, XL., and Fereres, A. (1998). Nonpersistent virus transmission efficiency determined by aphid probing behavior during intracellular punctures. Environ. Entomol 27, 583-591. 61. Achon, M.A., Pinner, M. Medina, V, and Lomonossoff, G.P. (1996). Biological characteristics of maize dwarf mosaic potyvirus from Spain. £^Mr. J P/fl«^Pa^Ao/, 102,697-705. 62. Wang, R.Y., Powell, G., Hardie, X, and Pirone, T.R (1998). Role of the helper component in vector-specific tranmission of potyviruses.^ Ge«. F/ro/. 79, 1519-1524. 63. Watson, M.A. (1938). Further studies on the relationship between Hyosciamus virus 3 and the dc^hid Myzus persicae (Sulz.) with special reference to the effects of fasting. Proc. R. Soc. Lond. /S/o/.y 125, 144-170. 64. Powell, G. (1993). The effect of preacquisition starvation on aphid transmission of potyviruses during observed and electrically recorded stylet penetrations. Entomol. Exp. Appl. 66, 255-260. 65. Wang, R.Y., and Pirone, T.P. (1996). Potyvirus transmission is not increased by preacquisition fasting of aphids reared on artificial diet. J. Gen. Virol. 11^ 3145-3148. 66. Bradley, R.H.E. (1952). Studies on the aphid transmission of a strain of henbane mosaic virus. Ann. Appl. Biol 39, 78-97. 67. Bradley, R.H.E. (1964). Aphids transmission of stylet-borne viruses. In "Plant Virology" (M.K. Corbett and H.D. Sisler, eds.), pp. 148-174. University of Florida Press, Gainesville, FL. 68. Bradley, R.H.E. (1961). Our concepts: On rock or sand? Recent Adv. Bot. 1, 528-533. 69. Martin, B., and Fereres, A. (1998). Behavior oi Aphis gossypii and Myzus persicae on resistant and susceptible Cucumis melo L. lines. In "Aphids in Natural and Managed Ecosystems" (XM. Nieto Nafria and A.F.G. Dixon, eds.), pp. 97-103. Universidad de Leon (Secretariado de Publicaciones). Leon, Spain. 70. Cole, R.A. (1997). Comparison of feeding behaviour of two Brassica pests Brevicoryne brassicae and Myzus persicae on wild and cultivated brassica species. Entomol. Exp. Appl. 85, 135-143. 71. Berger, PH., Zeyen, R.X, and Groth, XV (1987). Aphid retention of maize dwarf mosaic virus (potyvirus): Epidemiological implications. Ann. Appl. Biol. I l l , 331-344. 72. Zeyen, R.X, and Berger, P.H. (1990). Is the concept of short retention times for aphid-borne nonpersistent viruses sound? Phytopathology 80, 769-771. 73. Fereres, A., Blua, M., and Perring, T. (1992). Retention and transmission characteristics of zucchini yellow mosaic virus by Aphis gossypii and Myzus persicae (Homoptera: Aphididae). J. Econ. Entomol. 85, 759-765. 74. Burt, P.E., Heathcote, G.D., and Broadbent, L. (1964). The use of insecticides to find when leaf roll and Y viruses spread within potato crops. Ann. Appl. Biol. 54, 13-22. 75. Rains, B.D., and Christensen, C M . (1983). Effect of soil-applied carbofuran on transmission of maize chlorotic dwarf virus to susceptible field corn hybrid. J. Econ. Entomol. 76, 290-293. 76. Ferro, D.N., Mackenzie, XD., and Margolies, D.C. (1980). Effect of mineral oil and systemic insecticide on field spread of aphid-borne maize dwarf mosaic virus in sweet corn. J. Econ. Entomol. 73, 730-735. 77. Simons, XN. (1957). Effects of insecticides and physical barriers on field spread of pepper veinbanding mosaic virus. Phytopathology 47, 139-145. 78. Broadbent, L., Burt, P.E., and Heathcote, G.D. (1956). The control of potato virus diseases by insecticides. Ann. Appl. Biol. 44,256-273. 79. Rieckmann, W (1991). Zur Problematik der Eingrenzung nichtpersistenter Viren und ihrer Vektoren im Pflanzkartoffelbau. Gesunde Pflanzen 43, 155-159. 80. Reagan, T.E., Gooding, G.V, and Kennedy, G.G. (1979). Evaluation of insecticides and oil for suppression of aphid-borne viruses in tobacco. J. Econ. Entomol. 72, 538-540.
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81. Jayasena, K.W., and Randies, J.W. (1985). The effect of insecticides and a plant barrier row on aphid populations and the spread of bean yellow mosaic potyvirus and subterranean clover red leaf luteovirus in Viciafaba in Australia. Ann. Appl Biol. 107, 355-364. 82. Briggs, G.G., Elliott, M., and Famham, A.W. (1974). Structural aspects of the knockdown of pyrethroids. Pesticide Sci. 5, 643-649. 83. Wouters, W., and Van den Bercken, J. (1978). Action of pyrethroids. Gen. Pharmacol. 9,387-398. 84. Gibson, R.W., Rice, A.D., and Sawicki, R.M. (1982). Effects of the pyrethroid deltamethrin on the acquisition and inoculation of viruses by Myzus persicae. Ann. Appl. Biol. 100,49-54. 85. Gibson, R.W., and Campbell, C.C. (1986). Investigations into how cypermethrin controls the spread of potato viruses by aphids. In "Proceedings, 1986 British Crop Protection Conference— Pests and Diseases," pp. 997-1000. Brighton, U.K. 86. Argandona, VH., Corcuera, L.J., Niemeyer, H.M., and Campbell, B.C. (1983). Toxicity and feeding deterrency of hydroxamic acids from Gramineae in synthetic diets against the greenbug, Schizaphis graminum. Entomol. Exp. Appl. 34, 134—138. 87. Avilla, C , Collar, J.L., Arretz, P., and Fereres, A. (1993). Feeding behaviour and mortality of Myzus persicae (Homoptera: Aphididae) on pepper plants treated with systemic insecticides. Entomology (Trends inAgr. Sci.) 1, 23-29. 88. Hardie, J., Holyoak, M., Taylor, N.J., and Griffiths, D.C. (1992). The combination of electronic monitoring and video-assisted observations of plant penetration by aphids and behavioural effects of polygodial. Entomol. Exp. Appl. 62, 233-239. 89. Woodford, J.A.T., and Mann, J.A. (1992). Sistemic effects of imidacloprid on aphid feeding behaviour and virus transmission on potatoes. In "Proceedings, 1992 British Crop Protection Conference—Pests and Diseases," pp. 229-234. Brighton, U.K. 90. Powell, G., Hardie, J., and Pickett, J.A. (1993). Effects of the antifeedant polygodial on plant penetration by aphids assessed by video and electrical recording. Entomol. Exp. Appl. 68,193-200. 91. Harrewijn, P., and Piron, P.G.M. (1994). Pymetrozine, a novel agent for reducing virus transmission by Myzus persicae. In "Proceedings, 1994 British Crop Protection Conference—^Pests and Diseases," pp. 923-928. Brighton, U.K. 92. Givovich, A., and Niemeyer, H.M. (1995). Comparison of the effect of hydroxamic acids from wheat on five species of cereal aphids. Entomol. Exp. Appl. 74, 115-119. 93. Nauen, R. (1995). Behaviour modifying effects of low systemic concentrations of imidacloprid on Myzus persicae with special reference to an antifeeding response. Pesticide Sci. 44,145-153. 94. Collar, J.L., Avilla, C , Duque, M., and Fereres, A. (1997). Behavioral response and virus vector ability of Myzus persicae (Homoptera: Aphididae) probing on pepper plants treated with aphicidQS. J. Econ. Entomol. 90, 1628-1634. 95. Bradley, R.H.E., Wade, C.V, and Wood, FA. (1962). Aphid transmission of potato virus Y inhibited by oils. Virology 18, 327-329. 96. Bradley, R.H.E., Moore, C.A., and Pond, C.C. (1966). Spread of potato virus Y curtailed by oil. Nature (Lond.) 209, 1370-1371. 97. Vanderveken, J. (1968). Effects of mineral oil and lipids on aphid transmission of beet mosaic and beet yellows viruses. Virology 34, 807-809. 98. Simons, J.N., and Zitter, T.A. (1980). Use of oils to control aphid-borne viruses. Plant Dis. 64, 542-546. 99. Szatmari-Goodman, G., and Nault, L.R. (1983). Tests of oil sprays for supression of aphid-borne maize dwarf mosaic virus in Ohio sweet com. J. Econ. Entomol. 76, 144-149. 100. Gibson, R.W, and Rice, A.D. (1986). The combined use of mineral oils and pyrethroids to control plant viruses transmitted non- and semipersistently by Myzus persicae. Ann. Appl. Biol. 109, 465^72.
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101. Lowery, D.T., Sears, M.K., and Harmer, C.S. (1990). Control of turnip mosaic virus of rutabaga with applications of oil, whitewash and insecticides. J. Econ. Entomol. 83,2352-2356. 102. Lowery, D.T., Eastwell, K.C., and Smirle, M.J. (1997). Neem seed oil inhibits aphid transmission of potato virus Y to pepper. Ann. Appl. Biol. 130, 217-225. 103. Webb, S.E. (1993). Effect of oil and insecticide on epidemics of pot5rviruses in watermelon in Florida. Plant Dis. 11, 869-874. 104. Qiu, J.Y., and Pirone, T.P (1989). Assessment of the effect of oil on the potyvirus aphid transmission process. J. Phytopathol 121,221-226. 105. Wang, R.Y., and Pirone, T.P. (1996). Mineral oil interferes with retention of tobacco etch potyvirus in the stylets oi Myzus persicae. Phytopathology 86, 820-823. 106. Simons, J.N., McLean, D.L., and Kinsey, M.G. (1977). Effects of mineral oil on probing behavior and transmission of stylet-borne viruses by Myzus persicae. J. Econ. Entomol. 70, 309-315. 107. Powell, G. (1992). The effect of mineral oil on stylet activities and potato virus Y transmission by aphids. Entomol. Exp. Appl. 63, 237-242. 108. Collar, J.L., and Fereres, A. (1999). Mineral oil effect on aphid probing behavior and on PVY acquisition from pepper plants. In "Proceedings, 8th International Plant Virus Epidemiology Symposium," p. 43. Almeria, Spain. 109. Ammar, E.D., and Nault, L.R. (1991). Maize chlorotic dwarf virus-like particles associated with the foregut in vector and non-vector leafhopper species. Phytopathology 81,444-448. 110. Gemeno, C. (1992). Preference and feeding behavior of the blackfaced leafhopper Graminella nigrifrons (Homoptera: Cicadellidae): Implications in maize chlorotic dwarf virus transmission. M.S. Thesis. University of Kentucky, Lexington, KY 111. Wayadande, A.C., and Nault, L.R. (1993). Leafhopper probing behavior associated with maize chlorotic dwarf virus transmission to maize. Phytopathology 83, 522-526. 112. Harris, K.F., and Childress, S.A. (1983). Cytology of maize chorotic dwarf virus infection in com. Int. J. Trop. Plant Dis. 1,135-140. 113. McLean, D.L., and Kinsey, M.G. (1984). The precibarial valve and its role in the feeding behavior of the pea aphid, Acyrthosiphon pisum. Bull. Entomol. Soc. Am. 30,26-31. 114. Triplehom, B.W, Nault, L.R., and Horn, D.J. (1984). Feeding behavior of Graminella nigrifrons (¥oThQs). Ann. Entomol. Soc. Am. 11, 102-107. 115. Wayadande, A. (1992). Studies in leafhopper probing behavior and its role in MCDV transmission. Ph.D. Dissertation. The Ohio State University, Columbus, OH. 116. Spotti Lopes, J.R. (1996). Feeding behaviour of xylem-feeding leafhopper vectors of Xylella fastdiosa: Present knowledge and research questions. 3rd EPG Workshop, Lyon, France, Sept. 1-6, 1996.
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CHAPTER 6
Ingestion-Egestion Theory of Cuticula-Borne Virus Transmission KERRY F. HARRIS LISA JEAN HARRIS
/. Introduction Noncirculative plant viruses, recently also dubbed cuticula-borne [1-4], and their aphid vectors have been studied longer and by more scientists than any other plant virus-vector system. Researchers have always been duly impressed with the brevity and seeming simplicity of nonpersistent transmission of cuticula-borne viruses by aphids. In fact, the essence of that simplicity has fascinated, challenged, and ultimately eluded plant pathologists, entomologists, virologists, and of late, molecular biologists for over seven decades. Their exceptional fecundity compensates aphids for their small, delicate bodies and vulnerability to attack by a prodigious number of parasites and predators. Dense populations cause extensive crop losses by sucking sap from phloic cells, especially sieve cells and tubes. Small populations can devastate crops with potentially phytotoxic substances in the saliva secreted during probing and feeding. Direct or indirect crop damage related to honeydew secretion can also be costly. Add virus transmission to this already infamous repertoire, as well as worldwide geographic and climatic distribution and expansive host ranges and voild! one of the most formidable animals ever to thwart human efforts to grow food and fiber. Homopteran-bome plant viruses are either noncirculative or circulative [5]. Aphids acquire and inoculate noncirculative viruses via the maxillary food canal and can inoculate them to plants immediately after acquisition. These viruses are retained at putative aphid cuticular receptors (ACRs) on the lining of the feedingVirus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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apparatus lumen and hence are referred to as cuticula-borne [6, 7]. One can subcategorize noncirculative or cuticula-borne viruses as either nonpersistent or semipersistent based on numerous discernible transmission characteristics [5]. Nonprobing viruhferous aphids, such as alatae carried in air currents, can retain inoculativity for more than a day [8-10] and carry their "nonpersistent" payload over many miles. Starved aphids are also more efficient transmitters of virus [11]. Alatae are analogous to the long-range stealth bombers of our strategic air command. These flying, biological, precision microsyringes can alight in a crop and deliver their "smart bombs" before being gunned down by insecticide, an epidemiologist's worst nightmare. The resulting primary infection foci then serve as sources of virus for secondary in-field disease spread by alatae and apterae. An overwhelming majority of all insect vectors of plant viruses is in the order Homoptera. Aphididae contains more vector species of noncirculative viruses than the other homopteran families combined. Aphids reign supreme as nonpersistent transmission specialists. The green peach aphid, Myzus persicae Sulzer, alone transmits over 100 different viruses, most of them nonpersistent. The genus Potyvirus contains more nonpersistent viruses than other plant virus genera combined. It is therefore little wonder that researchers traditionally choose this virus-vector combination in experiments designed to elucidate the mechanism(s) of nonpersistent transmission. Cucumber mosaic virus (CMV) and Aphis gossypii are the traditional workhorses in research aimed at elucidating the mechanism of non-helper-assisted, nonpersistent transmission (see chapter 9). The most thoroughly studied semipersistent, noncirculative transmission is that of cauliflower mosaic virus (CaMV) by the aphids M persicae and Brevicoryne brassicae (see chapter 8). Unless otherwise noted, our comments herein mainly refer to aphids and potyviruses, usually M persicae and either potato virus Y or tobacco etch virus.
//. Terminology Those who study plant virus-vector interactions traditionally use incorrect terminology to describe the homopteran feeding apparatus. This is not surprising, in as much as most of the pioneering researchers in this specialty were plant pathologists, a fact that remains true today. We are referring to terms such as foregut, anterior foregut, fore-alimentary canal, anterior alimentary canal, and pharyngeal pump. Most of us johnny-come-latelies were aware of the problem but continued using the terminology, by now well established in the literature, for the sake of continuity. In 1994, Ammar et al [12] became aware of the problem and attempted to correct it by redefining the foregut "to include the esophagus, cibarium (including the pharynx), and the precibarium" and referring to the walls of the precibarium and cibarium as "intima." However, defining preoral structures such as precibarium and cibarium as postoral (i.e., parts of the foregut) and post-
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oral ones such as pharynx and intima as preoral only exacerbated the problem. The pharynx is not part of the cibarium and the cibarium is not part of the foregut. The homopteran feeding apparatus is a preoral structure whose contiguous food conduit extends from the distal common (food-saliva) duct to the maxillary food canal, antecibarium, cibarial valve, and postcibarium. The latter empties through the OS or true mouth into the pharynx of the foregut. The foregut and hindgut of aphids are derived from invaginations of the exoskeleton and hence lined by specialized cuticula called intima. The homopteran feeding apparatus comprises six exoskeletal lobes from the venter of the head: clypeolabrum anteriorly, paired lora anterolaterally, paired maxillary plates posterolaterally, and hypopharynx posteriorly [6, 7]. Another lobe, the labium, emerges from the median of the cervical venter. The cibarial valve divides the cibarium, mainly formed by the apposition of the epicibarium and hypocibarium, into antecibarium and postcibarium [6, 7]. The latter functions as the cibarial pump. The cuticular nature of the cibarium and the origin of its valve and postcibarial retractor muscles from the anteclypeus and postclypeus, respectively, attest to its evolutionary linkage to the preoral cibarium of the primitive orthopteroid cibarium. The os connecting postcibarium to intimalined, cellular pharynx coincides with the frontoclypeal suture and the distalmost descent of the frontal ganglion [6, 7]. Similarities among aphid, whitefly, and leafhopper feeding apparatuses confirm the ingestion-egestion theory (IET): virus is acquired by ingestion, carried internally on the cuticular lining of the feeding-apparatus lumen (cuticula-borne), and inoculated to plants by egestion. Vector researchers have generally accepted the I-ET since its introduction [13, 14]. (Our upgrading the ingestion-egestion model of nonpersistent transmission from hypothesis to theory is based on its withstanding 27 years of scrutiny and testing.) The postcibarium operates similarly to a reversible bellows. Contraction of the distal retractor muscles with the cibarial valve open and the pump closed posteriorly results in negative pump pressure and active ingestion. Conversely, a wavelike proximodistal relaxation of the retractor muscles with the cibarial valve open and the pump closed posteriorly results in positive pump pressure and egestion. A wavelike distoproximal relaxation of the pump retractors with the cibarial valve closed results in active "swallowing" of fluid from the postcibarium past the os into the pharynx of the foregut [6, 7]. Aphids control flow of sap from sieve elements. The small entrance to the maxillary food canal is a natural reduction valve. Additionally, the cibarial valve aperture can be varied between fully open and fully closed. The postcibarial retractor muscles must be sufficiently contracted to effect negative pressure in the pump relative to turgor pressure, even during socalled passive [15] ingestion in sieve elements. Relaxation of the pump retractor muscles and the natural inward collapse of the epicibarium with the cibarial valve open and the postcibarium closed posteriorly produce a force greater than turgor pressure. If the latter were not so, aphids, leafhoppers, and whiteflies would be
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unable to egest and therefore unable to inoculate plants with cuticula-borne viruses—^but of course, they can and do. The aphid salivary syringe, designed to receive saliva from the salivary glands and excrete it onto or into plants, is essentially the same as that described in whiteflies [6, 7]. A highly specialized development of the salivarium of orthopteroid insects, it consists of a strong exoskeletal invagination that is securely anchored to the front wall of the hypopharynx. The syringe with its efferent duct has a tulipshaped profile because when the piston retractor muscles are relaxed, the elastic piston (cuticular roof of the cupula) returns to its "normal" position following the concave contour of the cupula floor. The efferent duct leading to the maxillary food canal can be likened to the tulip's stem. The syringe is similar to a hypodermic syringe in both form and function [2]. Its chamber or cupula is analogous to the wide body of the hypodermic syringe, its roof or piston to the hypodermic's plunger, and the efferent duct and contiguous maxillary saliva canal to the hypodermic's narrow stem and attached needle, respectively. Retracting or lowering the piston creates negative or positive pressure within the cupula, respectively. Retracting the piston by contracting the piston retractor muscles creates vacuum that sucks saliva from the open afferent duct (cupula muscles relaxed) into the cupula. Subsequent contraction of the cupula muscles (afferent duct closed) and simultaneous relaxation of the piston muscles creates positive pressure to eject saliva from the cupula through the efferent duct into the salivary canal of the maxillae. The axes of rotation of the cupula sclerites are such that contraction of the cupula muscles causes the sclerites' medial margins to collapse inward, effectively closing the afferent duct aperture between them. Closure of the afferent duct aperture while the piston is lowered prevents back flow of saliva into the afferent duct. The piston angle and the insertion of piston muscles allow the back of the piston to rise while the front margin remains at rest over the opening to the efferent duct, effectively sealing it and preventing back flow. Hence the piston seemingly functions both as piston and as efferent-duct valve when retracting to enlarge the cupula, create vacuum, and draw saliva from the afferent duct. Misnomers used to describe the morphology of the homopteran feeding apparatus have carried over to virus transmission terminology. Some researchers use "foregut-bome" [12, 16] to define where and how aphids retain noncirculative viruses. However, an analysis of the homopteran feeding apparatus indicates that virions held at sites posterior to the cibarium (i.e., in the pharynx of the foregut) are nonegestible and hence nontransmissible [6, 7]. Others have reverted to referring to nonpersistent viruses as "stylet-borne" {sensu Kennedy et al. [17], carried at the maxillary tips). However, this reversion is premature unless, sensu stricto, a particular virus is demonstrably carried exclusively at the stylet tips or, sensu lato, exclusively in the maxillary food canal. To date, no such cases are known. In 1984, McLean and Kinsey [18] rediscovered the cibarial valve described by Davidson [19] in 1914 and Weber [20] in 1928 and concluded that aphids are anatomically incapable of egesting. McLean and Kinsey [18] further suggested
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that aphids inoculate noncirculative viruses when valve malfunctioning allows previously ingested virus to exit the feeding apparatus by "extravasation." Extravasation, a highly specialized term in medical pathology, refers to leakage of fluids, particularly blood, from vessels into surrounding intercellular spaces. It also links cuticula-bome virus inoculation to an abnormal, unpredictable, passive, nonpurposeflil, vector activity. Virus transmission, however, is a consequence of the normal, predictable, active, purposeful, vector activities of salivation, ingestion, and egestion. Obviously aphids [13], leafhoppers [21], and, by morphological similarity, whiteflies can and do actively salivate, ingest, and egest. The fact that aphids can ingest or egest for minutes at a time [13] indicates that they can sustain negative or positive pressure, respectively, within the feeding-apparatus lumen. The postcibarium is equally well equipped to induce ingestion or egestion [6, 7]. Nevertheless, a few researchers [22] persist in referring to noncirculative virus inoculation as occurring by extravasation rather than egestion. In this chapter, we follow the feeding-apparatus and transmission terminologies recommended by Harris and associates [6, 7].
///A.
Mechanism of Nonpersistent Transmission
Mechanical Contamination Hypotheses of the First Four Decades
Mechanical, inactivator, inactivator-behavior, stylet-plug, saliva-plug, mechanicalsurface adherence, modified mechanical, and mechanical-inactivator-behavior-compatibility are all descriptive titles of pioneering hypotheses of the mechanism of nonpersistent transmission. The hypotheses differ among themselves in how they explain nonpersistence, specificity, the importance of brief probes, the preacquisition starvation effect [11], and how, when, and where virus contacts, adheres to, and detaches from the stylets. All, however, are based on the assumption that virus is carried by the stylets in an essentially mechanical manner as proposed by Doolittle and Walker in 1928 [23]. This commonality among early hypotheses is understandable, in view of what researchers of the time knew or thought they knew about brief, superficial probing by aphids. Light microscopy revealed that aphids begin such probes by secreting a salivary flange in anticlinal grooves between adjacent epidermal cells. The flange continues as salivary sheath in the epidermis and sometimes in underlying mesophyll (again seemingly between cells) until stylet-bundle withdrawal. Occasionally, a daredevil would brazenly suggest that aphids must somehow ingest plant fluid during brief probes. However, such renegade thinking was quickly silenced by yet another dogma: Aphids cannot egest. Then, back we went to reshaping "round" data to fit the "square" mechanical hypothesis. In 1955, Bradley and Ganong [24, 25] implicated the maxillae as carriers of virus. Exposing the terminal 5-10 |im of the maxillae of viruliferous aphids to
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various chemical and physical (especially ultraviolet irradiation) antiviral treatments rendered them aviruliferous. Kennedy et al [17] labeled this more definitive mechanical hypothesis "stylet-borne." These findings [24, 25] would later take on new and different significance. B.
Ingestion-Egestion Theory
In 1969, electron microscopy of salivary sheaths revealed that at some point at least some superficial probes become intracellular [26]. (A later, more comprehensive study [27] would confirm what these data suggested, that transmission is an intracellular event.) In 1973, light microscopic observations of aphids feeding through membranes indicated that aphids routinely egested when, following prolonged periods of ingestion, carbon particles suspended in a feeding solution hampered fluid flow into the food canal [13]. Aphids free clogs by egesting in various combinations and sequences with other activities: stylet manipulation of the particles, watery saliva (Sw) excretion, ingestion, and egestion. Egestion is a natural, purposeful component of aphid (Homoptera: Stemorrhyncha) [13], leafhopper (Homoptera: Auchenorrhyncha) [21] and, based on their morphologically similar feeding apparatus and ability to transmit semipersistent cuticula-bome viruses, whitefly (Homoptera: Stemorrhyncha) [6, 7] feeding. Indeed, the fact that homopteran and hemipteran cibaria share a common plan indicates that members of both orders can egest. The aforementioned discoveries helped elucidate when, where, and how cuticula-bome vims transmission occurs. Aphids are cold-blooded; however, these gentle, cooperative, research animals are not malicious flying needles, spreading disease and death. Aphids probe to sample cytoplasm to determine host suitability. Unfortunately, as a consequence of host selection behavior, they can also acquire and inoculate nonpersistent plant viruses by ingestion and egestion; this is the ingestion-egestion theory (I-ET). Inoculated cells must survive for an infection site to develop. During host selection probes, aphids insert about 0.05 |Lim of the tips of the maxillae [28] into cells; secrete, ingest, and egest Sw; ingest, taste, and egest miniscule (compared to total cell volume) aliquots of C5^oplasm; and plug cell-entry sites and the salivary sheath canal with gelling saliva while withdrawing their stylet bundle [5]. These behavioral features make aphids ideal vectors of nonpersistent, cuticula-bome vimses. C.
Ingestion-Salivation Hypothesis
Proponents of the ingestion-salivation hypothesis (I-SH) [15, 29-31] claim that aphids acquire nonpersistent vimses by ingestion, carry them in the common (food-saliva) duct in the terminal 1-3 |Lim [28] of the maxillae, and subsequently inoculate them by saliva excretion. These proponents [15, 29-31] defend the common duct as the site of virus retention with statements such as: The I-SH is based
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on the fact that aphids inoculate nonpersistent viruses before any ingestion can take place [30]. Martin et al [29] state that a model in which egestion through the [maxillary] food canal injects virus into the plant [the I-ET] does not take account of the anatomical fact that the food and salivary canals in the maxillary stylets fuse at 2-8 jam from the tips. Tjallingii and Prado [15] make the same statements. In point of fact, the existence of the common duct in the terminal 2-A |im of the tips of the maxillae was a well- and long-established anatomical fact [28, 32, 33] when the I-ET was introduced [14]. Moreover, the common duct region of the maxillae received considerably more consideration than other regions of the maxillae before being dismissed as an unwholesome, most unlikely site for virus retention [14]. As we will soon demonstrate, there is good reason to believe that aphids do ingest before any virus inoculation takes place. Furthermore, the relative sequence of events with respect to ingestion and egestion or virus acquisition and inoculation or both has no significance in proposing or supporting any mechanism of nonpersistent virus transmission or in associating electrically recorded waveforms produced by plant-probing aphids with any particular aphid activity.
IV, Site of Virus Retention A.
Common Duct
Proponents of the I-SH do not refute the overwhelming, unequivocal evidence that both nonpersistent and semipersistent cuticula-borne viruses are retained proximal to the entrance to the maxillary food canal [12, 22, 34-38]. In the case of nonpersistent viruses, however, they [21,29-31] dismiss the confirmatory data based on the assumption that egestion is not a part of aphid host-selection behavior. They defend this latter assumption with the following statements: "The sequence of events in EPGs makes salivation a better candidate for inoculation than egestion, because egestion seems to occur after ingestion during prolonged aphid feeding on artificial systems [13]" [30]; "Moreover, it seems that aphids need to ingest first to determine host suitability" [30]; "Why should an aphid egest any sap material before sampling a new host? [30], and finally: "Recently, radiolabelled pot3rvirus particles were found to be retained especially in the distal third of the maxillary stylets (38) (i.e., stylet tips)" [29]. We will respond to these statements in reverse order. First, for Martin et al [29] to equate the 140 |im-long distal third of the maxillary food canal with the 1- to 3-|im-long common duct [28] is more than a minor stretch. Second, we respond to their rhetorical query with two of our own. Why would an animal capable of both salivation and egestion not utilize these abilities to wet and clean its feeding apparatus and gustatory sensilla of residue from a previous feeding or tasting? Furthermore, why would an aphid pass through its
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entire digestive system any plant fluid or artificial feeding solution that it has either not yet discerned as appropriate for ingestion or has discerned as distasteful or potentially harmful? Third, we think that aphids need to first secrete, ingest, and egest saliva before ingesting cytoplasm to determine host suitability. Fourth, superficial probing and phloem feeding are very different processes seeking totally different goals—host selection versus uninterrupted feeding, respectively. If anything, therefore, we would expect their "sequence of events" [29, 30] to be also different. Virus retained in the common duct is susceptible to drying out and possible physical (flushing) and chemical side effects of saliva secretion. Aphid saliva is known to contain pectin (middle lamella)- and cellulose (cell wall)-hydrolyzing enzymes, as well as phenols, phenol oxidases, and amine-loving quinones [14]. Aphids seal cell-puncture sites and the saliva-sheath lumen with gelling saliva at the end of a probe. They also initiate probes by excreting flange and sheath saliva. This represents volumes of saliva that flush the 1- to 3-|Lim long [28] common duct thousands of times over. Nonetheless, transmission occurs. In contrast to the vulnerability of virions presumably retained in the common duct, virions retained in the feeding apparatus proximal to the entrance to the maxillary food canal are out of harm's way. Artificially suspending viruliferous aphids stimulates them to excrete fine threads of gelling saliva and droplets of watery saliva (Sw). However, this salivation does not affect their inoculativity or rate of virus inoculation [39], a surprising fact given that, according to I-SH proponents, saliva excretion is what flushes virions from the common duct and inoculates them to cells. However, the story is different for viruliferous aphids allowed to probe glass "where aphids cannot ingest anything save possibly their own saliva" [39]. After a brief period on glass, aphids lose inoculativity at a rate comparable to that of aphids transferred directly from virus source plant to healthy test plant. As we will later show, simultaneous secretion and ingestion of Sw followed by egestion are natural components of the pregustation phase of host selection probes, ultimately effecting the release of virions from putative, aphid cuticular receptors (ACRs) and their inoculation to cells. This pregustation behavior also explains why virions can be seen in saliva deposited on glass by viruliferous aphids [40]. B.
Maxillary Food Canal and Cibarium
To taste a plant, aphids ingest cytoplasm (a "sap sample") through the maxillary food canal to gustatory sensilla in the vicinity of the cibarial valve [6, 7]. Cuticula-bome viruses have customized their conveyance system with methods for recognizing and clinging to their aphid carriers. Most, such as potyviruses [41] and caulimoviruses [42], produce viral encoded, proteolytic, helper protein(s) (HC), which form bridges linking putative ACRs to the N-terminal portion of the viral capsid-protein subunit (CPS). For cucumovimses [43], a conserved domain of the CPS, the pH-|3I loop.
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seemingly facilitates virion binding directly to ACRs. One assumes that ACR-HCCPS or ACR-CPS associations occur wherever ACRs are present and contact the surface of cytoplasm ingested from an infected plant cell containing compatible HC and CPS: common duct, maxillary food canal and cibarium, particularly the antecibarium. This assumption is well founded [12,22, 34-38]. Wang et al [38] allowed starved aphids to probe through Parafilm membranes into feeding solutions containing transmissible or nontransmissible virus-helper combinations. Immunogold-transmission electron microscopy (TEM) or autoradiography served to localize ACR-associated virions or label, respectively, in the feeding apparatus lumen. Wang et al. [38] observed gold-labeled virions in the distal part of the food canal in 56.0% of aphids examined. Or, from a different perspective, virions occurred in the proximal part of the food canal (34.6%) or in the cibarium (12.0%) of 46.6% of aphids examined. In the stylet-limited, autoradiography (isotopically labeled virions) experiments [38], they detected label in the distal third of the maxillae in 82% of the aphids, either exclusively or in combination with the central or proximal third. Or, again from a different perspective, over 50% of the aphids had label solely in the central or proximal; solely in the central and proximal, in the central or proximal third in combination, either singly or jointly, with the distal third; or evenly distributed throughout the maxillae. Analyses of these and other data [12, 22, 34-38] indicate that virus is retained in all areas where one expects contact among ACRs, HC, and CPSs. The most helpful aspect of the aforementioned immunogold-TEM and autoradiography experiments [38] is their design. Thanks to excellent controls, one can reasonably conclude the following: (1) ACR-HC-CPS reciprocity ("bridging" [41, 42]) is essential to virion retention. (2) Naturally terminated 1- to 2-minute probes on membranes mimic those on plants with respect to aphid host-selection behavior. (3) At the end of naturally terminated brief probes, feeding-solution samples (or cytoplasm, in plant probing) and non-ACR-associated virions are removed from the transmission-associated portion of the feeding-apparatus lumen prior to stylet bundle withdrawal. (4) Sample removal occurs mainly by sample egestion (preacceptance aphids) or by sample ingestion beyond the cibarial valve (postacceptance aphids). (5) The data, taken together, also suggest that virus inoculation occurs before acquisition; otherwise, aphids would either be unable to acquire and transmit virus or would do so much less efficiently, possibly not efficiently enough to ensure survival of the virus being transmitted. Data from recent electrical penetration graph (EPG) analyses of superficial probes confirm the latter suspicion [29, 30, 44, 45]. Wang et al [38] noted a bias toward virion retention in the distal portion of the feeding apparatus lumen: maxillary food canal versus cibarium, and distal versus proximal food canal. This bias reflects the nature of host selection behavior and the morphology of the feeding apparatus lumen. In preacceptance aphids, the surface of ingested cytoplasm contacts ACRs both coming (ingestion) and going (egestion). The diameter of the maxillary food canal increases from about 0.35
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|Lim distally to about 1.2 jiim proximally. The antecibarium is about 25 |Lim long, and its diameter increases from about 1.2 |Lim distally to about 2.0 |Lim proximally. The lumen diameter of a fully open cibarial valve is about 5 |im, as is the distal postcibarium. (The central postcibarium can expand to over 20 |im when open but narrows to about 3 |im, when open, at its proximal end.) In smaller-diameter, distal portions of the feeding-apparatus lumen, there is greater contact between ACR and sample surface and hence greater likelihood of successful (retentionwise) ACR-HC-CPS encounters. The latter can be conceptualized by imagining a roadway (the feeding-apparatus lumen) that begins as a three-lane highway and then broadens first into a four-lane freeway and then into a five-lane turnpike. Imagine too that people with paint brushes (ACRs) are standing alongside the inner and outer lanes of the road, marking cars (virions) with paint as they pass. A given "volume" of bumper-to-bumper traffic passes through the roadway. Only cars in the middle lane of the three-lane highway pass paint free (no ACR encounters and no retention). However, cars in the two middle lanes of the freeway and three middle lanes of the turnpike pass paint-free (no ACR contacts). Thus, one expects random sampling [38] to result in a distal bias with respect to the number of ACR-associated virions observed or detected. However, the existence of this bias is probably moot, since virions from ACRs in various portions of the feeding-apparatus lumen mix in a watery saliva (Sw) potpourri (Fig. 1, drawing Kh-la) prior to being inoculated to a cell (drawing KhIb) in the pregustation phase of host selection. Presumably, the 1.19% of aphids with label evenly distributed throughout their maxillae [38] converted to postacceptance status (less or no sample egestion) during their 10-minute virus acquisition feeding period. Moreover, they converted early enough to ingest a sufficient volume of sample through the feeding-apparatus lumen to saturate available ACRs with labeled virions.
y. Electrical Penetration Grapti Analysis A.
Electrical Penetration Graph System
Researchers can correlate recorded waveforms with stylet location and aphid activity by using a direct-current electrical penetration graph (EPG) system [15, 29, 30]. Cell penetration by the maxillae establishes a better electrical connection between aphid and plant, resulting in increased current flow and decreased potential, that is, a potential drop (pd). One can relate pd waveforms during host selection to aphid activities by studying the characteristics of waveforms in pd recordings (Fig. 1). One can determine when inoculation takes place [29] by interrupting probes of healthy test plants by individual viruliferous aphids at various time intervals in the pd, observing the test plants for symptom expression.
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EPG
INGESTION-SALIVATION HYPOTHESIS pd subphase: (activity) Transmission event:
pd phase : (activity)
< <-
<
- II-l (salivation)
II-2 -
->^<X
(?)
>i< (ingestion)
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I I
inoculation
- '<- acquisition —• <
(pre-gustation)
pd subphase: , t ^ - % ^ ^ . ^ (activity) (salivation/mgestion^
, Kh-lb (egestion)
Transmission event:
inoculation > l < acquisition >'
dissociation
Kh-2b (pre-acceptance egestion) acquisition
INGESTION-EGESTION THEORY
Fig. 1 A 5.5-s potential drop (pd) produced by Myzus persicae Sulzer. The pd is typical of initial (preacceptance), superficial host selection probes by efficient aphid vectors following preacquisition starvation. Data presented above and below the pd pertain to the ingestion-salivation hypothesis (ISH, see chapter 5) and ingestion-egestion theory (I-ET) of nonpersistent transmission, respectively. (The reader is referred to Table I and the text for a comprehensive presentation of the major differences between the I-SH and I-ET.) The letters EPG (electrical penetration graph) at the top left of the figure indicate the recording potential level before and after the pd or intracellular portion of the probe. Proponents of the I-SH [15, 29-31] subjectively divide (vertical dash-dash lines) the pd into subphases II-l, II-2, and II-3, based on the sequence of both transmission events during the pd (virus inoculation before acquisition) and ingestion-egestion events during membrane feeding (ingestion before egestion [13]). In the I-ET, the pd is divided (solid, vertical line) into pregustation (Kh-1) and gustation (Kh-2) phases based on the aphid's host selection needs and comparative examination of pd waveform attributes such as frequency and amplitude. Similar intraphase comparisons of waveforms, subdivide (vertical dash-dot-dash lines) phases Kh-1 and Kh-2 into subphases or waveform types: Khla and Kh-lb, and Kh-2a and Kh-2b, respectively. Drawings at the bottom of the figure are representative of aphid activities associated with individual pd waveforms or subphases. The drawings presume that the pd is one produced by a viruliferous aphid making an inoculation probe on a virusfree plant. Abbreviations: CW = cell wall; Cy = cytoplasm; Cy/Sw = Cytoplasm/watery saliva (Sw) mixture; F = maxillary food canal; Md = mandible; Mx = maxilla; P = plasmalemma; Ss = gelled sheath saliva. Arrows in drawings show the direction of fluid flow (Modified in part from Martin et ah [29] and Fereres and Collar [30].)
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and testing those plants symptomatic of infection for the presence of virus. Conversely, one can determine when acquisition takes place by similarly interrupting individual aviruliferous test aphids probing a virus source plant and then testing their inoculativity on healthy test plants. B.
Potential Drop Analysis According to the I-SH
The EPG in Figure 1 includes a recording of a 5.5-second potential drop (pd) typical of initial host selection probes by prestarved, efficient aphid vectors. The pd typifies ones that result in optimal virus acquisition and transmission. The EPG lettering in the upper left of the figure marks the potential level of the recording before and after the pd. Data posted above or below the pd pertain to the I-SH or I-ET, respectively. Dashed vertical lines indicate where in the pd (see section V. A) aphids inoculate (left line) and acquire (right line) virus (29, 30). On the basis of these transmission events, I-SH proponents divide the pd into three subphases: II-1, II-2, and II-3. They also conclude that egestion does not occur during pds because virus acquisition occurs before inoculation and egestion occurs after ingestion in aphid membrane feeding [15, 29, 30]. They associate waveform II-1 (Fig. 1) with Sw excretion and virus inoculation. Since the first Sw excreted through the common duct presumably would contain the highest concentration of virions, it should be possible to push forward the point of probing interruption to perhaps only about 0.1 second into subphase II-1 (Fig. 1). Furthermore, doing so ought to have little, if any, effect on the inoculation efficiency of viruliferous aphids compared with ones that are interrupted at the end of phase II-1 (Fig. 1). If pushing forward the interruption point does reduce aphid inoculation rates, then proponents of the I-SH must ask themselves if their hypothesis is built on bedrock or sand. Proponents of the I-SH have not yet associated subphase II-2 and the "archlets" portion of subphase II-3, together comprising over 50% of the pd, with any aphid activities (Fig. 1). Potential Drop Analysis According to the Ingestion-Egestion Theory
C. 1.
PD PHASES AND SUBPHASES
The pd in Figure 1 reflects a continuum of aphid activities associated with host selection. There are similarities and differences among the four subphases or waveforms comprised by the pd. For example, in frequency (Hz) and appearance, wave peaks at the end of waveform Kh-la resemble peaks at the start of Kh-lb more than they do ones at the start of Kh-la. In other words, waveforms Kh-la and Kh-lb apparently are "flip sides" of the pregustation phase of host selection (Fig. 1, Kh-1). Similarly, wave peaks at the end of waveform Kh-2a resemble peaks at the start of Kh-2b more than they do ones at the beginning of Kh-2a.
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However, wave peaks at the end of Kh-lb are strikingly different in appearance and frequency from those at the start of Kh-2a (Fig. 1). Thus, the pd appears to comprise two distinct host selection phases, pregustation (Kh-1) and gustation (Kh-2), and each of these phases is subdivided into subphases, Kh-1 a and Kh-lb, and Kh-2a and Kh-2b, respectively. Vertical, dash-dot-dash lines below the pd separate the different subphases or waveforms. Note that we have randomly pushed the left dash-dot-dash line slightly forward with respect to when inoculation supposedly occurs in the I-SH (the left dashed line above the pd). This indicates that in the I-ET, inoculation occurs at the start of the second pd subphase, Kh-lb, not at the end of the first (as in the I-SH). The four drawings at the bottom (Fig. 1) represent the aphid activity associated with each of the host selection subphases or waveforms. 2.
VIRUS INOCULATION
The first thing an aphid does after entering a cell is to salivate. Animals salivate as a pregustatory activity. (Think of Pavlov's experiments with dogs, admittedly on a higher scale than aphids but involving essentially the same principle.) An aphid first needs to wet and clean its food receptacle and gustatory sensilla of residue from previous sap sampling or feeding. Note that in the I-ET, inoculation requires ingestion-egestion of mainly Sw, not just egestion and not egestion of mainly cytoplasm. The aphid simultaneously secretes and ingests Sw (Fig. 1, drawing Kh-1 a); in addition to removing residue, this facilitates release of virions from their ACRs for subsequent inoculation by egestion (Kh-lb). In Kh-1 a, the wave of Sw sweeps virions from their ACRs as it advances up the feeding apparatus lumen toward the gustatory sensilla, swirling the virions into a Sw potpourri, irrespective of the virions' former ACR locations. Note that following cell puncture at the start of Kh-1 a, the potential continues to drop as Sw is drawn up the food canal. The latter reflects progressively increasing amperage and decreasing potential. The pd plateaus between Kh-1 a and Kh-lb when Sw contacts the dendritic plasmalemma of the gustatory sensilla in the vicinity of the cibarial valve. Having wetted and cleansed the sensilla with Sw, the aphid then egests the Sw, along with dissociated virions, into the cell (Fig. 1, Kh-lb) in preparation for the gustation phase, Kh-2, of host selection. In the I-ET, virion inoculation takes place at the start of pd subphase Kh-lb (Fig. 1). Since there are no degrees of inoculation and assuming a single-hit infection mechanism, most inoculations should and do occur at the start of Khlb. As one would expect, the rate of transmission by aphids interrupted at the end of Kh-lb is not significantly different from that by aphids interrupted at its beginning (or the end of subphase II-1 in the I-SH [29, 30]). Therefore, most aphids appear to acquire manyfold more virions than are needed for them to be inoculative. If this were not so, one would expect an increase in transmission by viruliferous aphids that are interrupted further into or at the end of subphase
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Kh-lb, since by egesting for a longer time they presumably also would egest larger volumes of inoculum. The repetition rate of waveform Kh-la (II-1 in the I-SH system) is 11-20 Hz [15]. This rate differs greatly from the 2- to 7-Hz waveform associated with Sw excretion [15, 45]. It is, however, very like the 8- to 25-Hz waveform associated with cibarial muscle activity and active ingestion [15]. Simultaneous Sw secretion and ingestion would involve contraction of the cupula muscles (afferent duct closed) of the salivary pump, relaxation of the piston retractor muscles of the salivary pump, and contraction of the cibarial valve and postcibarial retractor muscles [6, 7]. The transition from waveform Kh-la to Kh-lb is marked by a progressive increase in the amplitude of wave peaks and a progressive decrease in their frequency from 11-20 Hz to 3-9 Hz (Fig. 1) [15]. The latter reflects progressive decreases in muscle activity as the postcibarial retractors relax from their contraction levels at the end of Sw ingestion (Kh-la). Additionally, unlike waveform Kh-la, waveform Kh-lb does not involve the now relaxed cupula and piston retractors of the salivary pump [6, 7]. The increase in amplitude of Kh-lb wave peaks might reflect a decrease in the electrical connection between aphid and plant as Sw empties from the feeding-apparatus lumen into the cell (Fig. 1, drawing Kh-lb). 3.
VIRUS ACQUISITION
Egested Sw probably converts cytoplasm in the area of the cell puncture from gel to more easily ingestible sol. The 8- to 25-Hz frequency of waveform Kh-2a [15], like that of Kh-la, indicates cibarial muscle contraction, this time to ingest the cytoplasm/watery saliva (Cy/Sw) mixture to gustatory sensilla for tasting (Fig. 1, drawing Kh-2a). The aphid, having tasted, returns the Cy/Sw to the cell by egestion (drawing Kh-2b). A long-duration Kh-2 phase (subphase II-3 of the I-SH) is typical of early pds of efficient vectors and results in optimal transmission efficiency [29, 30]. Potential drops with a long Kh-2b waveform increase the likelihood of successfril (retentionwise) ACR-HC-CPS "hits" (or ACR-CPS hits in the case of cucumovirus transmission) because the Cy/Sw surface contacts ACRs coming (ingestion; Fig. 1, Kh-2a) and going (egestion, Kh-2b). The lower frequency of waveform Kh-2b reflects the decreasing involvement of the postcibarial retractors as they relax from their contraction levels at the end of Cy/Sw ingestion (Kh-2a). The higher amplitude of the wave peaks reflects weakening of the aphid-plant electrical connection as Cy/Sw empties from the food canal into the cell (Kh-2b). Note that previously egested virions (Fig. 1, drawing Kh-lb) can reenter the feeding apparatus lumen (drawing Kh-2a), possibly attaching again to ACRs. If so, aphids can be "recharged" by "used" or recycled virions. The potency of the recharge would progressively decrease during successive pds on the same or different healthy plants until the recharge reaches a noninfectious level. This recycling might explain, in part, why some viruliferous
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aphids, presumably those that acquire and subsequently recycle the most virions, are more successful than others in inoculating multiple plants in a succession of brief probes of inoculation test plants in a series. Ingestion-egestion of Sw accounts for virus inoculation (Fig. 1, Kh-la and KhIb). Ingestion-egestion of Cy/Sw, the gustation phase (Kh-2) of host selection, maximizes transmission efficiency. Hence, the I-ET involves even more ingesting and egesting than previously thought, not less or no egestion as claimed by proponents of the I-SH [15, 29-31]. Furthermore, the fluid ingested (ACR-HC-CPS dissociation) and egested (virion-HC or virion inoculation) during the pregustation phase of host selection (Kh-1) is mainly Sw, not plant cytoplasm.
W. Role of Watery Saliva in Transmission Anything relating to the role of Sw in transmission is of course still highly speculative. However, if the I-ET is correct (we would be willing to bet the farm, if we had one), Sw would seem to play an important role in releasing virions (Fig. 1, Kh-la) from ACRs for subsequent inoculation to plant host cells (Kh-lb). Before speculating about the molecular bases of how and where Sw effects ACR-HC-CP dissociation and hence virion release, we will briefly review what an analysis of current data tells us about the nature of potyvirus-aphid binding. Protein-protein binding is involved in virus retention. There are two major hypotheses regarding the mechanism of potyvirion binding, the direct binding hypothesis (DBH) and the bridge hypothesis (BH). In both hypotheses, a virusencoded proteolytic, helper protein (HC [or HC-Pro]) mediates the bridging process [46, 47]. The HC appears to function as a dimer (or possibly a trimer) with a molecular weight of 100-150 kDa [48,49]. According to the DBH, the HC attaches to CPS and effects a conformational change that allows direct binding of CPS to ACRs. However, an analysis of available data [41] favors the BH proposed originally in 1971 by Kassanis and Govier [50-52]. Blanc et al [42, 53, 54] recently presented a far more sophisticated molecular version of the BH. Exactly how HC forms a bridge connecting virion to ACR (ACR-HC-CPS) remains to be determined. However, a number of linear sequence motifs seemingly play important roles in the process. Changes (some as slight as a single amino acid substitution) in or sometimes around these motifs may transform a transmissible ACR-HC-CPS combination to a nontransmissible one. A DAG motif (aspartic acid-alanine-glycine) in the N-terminal region of the CPS appears to be the site at which HC binds to CPS [53]. Furthermore, a PTK motif (prolinethreonine-lysine) of HC is seemingly directly or indirectly involved in HC-DAG binding. Little is known about the ACRs; however, a KITC motif (lysineisoleucine-threonine-cysteine) of HC seems to function in binding HC to the putative ACR protein. Raccah et al [41] suggest two models of HC dimer bridg-
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ing. In the first, two HC molecules coassociate, with the first HC molecule attached on one side to an ACR and on the other to a second HC molecule which is bound to CPS (ACR-HC-HC-CPS). In the second model, a side-by-side HC dimer binds to a single ACR at one end and to DAG motifs of adjacent CPSs at the other (ACR-HC/HC-2DAGs). Based on the foregoing, we will now speculate about how Sw dissociates (Fig. 1, Kh-la) ACR-virion bridges built during the acquisition phase of a pd (Fig. 1, ICh-2a and Kh-2b). Secondary and tertiary molecular structure is known to function in animal virus host membrane recognition [55, 56]. Additionally, molecular conformation appears to enable direct binding of cucumovirus CP to ACRs (43). A highly conserved domain, the PH-pi loop, in the cucumovirus CPS appears involved in ACR-CPS binding (see chapter 9). If secondary or tertiary conformations or both are crucial to motif-motif recognition in ACR-HC-CPS bridge formation, then the mechanism by which Sw dissociates (or builds) ACR-HC-CPS bridges could be as simple as pH-mediated changes in protein electrostatic charge and conformation. Changes in pH can influence the structure, charge, and conformation of proteins and consequently can influence protein affinities and protein-protein interactions. Many amino acids that are chargeless at neutral to slightly basic pHs (7 to 8) become positively charged in a slightly acidic milieu (e.g., pH of 6 to 7). N-Terminal amines can be similarly changed from neutral to positive. Animal saliva is generally basic. Whether this is true of homopteran Sw remains to be determined. Also, the effects of Sw on the pH and composition of Cy/Sw (Fig. 1, Kh-2a) is not known. However, virions and HC in Cy/Sw ingested during subphase Kh-2a are able to bridge to ACRs. Hence, the design and execution of experiments to further elucidate the chemistry of aphid Sw appear to be worthwhile pursuits. One could also design experiments to determine which bridge linkage(s) Sw cleaves. (However, at this point, one cannot rule out the possibility of direct involvement or coinvolvement of known or unknown constituents of Sw that compete for or induce changes in bridge-binding sites.) We do know from membrane-feeding experiments followed by immunogold-TEM that HC by itself can bind with ACRs. Therefore, by combining these protocols with the EPG technique, one could interrupt viruliferous aphids probing a healthy test plant about two-thirds or so into waveform Kh-lb (or at least before the start of Kh-2a). If subsequent immunogold-TEM revealed little or no HC in the feeding apparatus, one could conclude (assuming appropriate controls) that bridge dissociation occurs at the ACR-HC link. On the other hand, the presence of HC would indicate a break at either the HC-HC link (perhaps the bridge's weakest) or the HC-CPS link. Cleavage of the HC-HC link would release HC as well as virions into the inoculated cell during subphase Kh-lb (Fig. 1). Then HC could influence events in planta, such as virion movement or other processes leading to infection site development and spread. It would also be fruitful to combine membrane feeding with the EPG technique followed by bioassay and immunogold-TEM or autoradiography. This combina-
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tion of technologies would permit researchers to vary the feeding solution (e.g., feeding solution only, functional versus nonfunctional HC, functional and nonfunctional ACR-HC-CPS combinations, feeding solution pH, etc.) in combination with different probe interruption points relative to host selection or virus transmission events. Relating pH changes in such experimental settings to changes in HC or HC-virion acquisition and inoculation might help elucidate the role of Sw in the transmission process.
VIL Semipersistent Transmission Proponents of the I-SH accept egestion as the mechanism of semipersistent transmission [15, 29-31]. (This concession probably relates more to an inability to conceive of any other way to get virions from the feeding apparatus lumen, especially the cibarium, to the plant than to a genuine acceptance of egestion and its role in homopteran probing and feeding and consequently cuticula-bome virus transmission.) However, the milieu for semipersistent inoculation probably is different from what is advocated by the I-SH proponents Ferreres and Collar [30]: "During salivation, it is unlikely that MCDV [semipersistent, leafhopper-borne, maize chlorotic dwarf virus] will be injected into the plant because most of the MCDV-like particles found attached to the foregut of G. nigrifrons [leafhoppers] were in the precibarium, cibarium and pharynx through which ejected saliva does not pass." With some semipersistent virus-vector combinations, vectors might require hours of feeding to saturate retention sites and maximize transmission. Virus retention can be heavily biased toward the postcibarium [22, 37]. The postcibarium represents an enormous surface area for ACRs and possible virus retention, as compared with mainly the maxillary food canal and antecibarium for nonpersistent virus transmission. (Hopefully, no one will jump the gun and begin calling semipersistent viruses "cibarium-borne." Or did we just do so?) Greater virus-carrying capacity explains the longer acquisition and retention periods of semipersistent transmission. Aphids simultaneously secrete Sw and ingest sap while feeding [15], creating a milieu, Cy/Sw, seemingly conducive to bridge formation and virus acquisition. Conversely, ingestion-egestion of Sw, which dissociates ACR-HC-CPS bridges, seems a better candidate for semipersistent virus inoculation than ingestion-egestion of Cy/Sw. Aphids control the flow rate of sap to postpone and minimize a wound healing response on the part of the plant and hence avoid the need for frequent feeding site changes. Nevertheless, given sufficient feeding time, presumably all aphids eventually face the problem of p-protein fibrils and callose clogging the entrance to the maxillary food canal, just as these substances clog pores in the sieve plates of ruptured sieve cells. Additionally, feeding in a virus-infected plant will coat the cuticular lining of the feeding apparatus with ACR-HC-virion complex. The latter
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Table I Comparison of the Ingestion-Egestion Theory and Ingestion-SaHvation Hypothesis of Nonpersistent Cuticula-Bome Virus Transmission. Ingestion-Egestion Theory
Comparisons General Amount of potential drop (pd) explained Relating pd waveforms to aphid activities
Virus inoculation Virus retention site(s)
Inoculation mechanism 11 to 20-Hz, first pd waveform [15, 30] How does salivation on glass reduce aphid inoculativity?
Why does salivation by suspended aphids not affect aphid inoculativity [39]? Virus Acquisition Acquisition mechanism How do some interrupted aphids acquire virus in the first pd-subphase [29, 30]?
Ingestion-SaHvation Hypothesis
All
Less than half
Based on host selection and waveform attributes
Based on the sequences of events during virus transmission and membrane feeding
Aphid cuticular receptors (ACRs) in feeding apparatus lumen Ingestion-egestion of watery saliva (Sw) Mimics cibarial muscle activity (8-25 Hz [15]) Aphid secretion/ingestion and egestion of Sw dissociates and flushes virions (Fig. 1, Kh-1) Virions are out of harm's way
ACRs in conmion (food-saliva) duct
Ingestion-egestion of cytoplasm/Sw (Cy/Sw) Some virions enter the food canal during Sw secretion/ingestion (Fig. 1, Kh-1 a)
Ingestion of Cy/Sw
Sw excretion through common duct ? (Sw-excretion waveform is 2-7 Hz [15]) Saliva flushes virions from common duct
? (Salivation should flush virions from the common duct and reduce rate of inoculation)
? (Virions must go against Sw flow, II-1, and survive both post- and preprobing salivation)
greatly increases the surface area of the Hning, increasing its coefficient of drag and slowing the flow of sap. Such ingestion-inhibiting forces would likely induce simultaneous secretion and ingestion of Sw to the postcibarium. Ingestion-egestion of Sw disrupts ACR-HC-CPS bridges and cleans the cibarial lumen and food canal of clogs. Thus, secretion and ingestion-egestion of Sw effects virion dissociation and inoculation in both nonpersistent and semipersistent transmission. However, there are noteworthy differences between these two types of cuticulabome virus transmission. Given the virion retention systems in play and the small size of the inocula at work, it is essential that inoculation precede acquisition in the hurried world of nonpersistent transmission. If it did not, virus acquisition
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probably would decrease to a level incapable of ensuring the survival of a virus. In the more slowly paced workings of semipersistent transmission, involving much larger retention areas and virus-carrying capacity, however, inoculation follows acquisition. These transmission types also differ in the purpose of Sw secretion and ingestion-egestion, pregustation in nonpersistent transmission versus "declogging" in semipersistent.
VIIL Concluding Remarks Table I compares the I-ET with the I-SH. The I-SH challenged us to update the I-ET to explain, more definitively, the mechanisms of cuticula-bome virus transmission. We hope that we have adequately answered that challenge. The feeding apparatus lumen beyond the common duct is the site of noncirculative virus retention. Furthermore, an analysis of available data on noncirculative transmission by aphids, leafhoppers, and whiteflies, including EPG recording data, indicates that secretion and ingestion-egestion of Sw makes a better candidate for inoculation than does saliva excretion alone. Hopefully, researchers will continue to push the envelope in testing and strengthening the ingestion-egestion theory. And should the theory endure another 27 years of scrutiny, we hope to be here to update it once again!
References 1. Harris, K.F., Pesic-Van Esbroeck, Z., and Dufifiis, XE. (1994). A morphological study of Bemisia organ systems of known importance in homopteran virus transmission. Phytoparasitica 22, 323-324. 2. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, IE. (1995). Vector anatomy of the sweet potato whitefly. In "Advances in Vegetable Virus Research." Proceedings of the 8th International Conference on Virus Diseases of Vegetables, July 9-15, Prague, pp. 49-52. 3. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Morphological and cellular bases of virus transmission by whiteflies. Proceedings of the 20th International Congress on Entomology, Florence, Italy, Aug. 25-31, p. 453. 4. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Morphological bases for whitefly transmission of viruses. In "Silverleaf Whitefly 1996 Supplement of the Five-Year National Research and Action Plan. USDA-ARS Publ. No. 1996-01, 34. 5. Harris, K.F. (1990). Aphid transmission of plant viruses. In "Plant Viruses" (C.L. Mandahar, ed.). Vol. 2, pp. \11-10A. CRC Press, Boca Raton, FL. 6. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Morphology of the sweet potato whitefly, Bemisia tabaci (Homoptera: Aleyrodidae) relative to virus transmission. Zoomorphology 116, U3-\56. 7. Harris, K.F, Pesic-Van Esbroeck, Z., and Duflus, J.E. (1996). Anatomy of a virus vector. In ''Bemisia: 1995 Taxonomy, Biology, Damage, Control, and Management" (D. Gerling and R.T. Mayers, eds.), pp. 289-318. Intercept Limited, Andover, U.K.
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8. Berger, RH., Zeyen, RJ., and Groth, J.V (1987). Aphid retention of maize dwarf mosaic virus (potyvirus): Epidemiological implications. Ann. Appl Biol. I l l , 337-344. 9. Zeyen, R.J., and Berger, RH. (1990). Is the concept of short retention times for aphid-borne nonpersistent viruses sound? Phytopathology 80, 769-771. 10. Fereres, A., Blua, M., and Perring, T. (1992). Retention and transmission characteristics of zucchini yellow mosaic virus by Aphis gossypii and Myzus persicae (Homoptera: Aphididae). J. Econ. Entomol. 85, 759-765. 11. Watson, M.A. (1938). Further studies on the relationship between Hyosciamus virus 3 and the aphid Myzus persicae (Sulz.) with special reference to the effects of fasting. Proc. R. Soc. Lond. Biol. 125, 144-170. 12. Ammar, E.D., Jarlfors, U, and Pirone, T.R (1994). Association of potyvirus helper component protein with virions and the cuticle lining the maxillary food canal and foregut of an aphid vector. Phytopathology %A, 1054-1060. 13. Harris, K.F., and Bath, J.E. (1973). Regurgitation by Myzus persicae during membrane feeding: Its likely function in transmission of plant viruses by vectors. Ann. Entomol. Soc. Am. 66,793-796. 14. Harris, K.F. (1977). An ingestion-egestion hypothesis of noncirculative virus transmission. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 165-220. Academic Press, New York. 15. Tjallingii, W.F., and Prado, E. (2001). Analysis of circulative transmission by electrical penetration graphs. In "Virus-Insect-Plant Interactions" (K.F. Harris, O.R Smith and J.E. Duffus, eds.), chapter 4 (this book). Academic Press, New York. 16. Nauh, L.R., and Ammar, E.D. (1989). Leafhopper and planthopper transmission of plant viruses. Annu. Rev. Entomol. 34, 503-529. 17. Kennedy, J.S., Day, M.F, and Eastop, VF (1962). "A Conspectus of Aphids as Vectors of Plant Viruses." Commonwealth Institute of Entomology, London. 18. McLean, D.L., and Kinsey, M.G. (1984). The precibarial valve and its role in the feeding behavior of the pea aphid, Acyrthosiphon pisum. Bull. Entomol. Soc. Am. 30, 26-31. 19. Davidson, J. (1914). On the mouth-parts and mechanism of suction in Schizoneura lanigera Hausmann. J. Linnean Soc. Lond. Zoology 32, 307-330. 20. Weber, H. (1928). Skelett, Muskulatur und Darm der Schwarzen Blattlaus Aphis fabae Scop. Mit besonderer Beriicksichtigung der Funktion der Mundwerkzeuge und des Darms. Zoologica 28, 1-120. 21. Harris, K.F., Treur, B., Tsai, J., and Toler, R. (1981). Observations on leafhopper ingestion-egestion behavior: Its likely role in the transmission of noncirculative viruses and other pathogens. J. Econ. Entomol. 74, 446-453. 22. Ammar, E.D., and Nault, L.R. (1991). Maize chlorotic dwarf virus-like particles associated with the foregut in vector and non-vector leafhopper species. Phytopathology 81,444-448. 23. Doolittle, S.R, and Walker, M.N. (1928). Aphis transmission of cucumber mosaic. Phytopathology 18, 143. 24. Bradley, R.H.E. and Ganong, R.Y (1955). Some effects of formaldehyde on potato virus Y in vitro, and ability of aphids to transmit the virus when their stylets are treated with formaldehyde. Can. J. Microbiol. 1, 783-793. 25. Bradley, R.H.E., and Ganong, R.Y. (1955). Evidence that potato virus Y is carried near the tips of the stylets of the di^Yiidi Myzus persicae (Sulz.). Can. J. Microbiol. 1,115-1S2. 26. Lopez-Abella, D. and Bradley, R.H.E. (1969). Aphids may not acquire and transmit stylet-borne viruses while probing intercellularly. Virology 39, 338-342. 27. Lopez-Abella, D., Bradley, R.H.E., and Harris, K.F. (1988). Correlation between stylet paths made during superficial probing and the ability of aphids to transmit nonpersistent viruses. In "Advances in Disease Vector Research" (K.F. Harris, ed.). Vol. 5, pp. 251-285. Springer-Verlag, New York.
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28. Forbes, A.R. (1969). The stylets of the green peach aphid, Myzus persicae (Homoptera: Aphididae). Can. Entomol 101, 31-41. 29. Martin, B., Collar, J.L., Tjallingii, W.F., and Fereres, A. (1997). Intracellular ingestion and salivation by aphids may cause acquisition and inoculation of nonpersistent transmitted plant viruses. J. Gen. Virol. 78, 2701-2705. 30. Fereres, A., and Collar, XL. (2001). Analysis of noncirculative transmission by electrical penetration graphs. In "Virus-Insect-Plant Interactions" (K.F. Harris, O.P. Smith and J.E. Dufifus, eds.), chapter 5 (this book). Academic Press, New York. 31. Collar, XL., Avilla, C , and Fereres, A. (1997). New correlations between aphid stylet paths and nonpersistent virus transmission. Environ. Entomol. 26, 537-544. 32. Forbes, A.R. (1969). The morphology and fine structure of the gut of the green peach aphid, Myzus persicae (Sulzer) (Homoptera: Aphididae). Mem. Entomol. Soc. Can. 36, 1-74. 33. Forbes, A.R. (1977). The mouthparts and feeding mechanism of aphids. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 83-103, Academic Press, New York. 34. Taylor, C.E., and Robertson, VM. (1974). Electron microscopy evidence for the association of tobacco severe etch virus with the maxillae in Myzus persicae (Sulz.). J. Phytopathol. 80,257-266. 35. Berger, PH., and Pirone, T.P (1986). The effect of helper component on the uptake and localization of potyviruses in Myzus persicae. Virology 153, 256-261. 36. Ammar, E.D., Gingery, R.E., and Nault, L,R. (1987). Interactions between maize mosaic and maize stripe viruses in their insect vector, Peregrinus maidis, and in maize. Phytopathology 11, 1051-1056. 37. Childress, S.A., and Harris, K.F. (1989). Localization of virus-like particles in the foreguts of viruliferous Graminella nigrifrons leafhoppers carrying the semipersistent maize chlorotic dwarf virus. J. Gen. Virol. 70, 247-251. 38. Wang, R.Y, Ammar, E.D., Thornbury, D.W., Lopez-Moya, XX, Pirone, T.P (1996). Loss of potyvirus transmissibility and helper-component activity correlate with non-retention of virions in aphid stylets. J. Gen. Virol. 11, 861-867. 39. Bradley, R.H.E. (1959). Loss of virus from the stylets of aphids. Virology 8, 308-318. 40. Hashiba, T, and Misawa, T. (1969). Studies on the mechanism of aphid transmission of styletborne virus. (Ill) On the adherence of the virus to the stylet. TohokuJ.Agr Res. 20, 159-171. 41. Raccah, B., Huet, H. and Blanc, S. (2001). Potyviruses. In "Virus-Insect-Plant Interactions" (K.F Harris, O.P. Smith, and XE. Duffus, eds.), chapter 10 (this book). Academic Press, New York. 42. Blanc, S., Hebrard, E., Drucker, M., and Froissart, R. (2001). Caulimoviruses. In "Virus-InsectPlant Interactions" (K.F. Harris, O.P. Smith, and XE. Duffus, eds.), chapter 8 (this book). Academic Press, New York. 43. Perry, K.L. (2001). Cucumoviruses. In "Virus-Insect-Plant Interactions" (K.F. Harris, O.P. Smith and XE. Dufifus, eds.), chapter 9 (this book). Academic Press, New York. 44. Collar, XL., and Fereres, A. (1998). Nonpersistent virus transmission efficiency determined by aphid probing behavior during intracellular punctures. Environ. Entomol. 27, 583-591. 45. Prado, E., and Tjallingii, W.F. (1994). Aphid activities during sieve element punctures. Entomol. Exp.Appl. 72,157-165. 46. Maia, I.G., and Bemardi, F. (1996). Nucleic acid binding properties of a bacterially expressed potato virus Y helper component-proteinase. J. Gen. Virol. 11, 869-877. 47. Carrington, XC, Cary, S.M., Parks, T.D., and Dougherty, W.G. (1989). A second proteinase encoded by a plant potyvirus genome. EMBO J. 8, 365-370. 48. Wang, R.Y., and Pirone, T.P. (1999). Purification and characterization of turnip mosaic virus helper component protein. Phytopathology 89, 564-567. 49. Thornbury, D.W., Hellmann, G.M., Rhoads, R.E., and Pirone, T.P (1985). Purification and characterization of potyvirus helper component. Virology 144,260-267.
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50. Kassanis B., and Govier, D.A. (1971). New evidence on the mechanism of aphid transmission of potato C and potato aucuba mosaic viruses. J. Gen. Virol. 10, 99-101. 51. Kassanis B., and Govier, D.A. (1971). The role of the helper virus in aphid transmission of potato aucuba mosaic virus and potato C. J. Gen. Virol. 13, 221-228. 52. Govier, D.A., and Kassanis, B. (1974). Evidence that a component other than the virus particle is needed for aphid transmission of potato virus Y. Virology 57,285-286. 53. Blanc, S., Lopez-Moya, J.J., Wang, R., Garcia-Lampasona, S., Thombury, D.W., and Pirone, T.P. (1997). A specific interaction between coat-protein and helper component correlates with aphid transmission of a potyvirus. Virology, 231, 141-147. 54. Blanc, S., Ammar, R.D., Garcia-Lampasona, S., Dolja, VV Llave, C , Baker, J., and Pirone, T.P. (1998). Mutations in the potyvirus helper component protein: Effects on interactions with virions and aphid stylets. J. Gen. Virol. 79, 3119-3122. 55. Rossmann, M.G., Arnold, E., Erickson, J.W., Frankenberger, E.A., GriflFith, J.P, Hecht, H.L., Johnson, J.E., and Kamer, G. (1985). Structure of a human common cold virus and functional relationship to other picomaviruses. Nature 317, 145-153. 56. Hogle, J.M., Chow, M., and Filmann, D.J. (1985). The three dimensional structure of poliovirus at 2.9 A resolution. Science 229, 1358-1365.
CHAPTER 7
Mechanism of Virus Transmission by Leaf-Feeding Beetles ROSE C. GERGERICH
/- Introduction The most widespread and important species of beetle vectors are in the family Chrysomelidae (leaf beetles), although vectors are also found in the families Coccinellidae (ladybird beetles), Curculionidae (weevils), and Meloidae (blister beetles). Adult beetles transmit plant viruses under field conditions. The larval stages of several chrysomelid beetles have also been reported as virus vectors [1-3], some being more efficient than their respective adults [1,2]. There are no reports of virus being acquired by larvae and carried through pupation to the adult. Beetles are vectors of viruses in the genera Sobemovirus, Comovirus, Bromovirus, Tymovirus, Machlomovirus, and Carmovirus. All beetle-transmitted viruses are relatively stable, single-stranded RNA viruses with isometric, 25- to 30-nm particles, which reach high concentrations in their plant hosts. Beetles play an important role in the ecology and epidemiology of many of these viruses in the field, but seed transmission and mechanical transmission by cultural practices are important factors with some of them. The transmission of viruses by beetles is characterized by short acquisition access and inoculation feeding periods, the absence of a latent period, and short persistence times in actively feeding beetles.
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Virus Acquisition: Beetie-Piant interactions
Beetles acquire virus quickly. Studies have demonstrated that they may become viruliferous after a single bite [4], although extended feeding times result in Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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increased transmission efficiency [5-7]. Beetles can acquire vims by drinking a solution of purified virus mixed with sucrose [8]. From the latter we conclude that the virus particle alone is sufficient for virus transmission and therefore that virus transmission is not dependent on a viral-encoded helper component, as is the case v/ith aphid transmission of Potyviruses and Caulimoviruses. There is no evidence that virus multiplication occurs in the beetle; for beetle-transmissible bean pod mottle virus in the bean leaf beetle, it has been shown that virus concentration in the beetle gradually declines after acquisition [9]. Plant-eating beetles lay their eggs on the leaves or in the soil near roots of their host plants. Emerging beetle larvae are not very mobile and often feed on the same plants that their parent fed on prior to egg deposition. If the parent beetle is viruliferous, there is an enhanced probability that emerging adults will begin feeding on a virus-source plant that was infected by the viruliferous parent beetle prior to egg laying. Other than this possibility, however, there is no evidence that adult beetles select and feed preferentially on virus-infected plants. Acquisition and transmission of viruses by leaf-feeding beetles are limited not only by the susceptibility of the host plants to the virus but also by the host range of the beetle vector. Most plant-feeding beetles will feed only on a limited series of host plants. Host selection by the beetle is determined by the presence of attractant and inhibitory substances in the plants [13a], which are detected by chemoreceptors on antennae and maxillary palpi [10]. The striped cucumber beetle, Acalymma vittata, can be induced to feed on bean plants by treating the leaves of the bean plant with an alcohol extract from butternut squash, a preferred host of this beetle. The beetle can also transmit southern bean mosaic virus (SBMV) from and to extract-treated bean [11]. The inability of the striped cucumber beetle to transmit SBMV in a bean host under normal conditions is therefore due to the feeding preferences of the beetle vector, not to the inability of this beetle to deliver the virus in a manner that might result in transmission.
///. Plant Virus-Beetle Interactions Many beetle-transmitted viruses, as well as some not transmitted by beetles, have been found in the regurgitant [12] and hemolymph [4, 13, 14] of beetles fed on virus-infected plants. This suggests that beetles have a circulative-like relationship with the viruses they transmit. However, some efficiently transmitted viruses are not found in the hemolymph of their beetle vectors but are present in regurgitant [14]. For example, bean pod mottle virus (BPMV) is not found in the hemolymph of viruliferous Mexican bean beetles, bean leaf beetles, or spotted cucumber beetles [14]. At least for these virus-beetle combinations, the relationship appears more like cuticula-bome [15, 16] or semipersistent type of vector transmission.
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The interaction of plant viruses with their beetle vectors appears to be governed by both the species of beetle and the virus. Although SBMV does not move from the gut to the hemocoel of the Mexican bean beetle, it does occur in the hemocoel of viruliferous bean leaf beetles and spotted cucumber beetles [14], which suggests that the type of beetle affects virus mobility within the insect. On the other hand, viral properties of the virus particle other than size appear to regulate movement into the hemocoel. Specifically, BPMy similar in size to SBMy moves into the hemocoel of the Mexican bean beetle, whereas SBMV does not [14]. An in-depth study of the interaction of SBMV with the spotted cucumber beetle, Diabrotica undecimpunctata howardi, revealed that virus enters the hemocoel of the beetle through the peritrophic membrane-lined midgut but not through the intima-lined foregut or hindgut [17]. Virus retention time in vectors can be an important factor in patterns of dissemination and overwintering by viruliferous vectors. On the basis of reports of virus retention in overwintering beetles [4] and the presence of virus in the hemolymph of beetle vectors [4, 12, 13, 18], it was assumed that ingested virus is retained in the hemocoel during long retention times [19,20]. Later studies, however, demonstrated that some plant viruses efficiently transmitted by beetles and retained in overwintering beetles, such as BPMV in the bean leaf beetle [21, 22], are not circulative in their vectors [14]. In laboratory studies, the retention time for SBMV in the Mexican bean beetle was found to be similar to that in two other beetle species, in spite of the fact that this virus is not found in the hemolymph of the Mexican beetle [23]. The capsid proteins of the beetle-transmitted comoviruses are partially degraded by proteases in beetle regurgitant, which changes the overall charge of the virus particles [8]. There is no evidence, however, to suggest that this partial digestion is necessary for comovirus transmission.
IV. Deposition of Virus in Beetie Regurgitant Several lines of evidence suggest that beetles regurgitate during feeding. When Mexican bean beetles or bean leaf beetles ingest bean leaves that have been infiltrated with the dye bromophenol blue, the beetles deposit blue regurgitant when they attempt to feed on bean leaves or wet filter paper disks. Additional evidence for beetle regurgitation comes from examination of beetle-feeding wounds using virus-specific immunofluorescent labeling. "Puddles" of deposited virus may be seen in the anticlinal grooves on the leaf surface at the edges of beetle feeding wounds [24]. Experimental analysis of regurgitant and its role in virus transmission is made possible by the fact that beetles can be induced to regurgitate by various stimuli [25], although the argument can be made that induced regurgitant may not be the same as naturally produced regurgitant.
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Regurgitant appears to be derived from maxillary and mandibular gland secretions and contents of the beetle foregut. Leaf-feeding beetles lack salivary glands, but the larvae and adults have gnathal glands associated with the maxillae and mandibles. In many Coleoptera, these glands open to the exterior on the basal inner surface of each maxilla and mandible [26]. Gnathal gland secretions are thought to constitute part of beetle regurgitant. The maxillary and mandibular glands of the Mexican bean beetle and the spotted cucumber beetle contain a central unbranched duct surrounded by a single layer of glandular epithelial cells [27]. This arrangement suggests a secretory ftinction for gnathal glands, but their actual function remains unknown. It appears that virus from beetle hemolymph is not transported into the gnathal glands and from there into regurgitant [27]. Evidence that the contents of the foregut are a component of regurgitant comes from studies of viruses that are not circulative in their beetle vectors. The legume isolates of tobacco mosaic virus (TMV) and SBMV are not found in the hemolymph of viruliferous Mexican bean beetles [14]; nevertheless, these viruses are present in regurgitant of viruliferous beetles and at feeding wound sites on bean leaves [24]. These results suggest that some of the components of regurgitant are derived from the foregut. Beetle regurgitant is a complex mixture of enzymes, which contains high levels of ribonuclease [28] and proteases [8], but plant viruses are not inactivated by these enzymes in virus regurgitant mixtures [29]. Initially, it was thought that beetle regurgitant contains an inhibitor, which selectively stops infection of all but the beetle-transmissible viruses. However, analysis with standard local lesion tests using mechanical rub inoculation of virus: regurgitant mixtures shows that infectivity of both beetle-transmissible and non-beetle-transmissible viruses is equally inhibited by regurgitant [25]. Unlike other virus-vector relationships, in which attachment or circulation in the vector determines plant virus retention and transmissibility [30, 31], many stable plant viruses that are not transmitted by beetles are retained within the beetle after acquisition and are infectious when deposited in regurgitant during feeding [12]. This led to the theory that the key to specificity of virus transmission by beetles is virus-plant interactions after virus is deposited in beetle regurgitant. Theoretically, the beetle's role is to acquire and retain virus for delivery via regurgitant to a receptive plant.
IV A.
Virus-Host Plant Interactions
The Gross-Wound Inoculation Technique
A pivotal advance in the study of virus transmission by beetles was the demonstration that the type of feeding damage inflicted by beetles is critical to whether
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virus deposited in regurgitant is able to infect a plant [25]. An inoculation method called the gross wounding technique was developed to mimic plant damage from beetle feeding and at the same time to apply virus-regurgitant mixtures to inoculation wounds. Gross-wound inoculation is accomplished by puncturing a leaf with the jagged edge of a glass cylinder that has been dipped in virus inoculum. Unlike the results with mechanical rub inoculation of virus-regurgitant mixtures, infectivity of beetle-transmissible and non-beetle-transmissible viruses was equally inhibited by regurgitant. Gross-wound inoculation of virus-regurgitant mixtures results in transmission of beetle-transmissible viruses only [25]. The action of regurgitant against non-beetle-transmissible viruses occurs with regurgitant from several different beetles and is effective in different hosts. The latter suggests that the effect is universal [25, 29]. Also, since non-beetle-transmissible viruses are transmitted by gross-wound inoculation in the absence of regurgitant in the inoculum, it appears that the combined effect of regurgitant and the inoculation procedure is necessary for selective transmission of viruses by beetles. B.
Role of Ribonuclease
Analysis of beetle regurgitant to determine what constituent(s) might be responsible for the selective inhibition demonstrated by gross-wound inoculation revealed that the component was a heat-labile macromolecule, possibly an enzyme [25]. Analysis of the enzymes in the regurgitant of several leaf-feeding beetles revealed high levels of ribonuclease activity. Incorporation of commercially available preparations of bovine pancreatic ribonuclease at activities similar to those found in beetle regurgitant into gross-wound inocula results in selective inhibition of non-beetle-transmissible viruses [28]. Regurgitant is a complex mixture of proteins and enzymes, some of which may interact with plant viruses [8]. Nevertheless, it appears that ribonuclease in regurgitant is the component preventing the transmission of all but beetle-transmissible viruses. Several lines of evidence indicate that the mechanism by which ribonuclease selectively prevents transmission of non-beetle-transmissible viruses is not due to selective destruction of these viruses. Non-beetle-transmissible viruses such as the legume isolate of TMV and tobacco ringspot virus (TRSV) are infectious when they are deposited in beetle regurgitant on the leaf surface [12]. Furthermore, non-beetle-transmissible viruses are infectious when purified from virusregurgitant mixtures [29] or when virus-regurgitant mixtures are diluted sufficiently to reduce the inhibition of ribonuclease in rub-inoculation infectivity assays [25]. The exception to this may be the aphid-transmitted cucumber mosaic virus, which is inactivated by ribonuclease in vitro [32]. Some beetle-transmissible viruses, such as cowpea chlorotic mottle virus and brome mosaic virus, swell at pH levels higher than 6.0 [33]. The pH of Mexican bean beetle regurgitant is about 5.5, suggesting that the RNA of these viruses would not be exposed to the RNase in beetle regurgitant [28].
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It is proposed that the specificity of virus transmission by leaf-feeding beetles is caused by the presence of RNase on the feeding wound edge, which prevents infection of plant cells wounded during gross-wound inoculation or beetle feeding. RNase is known to be a powerful inhibitor of virus infection, and undiluted regurgitant inhibits infection of both beetle-transmissible and non-beetle-transmissible viruses when inoculated by mechanical rub inoculation [25]. Beetletransmissible viruses appear to escape the effects of RNase at the wound edge by translocation in the vascular system (away from the RNase), which is followed by infection of unwounded cells. C.
Translocation of Beetle-Transmissible Viruses
Several lines of evidence suggest that the virus particles of beetle-transmissible viruses are more mobile in the vascular system than those of non-beetle-transmissible ones. Perhaps the best visual evidence for this phenomenon as it relates to virus transmission comes from virus-specific, immunofluorescent, localization studies of whole leaf pieces after virus has been deposited by viruliferous leaf-feeding beetles during feeding [24]. Beetle-transmissible viruses such as BPMV, SBMV, and blackgram mottle virus are present in veins radiating from the wound site, sometimes several millimeters from the wound edge. Non-beetle-transmissible viruses such as TRSV and TMV are only occasionally seen in short segments of veins leading from the wound edge. The importance of an inoculation technique that mimics beetle feeding by damaging the vascular system might be that it allows virus entry into the xylem where it can be translocated away from the wound edge. Additional evidence for translocation of beetle-transmissible viruses in the vascular system comes from studies in which young cowpea and bean seedlings that are severed at their base and placed in purified virus suspensions to determine which virus can translocate in the xylem to the upper portions of the plant [34]. The three beetle-transmissible viruses used in this study translocated into the growing point of the seedlings, whereas none of the non-beetle-transmissible viruses translocated under the same conditions. Immobilization of non-beetletransmissible viruses at the feeding wound edge is considered a key factor in preventing transmission of these viruses. An alternative explanation for the lack of translocation of non-beetle-transmissible viruses is that they interact with and are destroyed by cellular components. When tobacco rattle virus or TMV is infiltrated into plant leaves, the virus attaches in a specific end-on orientation to plant cell walls [35]. The attached virus particles become shorter with time and disappear completely within 5 days postinfiltration. This suggests that the virus particles are not only immobilized on the cell wall but also degraded. Immunofluorescent localization studies of virus in beetle feeding wounds [24] also indicate that, unlike beetle-transmissible viruses such as SBMV and BPMV, non-beetle-transmissible viruses such as TRSV and TMV are degraded in leaf veins within 2 to 3 days after beetle feeding.
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It is not clear what role, if any, degradation of immobilized viruses plays in preventing virus infection following virus deposition by leaf-feeding beetles. The plant and viral determinants that result in virus immobilization in the plant are unknown at this time. Evidence from heterologously assembled virus particles suggests that the viral capsid protein is one determinant of transmission. Heterologous virus particles, having the capsid of a beetle-transmissible virus (SBMV) and the RNA of a non-beetle-transmissible one (the legume isolate of TMV) are transmitted by beetles and produce TMV-infected plants [36]. It seems, therefore, that the virus coat protein mediates virus-beetle or virus-plant recognition and compatibility needed for beetle transmissibility. D.
Infection of Unwounded Plant Cells
Following translocation in the vascular system and escape from regurgitant RNase, beetle-transmissible viruses must infect unwounded plant cells if they are going to establish an infection in the plant. Several lines of evidence suggest that beetletransmissible viruses are capable of escaping the vascular system and infecting unwounded cells. By using an experimental system developed by Schneider and Worley [37], it was shown that beetle-transmissible viruses, but not non-beetletransmissible viruses, can move through the xylem of steam-killed sections of bean stems and establish infection in unwounded cells in the plant above the killed portion of the stem [34]. When seedlings of pinto bean, a local lesion host for SBMV, are cut at the base of the stem and placed in a solution of purified SBMy the virus is translocated into the growing point of the seedling, where it establishes an infection characterized by the appearance of local lesions in the developing leaves. More direct evidence for infection of unwounded cells comes from immunofluorescent localization studies [24] of virus infection sites following beetle transmission of SBMV and BPMV Five days after virus transmission, clusters of fluorescing cells representing sites of virus infection appear adjacent to veins leading from the feeding wound site, but not on the cut edge of the beetle feeding wound. This study provides direct, visual evidence for infection of unwounded cells by beetle-transmissible viruses following virus transmission by beetles. Infection of unwounded cells by beetle-transmissible viruses does not appear to be characteristic of all beetle-transmissible viruses under normal conditions of systemic spread of virus in a plant. When stem segments of bean plants were steamed, followed by mechanical inoculation of virus on carborundum-dusted leaves below the killed portion of the stem, only SBMV was able to infect the top portions of the plant. Other beetle-transmissible viruses such as BPMV and blackgram mottle virus remained restricted to the portion of the plant below the steam-killed segment of stem. Opalka et al [38] proposed a mechanism for rice yellow mottle sobemovirus (RYMV) transport through pit membranes of adjacent xylem cells during xylem vessel maturation. Their model describes a mechanism for translocation and accumulation of RYMV and other sobemoviruses such as
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SBMV in young xylem vessels during systemic spread of the virus. It does not, however, address the question of how sobemoviruses escape xylem vessels and infect adjacent, unwounded, living parenchyma cells. Infection after escape of the virus from xylem vessels following beetle feeding or gross-wound inoculation appears to be less efficient than inoculation by mechanical rubbing of carborundum-dusted leaves. Some plants that are susceptible to mechanical inoculation with infectious plant sap are not susceptible when inoculated by beetles or gross-wound inoculation [39]. This has implications in programs that screen for virus resistance using mechanical inoculation; plants susceptible to mechanical inoculation can be resistant to virus transmission by beetles. Hobbs and Kuhn [40] observed that cowpea cultivars differ in their susceptibility to the cowpea strain of SBMV in the field, even though these cultivars were equally susceptible to mechanical inoculation with undiluted infectious sap. Gross-wound inoculation of mixtures of infectious sap and regurgitant revealed a pattern of susceptibility among the cultivars similar to that seen under field conditions [39], suggesting that gross-wound inoculation is a more accurate method for detection of resistance to beetle-transmitted viruses.
VL Concluding Remarks Virus transmission by leaf-feeding beetles is a complex process in which virus intimately associated with a beetle vector is deposited in regurgitant in beetle feeding wounds and translocated in the xylem, ft-om which it escapes to infect unwounded leaf cells. The key to understanding the process is understanding the components of the virus, beetle, and plant that interact with each other to effect virus infection. The presence of RNase in regurgitant appears to limit transmission of most plant viruses by beetles. Another key factor in the transmission process is that the type of feeding damage caused by beetles allows virus to enter the xylem of the plant. The viral and plant components that regulate interaction of virus (both beetle-transmissible and non-beetle-transmissible) with the plant after deposition by beetles are not yet known.
References 1. Markham, R., and Smith, K.M. (1949). Studies on the virus of turnip yellow mosaic. Parasitology 39, 330-342. 2. Nault, L.R., Styer, W.E., Coffey, M.E., Gordon, D.T., Negi, L.S., andNiblett, C.L. (1978). Transmission of maize chlorotic mottle virus by chrysomelid beetles. Phytopathology 68,1071-1074. 3. Serjeant, E.R (1967). Some properties of cocksfoot mottle virus. Ann. Appl. Biol. 59, 31-38.
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4. Freitag, J.H. (1956). Beetle transmission, host range, and properties of squash mosaic virus. Phytopathology 46, 73-81. 5. Fulton, J.P., Scott, H.A., and Gamez, R. (1980). Beetles. In "Vectors of Plant Pathogens" (K.F. Harris and K. Maramorosch, eds.), pp. 115-132. Academic Press, San Diego. 6. Bakker, W. (1973). Rice yellow mottle virus. CMI/AAB Descriptions of Plant Viruses. No. 149. 7. Catherall, PL. (1970). Phleum mottle virus. Plant Pathol 19, 101-103. 8. Langham, M.A.C., Gergerich, R.C., and Scott, H.A. (1990). Conversion of comovirus electrophoretic forms by leaf-feeding beetles. Phytopathology 80, 900-906. 9. Ghabrial, S.A., and Schultz, F.J. (1983). Seroldgical detection of bean pod mottle virus in bean leaf beetles. Phytopathology 73,480-483. 10. Fischer, D.C., and Kogan, M. (1986). Chemoreceptors of adult Mexican bean beetles: External morphology and role in food preference. Entomol. Exp. Appl. 40, 3-12. 11. McLaughlin, M.R., and Gilbrath, G.M. (1978). Transmission of southern bean mosaic virus by the striped cucumber beetle, Acalymma vittata. (Abstr.) Proc. Am. Phytopathol. Soc. 4, 129-130. 12. Scott, H.A., and Fulton, J.P (1978). Comparison of the relationships of southern bean mosaic virus and the cowpea strain of tobacco mosaic virus with the bean leaf beetle. Virology 84, 207-209. 13. Slack, S.A., and Scott, H.A. (1971). Hemolymph as a reservoir for the cowpea strain of southern bean mosaic virus in the bean leaf beetle. Phytopathology 61, 538-540. 13a. Selman, B.J. (1973). Beetles—^phytophagous Coleoptera. In "Viruses and Invertebrates" (A.J. Gibbs, ed.), pp. 157-177. American Elsevier Pubhshing Co., New York. 14. Wang, R.Y., Gergerich, R.C., and Kim, K.S. (1992). Noncirculative transmission of plant viruses by leaf-feeding beetles. Phytopathology 82, 946-950. 15. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Morphology of sweet potato whitefly, Bemisia tabaci (Homoptera: Aleyrodidae) relative to virus transmission. Zoomorphology 116, 143-156. 16. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Anatomy of a virus vector. In ''Bemisia 1995: Taxonomy, Biology, Damage, Control, and Management" (D. Gerling and R.T. Mayer, eds.), pp. 289-318. Intercept Ltd., Andover, U.K. 17. Wang, R.Y., Gergerich, R.C., and Kim, K.S. (1994). Entry of ingested plant viruses into the hemocoel of the plant virus vector Diabrotica undecimpunctata howardi. Phytopathology 84, 147-152. 18. Slack, S.A., and Fulton, J.P. (1971). Some plant virus-beetle vector relations. Virology 43, 728-729. 19. Fulton, J.P, Gergerich, R.C., and Scott, H.A. (1987). Beetle transmission of plant viruses. Annu. Rev. Phytopathol. 25, 111-123. 20. Gergerich, R.C., and Scott, H.A. (1991). Determinants in the specificity of virus transmission by leaf-feeding beetles. In "Advances in Disease Vector Research" (K.F. Harris, ed.). Vol. 8, pp. 1-13. Springer-Verlag, New York. 21. Walters, H.J., Lee, F.N., and Jackson, K.E. (1972). Overwintering of bean pod mottle virus in bean leaf beetles. (Abstr.) Phytopathology 62, 808. 22. Mueller, A.J., and Haddox, A.W (1980). Observations on seasonal development of bean leaf beetle, Cerotoma trifurcata (Forster), and incidence of bean pod mottle virus in Arkansas soybean. J. Ga. Entomol. Soc. 15, 398. 23. Wang, R.Y., Gergerich, R.C., and Kim, K.S. (1994). The relationship between feeding and virus retention time in beetle transmission of plant viruses. Phytopathology 84, 995-998.
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24. Field, T.K., Patterson, C.A., Gergerich, R.C., and Kim, K.S. (1994). Fate of viruses in bean leaves after deposition by Epilachna varivestis, a beetle vector of plant viruses. Phytopathology 84, 1346-1350. 25. Gergerich, R.C., Scott, H.A., and Fulton, J.R (1983). Regurgitant as a determinant of specificity in the transmission of plant viruses by beetles. Phytopathology 73, 936-938. 26. Srivastava, U.S. (1959). The maxillary glands of some Coleoptera. Proc. R. Entomol. Soc. Lond. 34, 57-62. 27. Wang, R.Y. (1994c). Studies of virus circulativeness and retention in beetle vectors of plant viruses. Ph.D. Dissertation, University of Arkansas, Fayetteville, 106 pp. 28. Gergerich, R.C., Scott, H.A., and Fulton, J.R (1986). Evidence that ribonuclease in beetle regurgitant determines the transmission of plant viruses. J. Gen. Virol. 67, 367-370. 29. Monis, I , Scott, H.A., and Gergerich, R.C. (1986). Effect of beetle regurgitant on plant virus transmission using the gross wounding technique. Phytopathology 76, 808-811. 30. Pirone, T.P. (1991). Viral genes and gene products that determine insect transmissibility. Semin. F/ra/. 2,81-87. 31. Gildow, F.E., and Gray, S.M. (1993). The aphid salivary gland basal lamina as a selective barrier associated with vector-specific transmission of barley yellow dwarf luteoviruses. Phytopathology S3, 1293-1302. 32. Ehara, Y., and Mink. G.I. (1980). Cucumber mosaic virus and peanut stunt virus differ in an initial event that occurs during the infection of cowpea leaf epidermal cells. Virology 104,258-261. 33. Lane, L.C. (1974). The bromoviruses. Adv. Virus Res. 19, 151-220. 34. Gergerich, R.C, and Scott, H.A. (1988b). Evidence that virus translocation and virus infection of non-wounded cells are associated with transmissibility by leaf-feeding beetles. J. Gen. Virol. 69, 2935-2938. 35. Gaard, G., and de Zoeten, G.A. (1979). Plant virus uncoating as a result of virus-cell wall interactions. Virology 96, 21-31. 36. Mahmood, T., Gergerich, R.C, and Kim, K.S. (1993). Coat protein as a determinant of virus transmission by leaf-feeding beetles. (Abstr.) Phytopathology 83, 1375. 37. Schneider, I.R., and Worley, J.F. (1959). Rapid entry of infectious particles of southern bean mosaic virus into living cells following transport of the particles in the water stream. Virology 8, 243-249. 38. Opalka, N., Brugidou, C , Bonneau, C , Nicole, M., Beachy, R.N., Yeager, M., and Fauquet, C (1998). Movement of rice yellow mottle virus between xylem cells through pit membranes. Proc. Natl. Acad Sci. U.S.A. 95, 3323-3328. 39. Gergerich, R.C, Scott, H.A., and Wickizer, S.L. (1991). Determination of host resistance to beetle transmission of plant viruses. Phytopathology 81, 1326-1329. 40. Hobbs, H.A., and Kuhn, CW. (1987). Differential field infection of cowpea genotypes by southem bean mosaic. Phytopathology 77, 136-139.
CHAPTER 8
Caulimoviruses STEPHANE BLANC EUGENIE HEBRARD MARTIN DRUCKER REMY FROISSART
/- Introduction Because of the structure of plants, viruses require some form of plant damage to enter and infect a healthy host; therefore they have evolved a wide variety of strategies for their transmission from one plant host to the next. The most common means of transmission encountered among these viruses is vector transmission, in which virus transport is ensured by plant-feeding organisms that frequently change their host plant and even travel among different plant populations. A wide range of organisms has been used as vectors by different plant viruses (for a recent review, see [1]), and in all cases vector transmission can be defined by a cascade of phases and events: (1) the acquisition phase, during which the vector contacts an infected plant and picks up virus together with plant cell components and the virus reaches specific location(s) in or on the vector; (2) the retention phase, during which the virus establishes a specific molecular interaction with the vector that allows the virus to get ready for inoculation and remain ready long enough for the vector to move to another plant; and (3) the inoculation phase, during which the vector injects the virus in a host plant in an infective form. Virus inoculated by a vector in a noninfectious form, for whatever reason, is considered not transmitted. The best known vectors are insects in the order Homoptera, especially aphids. Aphid transmission has been studied and discussed thoroughly for over 50 years, and the classification of the different types of virus-vector interactions is still subject to modifications and adjustments to integrate new data and concepts [1, Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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2]. In contrast to some viruses that cross cellular barriers and complete a cycle in their vector's body (circulative), others are noncirculative [3,4]. Hull [1] categorizes noncirculative viruses as "externally borne," whereas Harris et al [5, 6] prefer "cuticula-bome" to connote that transmissible virions are carried externally (i.e., anterior to the os) at specific retention sites on the cuticula lining the lumen of the feeding apparatus. Based on thorough analysis of the morphology of the homopteran feeding apparatus, Harris et al [5, 6] concluded that virus located past the OS in the intima-lined pharynx of the foregut can no longer be egested and is therefore noninoculable. Two subcategories of noncirculative viruses, nonpersistent [7] and semipersistent [8], are usually differentiated on the basis of the time required for virus acquisition and inoculation, as well as the length of time that the vector can retain virus in an infectious form. All three parameters are shorter for nonpersistent than for semipersistent viruses. However, whereas results of acquisition, retention, and inoculation optimum times are quite consistent for nonpersistent viruses, in some cases variable results have been obtained for semipersistent viruses and, in particular, for cauliflower mosaic virus (CaMV) [9]. This led Day and Venables [10] to describe semipersistent transmission as an ensemble of atypical cases of nonpersistent transmission. Harris [4] interpreted this subcategory of noncirculative transmission as the result of the combined effect of virus distribution within plant tissues and feeding behavior of vectors. For example, a virus found only in the vascular tissue will be reached later by the vector than one present in the epidermal layers. For further details on the feeding behavior of vectors in relation to transmission of plant viruses, readers are referred to chapter 5. It is important to note that, even at present, there is no conclusive evidence of significant differences between the molecular mechanism of virus-vector interaction in nonpersistent and semipersistent transmission. This explains why, in a recent review, Pirone and Blanc [11] did not discriminate between these two transmission types. Rather, these authors used recent data on the molecular mechanisms of virus-vector interactions as the main criteria for defining two distinct viral strategies, the capsid strategy and the helper strategy. In the capsid strategy, the virus interacts directly with the vector via its coat protein, whereas in the helper strategy the virus-vector interaction is mediated by an additional virus-encoded nonstructural protein generally designated as helper. The helper strategy is frequently found among noncirculative viruses in the genera Potyvirus, Caulimovirus, Waikavirus, Sequivirus, and presumably Closterovirus and even the nematode-transmitted Tobravirus [11]. However, the helper strategy has been extensively studied only in the genera Potyvirus and Caulimovirus. In both groups, the molecular mechanisms of virus-vector interaction have been elucidated by researching the mode of action of the helper. For decades, a single model was believed to apply to all helpers, but we now have experimental evidence that the helpers in different virus groups may not all function similarly. This is illustrated by comparing the aphid transmission of
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caulimoviruses, the primary focus of the present review, with that of potyviruses, addressed in chapter 9.
//.
The Virus
Members of the genus Caulimovirus are DNA viruses with a pregenomic RNA template for replication via reverse transcription. This genus belongs to the pararetroviruses, a supergroup including caulimo-, badna- and hepadnaviruses. This supergroup is a paraphyletic taxon thought to be phylogenetically close to retroviruses. The two most important differences between retro- and pararetroviruses are that (1) the transmissible virus particles of the latter contain doublestranded DNA, whereas those of retroviruses contain RNA and (2) whereas retroviruses integrate their genetic material in the cellular genome [12], pararetroviral DNA usually accumulates within the host cell's nucleus as multiple copies of a minichromosome, although one case of integration was recently reported [13]. On the basis of the similarities in reverse transcriptase sequences [14] and structural and functional criteria [15], caulimo- and badnaviruses seem to be phylogenetically closer to retroviruses than are hepadnaviruses. Caulimoviruses infect only dicots, and all known vectors belong to the family Aphidinae. Cauliflower mosaic virus (CaMV) represents the t3^e species of this genus and has been the focus of most studies on caulimoviruses for over 30 years. Most information on caulimovirus biology was obtained with CaMy although some data concerning other members of the group recently became available and show that their genome organization and gene expression strategies are similar to those of CaMV [16]. Most important for the present review is the work of Markham and Hull [17] demonstrating that similar virus-vector relationships are involved in transmission of various caulimoviruses. These authors showed that the helpers controlling aphid transmission of figwort mosaic caulimovirus (FMV) and carnation etch ring caulimovirus (CERV) were equally able to mediate the aphid transmission of CaMV Since then, all further studies on the molecular mechanisms of caulimovirus aphid transmission have been carried out on CaMV This is the reason why this review will focus on the CaMV system nearly exclusively A.
Genome Organization and Gene Expression Strategies
The CaMV genome is a double-stranded DNA, containing between 7,610 and 8,033 base pairs, depending on the virus strain. The DNA is circular, with three sequence discontinuities where the strands are not covalently linked, and contains seven major open reading frames (ORFs) separated by few intergenic regions [16]. Two intergenic regions have been extensively characterized, one corresponding to the 35S promoter, which controls the transcription of the pregenomic 35S
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Fig. 1 Genetic map of cauliflower mosaic virus. Thick arrows: Open reading frames (ORFs) encoding proteins (for function, see text). Thin arrows: The two major viral transcripts. Thin line: Intergenic regions.
RNA, and the other corresponding to the 19S promoter, which directs synthesis of a 19S subgenomic RNA (Fig. 1). The 35S RNA is the template for reverse transcription and serves as a polycistronic messenger RNA for most viral gene products. The monocistronic 19S RNA is translated by the host's cellular machinery, independently of the polycistronic 35 S RNA. Additional minor RNA species have been described and are suspected to be either mono- [18] or polycistronic RNAs originating from putative additional promoter(s) [19] or from splicing of the pregenomic 35S mRNA [20]. The amount of RNA produced may vary depending on the infected tissue in the host plant. For instance, Benfey and Chua [21] demonstrated that, in addition to a core containing the TATA box, the 35S promoter exhibits two domains involved in tissue or organ specificity or both and defined as region A for expression in roots and region B for expression in leaves. As mentioned above, CaMV 35S RNA is a polycistronic mRNA. Translation of several successive ORFs from a single mRNA is usually very inefficient in eukaryotic cells, because only 5' proximal ATG start codons are used for initiating translation by scanning ribosomes (the regular scanning model in eukaryotic cells is reviewed in [22]). Polycistronic translation of CaMV 35S RNA in host plants has been thoroughly studied and several specific mechanisms have been described: (1) ribosome shunt, allowing the bypass of several short untranslated ORFs located in the 35S leader sequence (a feature known to be inhibitory for downstream translation [23, 24]); and (2) relay race [25, 26], in which ribosomes reinitiate protein synthesis at the nearest start codon after passing a termination codon. Fiitterer and Hohn [27] demonstrated that the latter process plays an important role in CaMV 35S RNA translation and depends on the presence of a
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viral translation trans activator (TAV). Although leaky scanning has never been described in caulimoviruses, it is frequently used to express different proteins from viral RNA [28] and has been reported for the translation of the polycistronic pregenomic RNA of a badnavirus [29]. The genome of caulimoviruses contains seven major ORFs, of which all but ORF VI are expressed from the 35S RNA. Functions associated with various CaMV gene products have recently been extensively reviewed [16] and are briefly summarized as follows. ORF I codes for PI protein involved in cell-to-cell movement and systemic spread. ORF II encodes the CaMV helper, also designated P2 or aphid transmission factor (ATF), and is the main focus of the present review. Polypeptides resulting from translation of spliced RNAs and corresponding to inframe frisions between ORFsI and II (three splice donor sites in ORF I and one splice acceptor site in ORF II) have been detected in infected plants. (The functions of such fusions, if any, remain to be determined and will not be further discussed in this review.) No clear biological function had been attributed to the ORF III product (P3) until the very recent demonstration of its involvement in aphid transmission. This discovery, which surprisingly revealed P3 as being as important as P2 in aphid transmission of CaMV, will be discussed in detail below. ORF IV codes for a precursor protein (P4), which is posttranslationally cleaved to give a set of structural polypeptides making up the viral capsid. The molecular weight of the main capsid components ranges between 35 and 44 kDa. ORF V codes for the reverse transcriptase (C-terminal domain) and is also involved in P4 maturation via an aspartic acid proteinase activity associated to its N-terminus. ORF VI is translated from the 19S RNA and codes for a multifunctional protein (P6). The most documented frmction of P6 (also designated TAV, see above) is that it transactivates the expression of the other genes from the polycistronic 35S RNA, especially that of ORFs III, IV, and V The protein P6 is also the major component of the so-called vacuolar inclusion bodies which is the site of viral DNA synthesis, accumulation of all virus gene products, and virus assembly. Moreover, P6 seems to have a nuniber of additional effects (either direct or indirect) on the CaMV infection cycle [16]. The product of ORF VII has never been found in planta; hence the role of this gene remains a mystery. The kinetics of protein expression have been studied by several authors and the results were very consistent whether the experiments were carried out on entire plants [30, 31] or with protoplasts [32, 33]. The proteins PI, P5, and P6 are expressed earlier than P3, whereas P2 and P4 are the late accumulating proteins [32]. These differences in expression timing suggest that each gene may be regulated differentially. B.
The Cauliflower Mosaic Virus Infection Cycle
After entry into a host cell, the viral DNA reaches the nucleus, and the discontinuities on both DNA strands are repaired to yield a minichromosome-like form of the CaMV genome, which is the template for transcription. The 19S and 35S
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RNAs migrate into the cytoplasm to trigger protein expression and virus replication. Reverse transcription and viral DNA packaging seem to take place in the vacuolar inclusion bodies of infected host cells [34]. There are two sorts of inclusion bodies, both located in the cytoplasm: (1) the electron-dense or vacuolar inclusion bodies (EDib), which have a matrix composed mainly of P6 [35] and are where most, if not all, viral genes are expressed and virus particles accumulate, and (2) the electron-lucent inclusion bodies (ELib), whose matrix is almost exclusively composed of P2 and contains only a few scattered virus particles [36]. Although the latter may represent the transmissible complex, no definitive function has been demonstrated for this type of inclusion body. Virus particles somehow move away from the EDib and migrate to the plasmodesmata, through which, with the help of PI, they colonize the adjacent cells or enter the vascular tissues. The frequent appearance of variants has been described during virus DNA replication [37]. An analysis of these variants revealed many modifications within the sequence of gene II. Interestingly, deletions or point mutations within gene II were similar to variations previously described in naturally occurring isolates, thus indicating that this type of variation may be a recurrent phenomenon during the CaMV replication cycle. Gene II is known to be the only gene dispensable for virus infection [38], although it is required for aphid transmission. It is possible that frequent variations within gene II, particularly deletions, are observed because they are not counterselected owing to the dispensability of this gene in experiments using mechanical inoculation. Genomic DNA recombination events have been described in CaMV; coinfection of host plants with two noninfectious, full-genome-length clones (each altered in a different region) yielded fully functional recombinant progeny [39]. Recombination can occur in various regions of the CaMV genome and seems to be rather frequent during the replication cycle [39-41]. However, most experiments either were done under selection pressure (only a recombination event could restore viability of the progeny, the parental genomes being noninfectious and therefore counterselected) or consisted of sequence analysis allowing the detection of genomes resulting from past recombination events. Hence, in the absence of selection pressure, a precise evaluation of the recombination rate has never been proposed.
///. Biology of Caulimovirus Transmission by Aphids A.
Classifying Cauliflower Mosaic Virus Transmission
Cauliflower mosaic virus is transmitted by at least 27 aphid species [3], the two most studied vectors being Myzus persicae and Brevicoryne brassicae. Transmission of CaMV via seeds or pollen has never been reported in nature, nor has transmission by any other type of vector or by plant-to-plant contact. In laboratories, CaMV is readily inoculated mechanically to host plants [42].
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In early work on CaMV aphid transmission, it was noted that the relationships between the virus and its vector were showing "irregularity" [9]. Presumably as a result of this atypical behavior in aphid transmission, different authors have not always agreed on whether CaMV transmission was to be considered as nonpersistent or semipersistent [3, 9, 43]. Although they concluded that CaMV transmission by M. persicae resembled typical nonpersistent transmission, Chalfant and Chapman [44] reported puzzling observations when B. brassicae was used as a vector. In this case, CaMV exhibited a very unusual "bimodal" pattern, showing features of both nonpersistent and persistent (also called circulative) viruses. When assessing the effect of the duration of the acquisition feed on virus transmission, they noted that there were two optimal time points, one after a few minutes (characteristic of nonpersistent transmission) and the other after several hours (more characteristic of persistent transmission). A similar phenomenon was later reported for pea seed-borne mosaic potyvirus (PSbMV) transmission by the "New England" biotype of Macrosiphum euphorbiae [45]. Not all vectors transmit CaMV or PSbMV in a bimodal manner, thus demonstrating that this unusual type of transmission is not a feature attributable to the virus itself Several authors have discussed this point, and it appears that bimodal transmission is not a combination of nonpersistency and persistency, but rather results from variations in noncirculative transmission due to the influence of many factors, such as aphid saliva, virus distribution in the plant, and vector behavior, plus a number of unknown others [45-47]. Markham et al [47] finally put an end to this question of bimodality by thoroughly reinvestigating CaMV aphid transmission by several aphid species and comparing the results with those for a typical nonpersistently transmitted potyvirus. Although an irregular pattern in various CaMV aphid transmission trials was consistently observed, the authors used a statistical analysis of their results to rule out the existence of a true bimodal phenomenon. They thereby demonstrated definitively that CaMV is transmitted in a semipersistent manner. The last decade has brought forth much information on the molecular interactions between virus and vector during noncirculative transmission. We believe that, preferentially, the viral molecular strategy for transmission should be taken into consideration. Consequently, we favor the use of capsid and helper strategy classifications, as proposed by Pirone and Blanc [11]. According to that view and owing to the fact that the requirement of a helper factor (described in detail in the following sections) has been demonstrated for CaMV, we prefer stating that it is transmitted in a noncirculative manner following the helper strategy. B.
Evidence for Helper Dependency in Aphid Transmission of CaMV
The first puzzling information that paved the way for the discovery of helper dependency in aphid transmission of CaMV came from Pirone and Megahed [48], who demonstrated that aphid transmissibility was lost upon virus purifica-
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tion. As for potyviruses, the characterization of nontransmissible isolates threw some light on this phenomenon. Lung and Pirone [49] reported convincing evidence that the lack of aphid transmission of two CaMV isolates (Campbell and CM 1841) could not be attributed to differential accumulation or distribution of the virus in infected leaves, nor was it attributable to some alteration of the purified virus particles. These authors demonstrated that these two CaMV isolates are in fact transmitted from plant to plant by aphids when virus-source plants are coinfected with a transmissible isolate (cabbage B). Campbell isolate was also transmitted from singly infected plants when aphids were first fed on a plant infected with cabbage B, but not when the acquisition sequence was reversed. In agreement with similar observations reported earlier on potyviruses [50, 51], Lung and Pirone [49] concluded that a helper or aphid transmission factor (ATF) was also required in caulimovirus transmission. This factor could be the virion itself or another factor produced in plants infected with a transmissible strain. A year later [52], Lung and Pirone confirmed these findings using purified CaMV preparations as the virus source. The precise acquisition sequence of various components by the aphids (ATF and purified virus simultaneously or ATF first and then purified virus) was confirmed to be essential for successful aphid transmission. Furthermore, when aphids probed first on purified cabbage B virions and then on purified Campbell virions, transmission did not occur. This indicated that the cabbage B virion itself was not the ATF. The similarity between these results and those obtained earlier on potyviruses [53] prompted an interesting experiment in which the ATF of caulimovirus was replaced with that of the potyvirus and vice versa. In that way. Lung and Pirone [52] clearly demonstrated that despite apparent similarities observed in the caulimo- and potyvirus systems, the ATFs (or helpers) involved in aphid transmission were different and specific for each virus group.
IV. Identification of Aphid Transmission Factor(s) A.
The Product of Cauliflower Mosaic Virus Gene II is the Aphid Transmission Factor
An important step toward the understanding of CaMV biology in general and of aphid transmission in particular was the discovery that purified DNA was infectious [54]. Simply rubbing leaves with purified CaMV DNA and an abrasive powder resulted in high rates of host plant infection. Even more important was the follow-up study demonstrating that a frill-length clone of the CaMV genome was similarly infectious [55]. Indeed, this discovery opened a whole new range of possibilities for research, such as chimeric virus construction and disruption of gene frmction by mutagenesis or gene replacement in the complete viral genome. This
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and the availability of several nontransmissible but infectious CaMV isolates allowed a number of authors to independently map the genome region responsible for aphid transmission [38, 56-58]. All results were consistent with the identification of the CaMV gene II as the gene encoding the aphid transmission factor. A polypeptide of 18 kDa (now called ATF or P2), copurifying with CaMV inclusion bodies, was found to be associated with the presence of an intact gene II in transmissible isolates [56, 58]. It was later demonstrated that some nontransmissible strains of CaMV (Campbell and CM 1841) were nevertheless producing P2 on plant infection [59] and that their lack of transmissibility was due to a single point mutation in P2 (glycine to arginine) at amino acid position 94. In vitro aphid transmission testing was used to confirm that the ATF biological activity was present in the viroplasm-enriched fraction [60] and associated with P2 [61]. However, because of recurrent failures in purifying P2 due to problems of solubility, these two studies were performed with rather crude extracts containing a large number of additional virus or host plant proteins. Hence, direct and definitive evidence that P2 is indeed the aphid transmission factor of CaMV was lacking until P2 was produced in a heterologous expression system. The expression of CaMV gene II in cultured insect cells using a baculovirus recombinant was first done by Espinoza et al [62]. Unfortunately, these authors encountered similar solubility-related problems during purification trials. Blanc et al. [63] further investigated this point and determined the conditions for recovery of biologically active ATF from crude extracts of cultured insect cells producing P2. Aphids that first acquired the baculovirus-expressed P2 became capable of transmitting several naturally nontransmissible isolates of CaMV This system marked a turning point in the study of CaMV aphid transmission, and most of the known biological properties of P2 described below were derived from it. B.
Cauliflower Mosaic Virus Aphid Transmission Requires an Additional Helper Factor
A surprising observation, reported by Blanc et al. [63], was that once the aphids were fed a baculovirus-expressed, P2-containing solution, they could transmit a naturally nontransmissible isolate of CaMV acquired from infected plants or crude extracts but not from purified virus preparations. This was puzzling because purified potyvirus particles are readily acquired and transmitted by aphids that first probe a solution containing highly purified HC-Pro (the transmission factor of potyviruses). The authors tentatively proposed two explanations: (1) the virus purification procedure somehow altered the CaMV coat protein, thus rendering it nonfunctional for aphid transmission, and (2) an additional unknown factor ("the missing factor"), originating either from plant or virus genome, was necessary for the transmission process and was lost upon CaMV purification. At that time, the latter explanation seemed unlikely because purified CaMV preparations had previously been described as suitable for virus
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acquisition by aphids in experiments in which P2 was provided from plants [52] or viroplasm-enriched fractions [60]. Nonetheless, a putative missing factor could not be totally ruled out, because it might have been acquired in these cases during the first feed of the aphid, together with P2. Following P2 feeding, aphids did not transmit purified CaMV mixed with crude extracts of healthy plants, nor did they transmit it from a mixture containing extracts from plants infected with an unrelated virus. In contrast, when purified CaMV Cabb B-JI (a transmissible isolate) virions were mixed with a crude extract from plants infected with the nontransmissible CM4-184 isolate, both were transmitted by aphids that had previously acquired P2 ("P2-loaded aphids"). This suggested that the putative missing factor was of viral origin (Blatt and Blanc, unpublished results). It was not until very recently that this problem was elegantly solved by Leh et al [64], who demonstrated that P2-loaded aphids transmitted highly purified virus particles only when they had been preincubated with the CaMV ORF III product, expressed from bacteria. Thus, these authors definitely and directly established the existence of an additional viral factor in the molecular mechanisms of CaMV transmission by aphids. This result is of fiirther relevance because it represents the first demonstration of a possible involvement of two aphid transmission factors in the helper strategy of noncirculative viruses. It also assigns the first described biological function to CaMV ORF III.
V. Characterization of Aphid Transmission Factor(s) A.
Biological Properties of P2
The product of CaMV ORF II is a polypeptide of 18 kDa (PIS, also designated here as P2 or ATF) consisting of 159 amino acid residues. Negative staining and immunoelectron microscopy demonstrated that P2 expressed in insect cells via a baculovirus recombinant can self-assemble into a highly organized paracrystalline structure [65] (Fig. 2). Paracrystals were first described in infected plant cells as arc-shaped [66]. Blanc et al. [65] also observed paracrystals in partially purified CaMV inclusion bodies. Two types of inclusions were observed, one clearly corresponding to EDib (mentioned above) and the other consisting of large paracrystalline aggregates containing a few scattered virions. The latter was somewhat similar to the ELib described by Espinoza et al. [36]. The characteristic paracrystalline structure indicating the presence of P2 allowed its unequivocal detection close to, or even inside, intact or partially degraded EDib (Blanc and Louis, unpublished). This observation renews the controversy of whether P2 is present in the EDib [58] or is exclusively located in the ELib [36]. To resolve this apparent discrepancy, Pirone and Blanc [11] proposed that the two types of inclusions could represent different stages in the maturation of the CaMV viroplasm. Furthermore, since the two inclusion types are often observed close to each other
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Fig. 2 Cauliflower mosaic vims (isolate Cabb B-JI) aphid transmission factor (ATF) paracrystal. P2 produced in Sf9 cells via a baculovirus recombinant was extracted with phosphate-buffered saline prior to negative staining with ammonium molybdate. (A) Paracrystal. (B) Detailed view: A paracrystal consists of a repetitive series of parallel layers (black arrowheads) of closely associated subunits (white arrowheads), the interlayer and intersubunit periodicity being 13 nm and 4 nm, respectively. (C) Paracrystals colocalize with microtubules. The gray arrowheads point to microtubule protofilaments. (Bar = 50 nm in A and C, and 20 nm in B.)
or even in contact [36, 62], the ELib might originate from the EDib via protein (P2) export. Indeed, the latter is believed to be the site of all viral protein synthesis. If this should be true, it would be easy to imagine an intermediate-stage EDib in which, in addition to P6, some P2 could also be detected. The paracrystals consist of a repetitive series of parallel layers (interlayer periodicity of 13 nm) of closely associated subunits. The distance between adjacent subunits within a layer is 4 nm. Each subunit within one layer seems to be connected to the corresponding subunit of the next layer by a filament-like structure. The paracrystals could be solubilized in the presence of high EGTA concentrations to provide the biologically active form of the CaMV aphid transmission factor, which in turn could be specifically reassembled into typical paracrystals on addition of calcium chloride and removal of EGTA [65]. Because of the lack of a satisfactory purification protocol for P2, the foregoing experiments were carried out in insect cell crude extracts that contained many contaminants. Hence, one cannot definitely conclude from the experiments that P2 is the exclusive constituent of the observed paracrystals. Among many P2 mutants analyzed, those able to form paracrystals are biologically active when solubilized, whereas all variants unable to form paracrystals
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remain inactive on solubilization (Blanc, unpublished). Nevertheless, experimental proof of a specific function of the paracrystals in aphid transmission is still missing, and the domains involved in their formation have not yet been identified. The propensity of P2 to form paracrystals is the main feature that permitted the discovery of its most striking property. Blanc et al. [67] reported the existence of a specific interaction between P2 and the microtubular network in P2-producing cells. Although this interaction seems surprising and its significance is not yet understood, it cannot be overlooked in the present review. In insect cells infected with a baculovirus recombinant expressing P2, a very large amount of modified microtubules (MT) was observed in the cytoplasm. These microtubules were thicker than normal (35 to 40 nm instead of 25 nm), and P2-specific gold-labeling of thin sections of infected cells demonstrated that P2 colocalized with the microtubule-like elements. This microtubular network, consisting of microtubules coated with P2, is extraordinarily stable and is the last remaining structure in the culture medium that is still easily detectable 3 days after cell death. When its stability was assessed in vitro, P2-microtubule complexes remained intact for 6 hours under various microtubule depolymerizing conditions such as low temperature, dilution, and calcium, or a combination of all three. The P2 produced in infected insect cells is partially solubilized only in the presence of 500 mM NaCl in the extraction buffer. At this high ionic strength, P2 still binds to taxol-stabilized insect and mammalian microtubules. This property seems to be unique among the known microtubulebinding proteins, which are usually released from microtubules at this ionic strength. Indirect immunofluorescent double labeling of P2 and tubulin in tobacco protoplasts demonstrated that P2 expressed via the tomato bushy stunt virus expression system [68] also interacted with plant cell microtubules [67]. The P2 domain responsible for the interaction with microtubules has not yet been determined. Some of our unpublished preliminary results, however, indicate that at least two domains are involved, located near the N- and C-termini, respectively. Further work is necessary to characterize the microtubule-binding domain and to investigate the P2-microtubule interaction during the CaMV infection cycle. Relationships between the host cell cytoskeleton and plant viruses are very poorly documented except for some viral cell-to-cell movement proteins reported to colocalize with microtubules in infected plant cells [69-71]. Although its precise role has not yet been defined, the interaction between microtubules and various cellto-cell movement proteins is somehow understandable. Cell-to-cell movement implies virus transport within or between cells, and the pivotal role of microtubules in the transport of protein and organelles within a cell is well established [72]. In contrast, the role of the CaMV aphid transmission factor seemingly comes into play inside the aphid stylets. It is hard to imagine that microtubules could intervene in an extracellular context. Nevertheless, several hypotheses can be suggested for integrating the microtubule-P2 interaction in the molecular mechanisms of CaMV transmission by aphids. This will be further detailed in section VI. Using various truncated bacterially expressed P2 fusions, Schmidt et al. [73] demonstrated that a 31 amino acid C-terminal domain of P2 specifically interacts
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with partially purified virus particles. Interestingly, the very same domain has recently been reported to interact also with P3 [64]. In further investigating this phenomenon, Leh et al. of the same group have convincingly demonstrated that P2 does not bind directly to highly purified CaMV particles but does so through an attachment to P3, which in turn is associated with coat protein [74]. The organization of P2 into paracrystalline structures is undeniable evidence of the existence of one or several domains involved in self-interaction. Preliminary results indicate that a C-terminal domain of P2 can interact with a homologous domain on another molecule (Blanc and associates unpublished). Predictions of secondary structures of the C-terminus of P2 anticipate the occurrence of two ahelices with a high probability of coiled coil formation [64]. Coiled coil structures result from the association of two or more a-helices and are shaped like braids. They can comprise a-helices originating from one or several molecules [75]. Taken together, these data indicate that P2 interacts with itself via a region located in the C-terminal half, which totally overlaps with the region involved in the interaction with P3. Furthermore, the microtubule-binding domain seems to be composed of at least two motifs of P2 located near the N- and C-termini, respectively, the latter overlapping again with the region of P2 involved in interaction with itself and with P3. This raises the possibility of precisely tuned interactions among P2, P3, and microtubules. Information on the three-dimensional structure of these protein partners would help elucidate their complicated interactions. Unfortunately, the lack of a satisfactory purification procedure has so far precluded all attempts to study the structure of P2. B.
Biological Properties of P3
Open reading frame III codes for a 129-amino-acid, basic protein of 15 kDa molecular mass. The corresponding protein P3 was initially recognized by genetic analysis as being indispensable for viability-infectivity of CaMV [76, 77]. Later on, it was identified as being associated with the icosahedral CaMV capsid, which is basically built of P4 [78, 79]. Studies based on Escherichia c6>//-expressed P3 revealed that it contains a domain that nonspecifically binds nucleic acids, mapped by deletion and substitution analysis to the C-terminal amino acid residues 112-121 [80]. Further work by Jacquot et al [81], who examined systematically introduced deletions in ORF III of the complete genome, revealed that all but amino acids 61-80 were required for infectivity-viability of CaMV The nucleic acid-binding domain seemed not to be the only one involved in infectivity, because nonviable P3 mutants that still bound nucleic acids were also characterized. The N-terminal coiled coil structure and oligomerization of P3 into a parallel tetramer was elegantly demonstrated by Leclerc et al. [82], who analyzed a synthetic oligopeptide corresponding to the 29 N-terminal amino acids of P3. These authors confirmed their results by showing that the P3 N-terminus interacted with itself in yeast two-hybrid screen and by analytical ultracentrifrigation of bacterially expressed wild-type P3.
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An important advance in understanding the biological function(s) of P3 was achieved by Leh et al [64], who established that P3 was required for CaMV aphid transmission and more precisely, that amino acids 1-26 were essential in this regard. They further demonstrated by protein overlay assays that this N-terminal region of P3 was implicated in binding to P2, which suggested a link between the ATF function of P2 and P3. In this report, the authors proposed, among other hypotheses, that P3 could mediate an indirect attachment of P2 to virions, thus intuitively suggesting an interaction between P3 and the capsid protein (P4). This crucial point for elucidating the mode of action of P3 was experimentally proved by Leh et al [74] using similar protein overlay assays. It was demonstrated that P3 binds not only to highly purified virions but also to bacterially expressed P4, which thus indicated that the capsid protein domain involved in this interaction is unlikely to result from the association of two or more capsomers in the mature virus particles. The P3 region responsible for capsid binding was mapped to a large domain located in the C-terminal half and spanning amino acids 60-110. Disrupting this interaction systematically also impaired the aphid transmission activity of the corresponding P3 mutants. Taken together, these data derived from biochemical analysis, mutagenesis, and protein interaction assays, when combined with structure predictions, suggest a bipartite organization [81] for P3: with an N-terminal, a-helical, leucine-rich, coiled coil implicated in P3 self-interaction (oligomerization) and binding to P2, and a C-terminal half involved in coat protein and nucleic acid binding. The two parts are presumably separated by a central flexible region [82]. As a concluding comment on the biological properties of the CaMV ATFs, it is interesting to note that the P3 region involved in P2 binding is also involved in self-interaction, the very same feature that was emphasized for P2 in the previous section. This puzzling observation opens the way for the putative regulation mechanisms discussed in section VII.
W. Mode of Action of Aphid Transmission Factor(s) A.
The Bridge Hypothesis
A simple but convincing model to explain helper-mediated noncirculative transmission of plant viruses by aphids was developed over a period of 25 years. Because it was mainly obtained from data on potyviruses, we will first briefly summarize this model and its supporting evidence and then discuss its possible extrapolation to caulimoviruses. The readers are referred to chapter 9 for a comprehensive review on potyvirus aphid transmission. The model, first introduced by Govier and Kassanis [53], was named the bridge hypothesis by Pirone and Blanc [11]. The bridge hypothesis presumes that the
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Aphid ci^ulartifitff withl)Mrf-$it0s() Fig. 3 Schematic representation of binding of a potyvirus and cauliflower mosaic virus (CaMV) to the aphid's mouthparts. The viruses are drawn in a median section to delineate the rod-shaped structure of potyviruses compared with the icosahedral structure of CaMV The light gray polygons represent the virus capsids. Of the potyvirus only a short part is shown; its extensions are indicated at the ends. The binding of potyviruses is mediated by a single protein (HC-Pro), whereas the CaMV virion is attached to the aphid binding site via two proteins, P2 and P3. Note that owing to its elongated shape, the potyvirus particle may be attached to the aphid by several HC-Pro molecules, whereas the CaMV virion probably forms only one "bridge" with the vector's binding sites.
bridge component (helper protein HC-Pro for potyviruses or ATF for CaMV) contains two domains, specifically binding to aphid stylets and to virus coat protein, respectively, and thus "gluing" the virus to the aphid's mouthparts (Fig. 3). This provision coincided with data on zucchini yellows mosaic potyvirus and tobacco etch potyvirus showing that the HC-Pro contained an N-terminal domain, which was most probably implicated in interaction with the aphid [83] and a Cterminal domain, which was responsible for virus binding [84]. The bridge hypothesis further requires that should only one of these domains (or their respective binding sites in the aphid or on the virus particle) be nonfunctional, transmission will not occur. The latter has been demonstrated for many pot3rvirus mutants (for an extensive review, see chapter 9). Direct evidence for HC-Pro-mediated retention of potyvirus particles has been microscopically demonstrated by Pirone's group [85-87]. The bridge hypothesis is now generally accepted and has been adapted to other noncirculatively transmitted plant viruses as well. B.
The Case of Caulimoviruses
The bridge hypothesis was also introduced to explain aphid transmission of viruses in the genus Caulimovirus. Earlier results were in agreement with the original concept of the bridge hypothesis, but recent findings by Leh et ah [64]
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indicate that in the case of CaMV and probably also other caulimoviruses, aphidmediated transmission cannot be explained by a single protein linking the virions to the aphid's cuticular surfaces. Instead, both P2 (the originally identified CaMV ATF) and P3 in concert are involved in transmission. Two modes of action were proposed in this work: either P2 would bind to P3 already attached to P4 (P3 serving as an adapter) or P3 would bind to P2 and render it competent for attaching to the virus coat protein P4. Since a P4-binding region, required for aphid transmission, was recently identified in the C-terminal half of P3 [74], the latter hypothesis seems highly improbable. Hence, the bridge hypothesis must be modified to fit aphid transmission of CaMV Whereas in the potyvirus system a single viral protein (HC-Pro) is sufficient to act as a bridge between the virus coat and the aphid binding sites, in caulimoviruses two proteins together seem to fulfill the ATF function: P2 mediates binding between the aphid's cuticular surface and P3, whereas the virus coatassociating protein P3 mediates binding of P2 to the virus particle (Fig. 3). This double-layered model provides a more tightly regulated attachment of CaMV to the vector, and consequently more tightly regulated transmission, than does the single protein link system of potyviruses. Preliminary results indicate that a precise acquisition sequence of all three components (P2, P3, and virion) by aphids is required for successful transmission (Froissart, Drucker, and Blanc, unpublished data). However, further work is needed to understand the kinetics of formation of the transmissible complex, which could occur in CaMV-infected leaves, probing aphids, or both. Another feature that biochemically distinguishes the helpers of the two virus genera (Poty- and Caulimovirus) is the tendency of P2 to form huge paracrystalline polymers. On the other hand, HC-Pro forms a soluble dimer [88, 89] and is not known to polymerize into a highly ordered structure. It has been suggested that this property keeps P2 inactive in planta [11, 65]. The situation in the vector is unknown because direct data on P2 retention in aphids or its conformation are lacking. It is conceivable, though, that in the aphid's stylets the contact surface of CaMV particles with cuticular binding sites is expanded by a mesh of polymerized P2 molecules interfaced by a few P3 molecules to the virus coat. This expanded contact area would improve attachment of the virions during retention and facilitate their release during feeding of the aphid on another plant. This P2-P3 mesh might be important for aphid transmission. The icosahedral structure of CaMV offers only a few direct contact sites for vector interaction. Potyvirus morphology (long flexous fibrils), on the other hand, affords much more contact between virions and the aphid's cuticle. A problem related to the mode of action of CaMV ATFs is discerning how the MT-binding property of P2 fits into the bridge hypothesis. Although there is no definitive evidence demonstrating that the P2-MT interaction is actually part of the transmission process, such involvement can be postulated in three different ways [67]: (1) the P2-MT binding may be important, before virus acquisition by aphids, for a correct localization of P2 or the transmissible complex or both in
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infected plant cells (i.e., accumulation in ELib); (2) P2 may bind to a tubulin-like receptor site(s) on the cuticle of the aphid feeding apparatus; and (3) the aphid could inoculate P2 along with virus particles into the new host plant, where an association of P2 with the ph3^ocytoskeleton may be involved in very early stage infection, such as virion transport to the nucleus. In all cases, the putative participation of tubulin or MT or both in noncirculative transmission of plant viruses is a novel concept, which deserves further investigation.
VII. Regulation of Aphid Transmission Factor's Function(s) Aphid transmission of CaMV involves the formation of a transmissible P2-P3-virion complex able to interact with receptor site(s) in the aphid's feeding apparatus. The mechanisms responsible for the formation of this complex are not well known, nor is the molecular machinery controlling when and where the complex will be formed. Although direct experimental data are missing, we believe that one can intuitively perceive both the existence of and need for a tight regulation in this process. The most convincing elements favoring this point of view are developed below, and some of them have been discussed recently [11]. In addition to their role in transmission, P3, the capsid protein P4, and perhaps P2 play other important parts during the viral cycle. Although dispensable in most host species, P2 has been implicated in light sensitivity [90] and in forming a stable complex with cellular microtubules [67]. Clearly, P3 has additional functions required for the infectivity-viability of the virus [81] and possibly for compaction of viral DNA and in coat assembly [82]. The capsid protein may interact with PI for virus cell-to-cell movement [32] and with DNA and itself for encapsidation and binding to P6 [34]. The assembly of all three components into a transmissible complex obviously has to be tightly regulated to allow each component to fulfill its multiple functions in a coordinated fashion. Little is known about how this coordination is achieved, but several modes of regulation are possible. The genes of CaMV are differentially expressed, with gene I, V, and VI products being the early proteins, followed by P3 and finally P2 and P4 accumulation [32]. This timely coordinated pattern of protein expression may be controlled by various mechanisms operating mainly posttranscriptionally. The expression of P2 was found to follow different accumulation kinetics than that of PI (see above). This fact makes it unlikely that the two genes are translated by the exact same mechanism from the fiill length 35S RNA, that is, by a TAV-assisted relay race mechanism [27]. Differential gene expression could result from RNA splicing, and indeed, several spliced viral RNAs (up to 70% of the total viral RNA) have been detected in CaMV-infected plants. The splice acceptor site used to produce these subgenomic RNAs is located within gene II. Most relevant for our discussion is the conclusion of Kiss-Lazslo et ah [20] that the splicing events could con-
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trol the expression of several genes, including genes II, III, and IV This could ensure a controlled formation of virus particles and ATF(s) complex by providing the needed components at the appropriate time, thus preventing unwanted interaction of its (and other) constituents during unfavorable phases of infection. Not yet resolved is the problem of the presence of two types of virus proteincontaining inclusion bodies in infected plant cells. Do they represent different functions or different stages of maturation? Whatever the case, it is possible that compartmentalization of various viral proteins helps to regulate their functions. In particular, a spatial separation of the transmissible complex components could prevent their premature association. This view could also apply to the biologically inactive P2 paracrystals purported to "protect the virus against properties of P2 that may be necessary for aphid transmission but detrimental for the virus cycle" (i.e., MT-P2 interaction) [11]. Whether there is a simple equilibrium between soluble and paracrystalline P2 or whether it is converted to its active form by a biological mechanism or modification remains to be resolved. It is conceivable, however, that a drastic impact such as puncture of a cell by an aphid's stylets might be accompanied by major physical or chemical changes promoting solubilization of P2 in the cell. Another interesting possibility that might be important for regulation of the CaMV-transmissible complex is direct protein-protein interactions. Both P2 and P3 contain coiled coils that are in general known to form homologous and heterologous protein contacts. It is interesting to note that the P3 a-helical domain is able to interact not only with another P3 molecule but also with the a-helical domains of P2 [82, 64]. Vice versa, the a-helical domains of P2 have been shown to interact with P3, and our own preliminary results indicate that they also undergo self-association. How far coiled coil formation influences P3-P3, P3-P2, and P2-P2 interactions is not known, but many scenarios are imaginable that might modulate the activities of one, both, or even more involved protein partners.
VIIL Concluding Remarks The overall data summarized in this review illustrate the level of complexity that can underlie an apparently simple phenomenon such as noncirculative vector transmission of plant viruses. Over the past 50 years, our understanding of this type of relationship between virus and vector has increased dramatically, and caulimoviruses probably provide one of the best examples of our gradually evolving conceptualization of virus-vector-plant transmission cycles. The initial view of vector transmission as a passive nonspecific contamination of the vector's stylet tip followed by a simple mechanical inoculation process, has now evolved to a realization that vector transmission is a complex and crucial function in the infection cycle of viruses. Indeed, in the case of caulimoviruses, at least three genes (II, III, and IV) out of a total of six are involved, one way or another, in vec-
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tor transmission, with one (gene II) being totally devoted to it. The frequent multi functionality of plant virus proteins (HC-Pro and CP in potyviruses as well as CP and P3 in caulimoviruses) implies that tight regulatory mechanisms are likely to control the input of each gene in the vector transmission process, in contrast to other functions in which they also intervene. This particular aspect will certainly be one of the main focuses of future research on aphid transmission of caulimoviruses. Not only for caulimoviruses but also for noncirculative viruses generally, another aspect of transmission requiring research is the vector side of the process. The vector components recognized by viruses and presumably located at the surface of the cuticular lining of the mouthparts are totally hypothetical. Although the attachment of potyvirus HC-Pro has been demonstrated at these vector sites, even the chemical nature of the putative receptor is unknown. Among a number of important questions related to this hypothetical binding site are: Is it responsible for at least some of the vector specificity associated with plant virus transmission? Is the same family of receptor molecules used by different noncirculative viruses? The answers to these questions are critical, because this molecule would be an ideal target in nonconventional strategies for virus disease control. Beyond the complexity of virus-vector interaction, the case of caulimoviruses is of further interest because it demonstrates that more than one molecule can typify a helper strategy. As we indicated when describing the CaMV ATF's mode of action, the involvement of two transmission factors in the caulimovirus-vector relationship is not incompatible with the bridge hypothesis originally developed for potyviruses but rather is an extension of it. Whether we consider the former or the latter virus group, there is (or are) still one (or two) nonstructural virusencoded factor(s) mediating an indirect attachment of virus to the vector receptor. This raises a question concerning the actual definition of what should or should not be designated as the helper in a helper strategy? Indeed, although the answer remains clear for potyviruses where there is only one candidate, HC-Pro, the situation is less clear for caulimoviruses. Although P2 was once believed to be the only CaMV helper, there are now three possibilities: (1) P2 is indeed the only helper and P3 would be more correctly referred as a capsid component; (2) P2 and P3 can both be equally considered helpers; and (3) neither P2 nor P3 but rather the P2-P3 complex is the CaMV helper. Deciding which of these three propositions is correct awaits both a clear definition of helpers and further information about the mode of action of P3 (and P2), especially data on the pattern and kinetics of the formation of the CaMV transmissible complex, P2-P3-virus.
Acknowledgment We gratefully acknowledge the assistance of Roger Hull and Thomas Peter Pirone for critically reading our manuscript and making valuable suggestions for improvement.
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41. Kohli, A., Griffiths, S., Palacios, N., Twyman, R.M., Vain, P., Laurie, D.A., and Christou, P. (1999). Molecular characterization of transforming plasmid rearrangements in transgenic rice reveals a recombination hotspot in the CaMV 35S promoter and confirms the predominance of microhomology mediated recombination. Plant J. 17, 591-601. 42. Matthews, R.E.F. (1991). "Plant Virology," 3rd ed., Academic Press, New York. 43. Van Hoof, H.A. (1954). Verschillen in de overdracht van het bloemkoolmosaickvirus bij Myzus persicae Sulzer en Brevicoryne brassicae L. Tijdsch Plantenziekten 60, 267-272. 44. Chalfant, R.B., and Chapman, R.K. (1962). Transmission of cabbage viruses A and B by the cabbage aphid and the green peach aphid. J. Econ. Entomol. 55, 584-590. 45. Lim, W.L., and Hagedom, D.J. (1977). Bimodal transmission of plant virus. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosh, eds.), pp. 237-251. Academic Press, New York. 46. Harris, K.F. (1983). Sternorrhynchous vectors of plant viruses; virus-vector interactions and transmission mechanisms. ^
59. Harker, C.L., Woolston, C.J., Markham, PG., and Maule, A.J. (1987). Cauliflower mosaic virus aphid transmission factor protein is expressed in cells infected with some aphid non-transmissible isolates. Virology 160, 252-254. 60. Rodriguez, D., Lopez-Abella, D., and Diaz-Ruiz, J.R. (1987). Viroplasms of an aphid-transmissible isolate of cauliflower mosaic virus contain helper component activity. J. Gen. Virol. 68, 2063-2067. 61. Espinoza, A.M., Markham, P.G., Maule, A.J., and Hull, R. (1988). In vitro biological activity associated with the aphid transmission factor of cauliflower mosaic virus. J. Gen. Virol. 68, 1819-1830.
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CHAPTER 9
Cucumoviruses KEITH L. PERRY
/. Introduction Within three decades of the first description of tobacco mosaic virus as a fiherable infectious agent, Cucumber mosaic virus (CMV) had been described [1] and was shown to be transmissible by an aphid vector [2]. Remarkably, this initial report described a number of the key features of CMV transmission in particular and nonpersistent transmission in general. These properties were as follows: (1) aphids can acquire and transmit the virus after very brief periods of feeding (".. .aphids can transmit the disease after feeding for 5 minutes"); (2) viruliferous aphids lose their ability to transmit shortly after feeding on a healthy plant (".. .the minute amount of virus thus carried is exhausted during the first feeding period"); and (3) viruliferous aphids confined in a test tube lose the ability to transmit virus [2]. The concepts were presented in abstract form, and it was not until a number of years later that experimental evidence for CMV transmission by aphids was first published [3, 4]. This work showed that aphids feeding on plants infected with both CMV and tobacco mosaic virus only transmitted one of the viruses, CMV This led to the recognition that "the relation between aphid and virus may not be purely mechanical" [3]. One of the primary goals in contemporary research on vector transmission is to understand, at the molecular level, how viruses are acquired and transmitted. This chapter summarizes the seven decades of work on CMV transmission by aphid vectors and provides a perspective on the direction of current research.
Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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Viral Genome and Cucumovirus Transmission
Cucumber mosaic virus is a positive-strand RNA virus with three genomic RNAs separately encapsidated into icosahedral particles [5]: RNAs 1 and 2 each encode one protein involved in viral replication [6], and RNA 3 encodes two proteins, the coat protein (CP) and a protein associated with cell-to-cell movement [7]. There is a fifth viral protein encoded by RNA 2, which also affects virus movement [8]. Determinants for aphid transmission have all mapped to the CP gene on RNA 3 [9-12]. There are multiple strains of the virus whose complete genomes have been sequenced, although most of the available sequences are for RNA 3 or the CP gene alone. An additional satellite RNA is sometimes present and is encapsidated in virions [7]. There are two major subgroups of the virus, which can be distinguished by both serological methods and nucleic acid analyses [7]. Recently, a further division of subgroup I strains into lA and IB has been made based on analyses of RNA 3 sequences from 53 strains [13]. There are no reports of differences in transmission that correlate with CMV subgrouping, ahhough ecological differences correlating with subgroup have been described [14, 15]. Cucumber mosaic virus is a member of the family Bromoviridae and the genus Cucumovirus [16]. There are two other cucumoviruses, peanut stunt virus (PSV) and tomato aspermy virus (TAV). All three viruses share the same genomic organization, and the genome of at least one strain of each has been sequenced. The RNA 3s of these viruses are genetically interchangeable and can be substituted to create reassortants [7]. This was the technique originally used to demonstrate that aphid transmissibility maps to RNA 3 [17]. RNAs 1 and 2 can be exchanged between strains of CMV but not between viruses. The best studied of these viruses is CMV, and this is especially true with respect to transmission. In fact, although PSV can be transmitted by aphids, there are few reports [18, 19], and aphid transmission is often inefficient and erratic (Perry, unpublished data). There are greater numbers of reports on TAV transmission by aphids, some of which indicate vector specificity [10, 20-24]. The prevalence of both TAV and PSV may be due to spread by means other than aphids, such as vegetative propagation in the case of TAV and mowing of cover crops in the case of PSV All three viruses have been reported to be seedborne in at least one host [5, 25, 26]. Of significance to a genetic analysis of vector transmission has been the availability of complementary DNA copies of all three CMV genomic RNAs [7]. These cDNAs can be transcribed into infectious RNAs and mechanically inoculated onto plants to give rise to a systemic infection [27]. This reverse genetic system has allowed in vitro mutation and manipulation of the cDNAs to genetically map viral phenotypes such as aphid transmission [7, 11, 12].
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Vector Transmission of Cucumber Mosaic Virus
Cucumber mosaic virus is transmitted by aphids with extraordinary efficiency. Up to 50% or more of aphids transferred from infected to healthy plants can transmit the virus [12, 28]. In the field, the aphid-mediated spread of CMV is sufficient to devastate entire crops. Cucumber mosaic virus can be acquired and transmitted to a new host within seconds to minutes. The mode by which the aphid transmission of CMV occurs is termed nonpersistent because the virus does not persist in the vector [29]. This type of transmission is classified as noncirculative to differentiate it from the circulative viruses, which enter the food canal and digestive system, cross membrane barriers, circulate, and exit via the salivary glands [30, 31]. There are five additional genera of nonpersistently transmitted viruses, namely, Potyvirus, Alfamovirus, Fabavirus, Carlavirus, and Caulimovirus, although caulimoviruses are further described as semipersistently transmitted [32]. Of these, the transmission of the potyviruses has been the best studied (see chapter 10). There are three hallmarks of nonpersistent transmission as exemplified by CMV: (1) rapid virus acquisition from infected plants and transmission to healthy plants (seconds to minutes); (2) a short retention period of the virus within the aphid (minutes to hours); and (3) an increased rate of transmission following preacquisition fasting of aphids [32]. Nonpersistent transmission is experimentally performed by placing a starved aphid onto the leaf of a virus-infected plant. The aphid begins to feed and penetrates the plant cell(s) with its needlelike mouthparts. This feeding is naturally terminated or artificially interrupted by the investigator, who picks the aphid up with a small brush and transfers it onto a leaf of an uninfected plant. The transferred aphid begins to feed and in doing so introduces the virus to a new site of infection. An alternative method involves allowing aphids to crawl from infected plants onto closely placed uninfected plants. In any event, test plants are subsequently treated with an insecticide, and aphid transmission of virus is scored by observing plants for the appearance of disease symptoms after 5 to 21 days, depending on the host and environment. Strains of CMV may vary in the efficiency with which they are transmitted by aphids [33]. Not all aphids are the same, in that different species may or may not transmit CMV and vector species transmit virus with varying efficiencies [34-37]. In fact, biological clones of the same aphid species may differ in their abilities to transmit CMV [38]. Aphis gossypii and Myzus persicae are the vectors used most commonly in studies of nonpersistently transmitted viruses, but a wide variety of aphids have the ability to transmit CMV [36]. In laboratory assays, A. gossypii appears to be the more efficient of the two vectors of CMV [12, 39]. A number of factors have been described as influencing the efficiency of transmission: temperature, the host plant species used as the infected source of inoculum, the age and the number of days postinoculation of the virus-source plant when used; and the con-
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centration of CMV in source leaves [40-43]. The presence of satellite RNA can negatively affect the transmission rate, and this may be a function of a satellite-mediated reduction in the concentration of virus in source leaves [44-^6]. Virions of CMV and other cucumoviruses can be transmitted by aphids without the need for an additional accessory protein [9,10,47]. Aphids fed through membranes on solutions of purified virions are able to acquire and transmit CMV This is in contrast with potyvirus transmission, which requires the acquisition by aphids of a virally encoded helper protein [48]. The discovery of this requirement followed the observation that purified potyviruses acquired by aphids through membranes could be transmitted only if the aphids had prior or simultaneous access to a virus-free supernatant prepared from infected plants. The active principle of the supernatant was termed the helper component (HC), and it was hypothesized that HC functions by acting as a "bridge" to connect virus particles and the vector [49]. The binding of potyviruses in the mouthparts of the vector has been visualized by transmission electron microscopy [50]. More recently, compelling evidence for the role of HC in transmission has been provided by Pirone's group [51, 52]. They have shown that in cases of vector-specific transmission, specificity was conferred by the HC and that transmissibility was highly correlated with retention of virus in the stylets [51]. The N-terminal domain of the HC is important for the binding of HC in the vector [52]. Even though CMV does not appear to require an HC for its transmission by aphids, it is believed that the underlying mechanisms of its transmission are very similar to those of potyviruses (discussed in section VA).
IV. structure of Cucumber Mosaic Virus It is useful to consider the structure of CMV in relation to the closely related bromovirus cowpea chlorotic mottle virus (CCMV), especially since there is a highresolution structure available for the latter [53]. Both these groups are in the family Bromoviridae, although the bromoviruses are transmitted by beetles, not by aphids [16]. Virions of CMV and CCMV are spherical with icosahedral (T=3) symmetry and contain 180 protein subunits arranged as clusters of pentamers and hexamers [5, 54]. These two viruses differ with respect to their stability and assembly in vitro. Virions of CCMV are relatively resistant to ribonuclease and can be assembled in vitro to form empty capsids [55]. Cucumber mosaic virus is sensitive to ribonuclease [56], and although particles can be assembled in vitro, this assembly is dependent on the presence of RNA [57]. Protein-protein interactions are important for the stability of both viruses, but for CMV, RNA-protein interactions are paramount. Structural studies of CMV have shown that it bears a remarkable similarity to CCMV [54]. This is illustrated by the cryo-electron microscopy images shown in
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Fig. 1 Cryo-electron microscopic reconstruction of particles of cowpea chlorotic mottle virus (CCMV) and cucumber mosaic virus (CMV) at 23 A resolution. Both viruses are in the family Bromoviridae and are structurally related, with similar arrangements of pentamers and hexamers. The viruses differ in that CCMV and other members of the genus Bromovirus are transmitted by beetles, whereas CMV and other members of the genus Cucumovirus are transmitted by aphids. (The reconstructions were kindly provided by N. Olson, T. Baker and J. Johnson and are described by Wikofif et al. [54]).
Figure 1. The CMV virion is slightly larger than the CCMV and has been characterized as being in a permanently swollen state, a reference to the ability of CCMV to undergo structural transitions and "swell" [55]. For both these viruses, the size and orientation of the P barrels and the stabilizing interactions for hexamer formation appear to be conserved [54]. In developing mechanistic models underlying the process of CMV transmission, a knowledge of the structure and surface architecture of the virus would be extraordinarily useful. Of particular significance in this regard is the fact that the primary amino acid sequences of cucumoviruses and bromoviruses can be aligned [54]. This allows one to predict, based on the CCMV structure, regions of the CMV CP that are exposed on the surface of virions and available for interactions with the aphid vector. A model for the folded CP of CMV is shown in Figure 2. Based on the alignment of the viral CP sequences and independent secondary structure prediction analyses, one can be fairly confident about the overall fold of the CMV CP and about the amino acid sequences of specific p strands and intervening loops. Using this model, one can predict the location of amino acid positions 129 and 162, which have been shown to influence transmission (discussed in section VB.) Position 129 is located in the EF loop and is predicted to be on the surface of virions, whereas 162 is clearly buried in the fold structure [54]. Recently, a 3.1 A resolution structure for CMV has been obtained by T Smith at Purdue University working in collaboration with this laboratory (unpublished results). The structure is consistent with earlier interpretations of surface domains of the virus, although there are unique internal structural motifs not found in CCMV
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fiH-ai loop (190)-KDDALETDE-(198)
Amino acid position 129
Amino acid position 162
COOH Fig. 2 A ribbon model of a cucumber mosaic virus (CMV) coat protein subunit showing the pH-(3l loop sequence and the relative positions of sites important in aphid transmission. The orientation is such that the loops at the top of the figure are proximal to the surface of the virus. The amino and carboxytermini are internal to the virus particle and play a role in subunit-subunit and subunit-RNA interactions. Each of the eight p strands in the folded polypeptide is labeled. Alterations in the pH-pI loop dramatically affect aphid transmission, and the amino acid sequence for CMV strain Fny is indicated at the top of the figure. The coat protein consists of 218 amino acids, and the pH-pI loop sequence spans amino acid positions 190 to 198 (indicated in parentheses). Of the nine amino acids in the pH-pi loop, the seven shown in bold are invariant in all CMV coat protein genes that have been sequenced. The aspartic acid (D) at position 192 is accessible to chemical labeling. Among the spontaneous CMV mutants defective in aphid transmission, coat protein amino acid positions 129 and 162 are determinative and predicted to reside in the pE-pF and pG-PH loops, respectively. The amino acid at position 162 is internal in the virus structure, whereas that at position 129 may be exposed on the virion surface. The ribbon model is based on the X-ray crystallographic structure of cowpea chlorotic mottle virus by Speir et al. [53] and the analysis of Wikoflfe^ al [54].
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V. Mechanisms of Aphid Transmission of Cucumoviruses A.
Working Model for Nonpersistent Transmission
A working model for nonpersistent transmission of plant viruses by aphids can be envisioned as follows: (1) virions are taken up and bind in the maxillary food canal or elsewhere in the distal portion of the feeding apparatus; (2) virions are retained in an infectious state for the time it takes a vector to migrate onto an uninfected plant; and (3) virions are released from adsorbed sites during feeding and delivered to the site of infection. These steps are useful to consider when trying to understand how the surface architecture of virions physically interfaces with the vector or to explain the basis for defects in transmission observed in virus mutants. A large body of experimental evidence supports this model and has been reviewed [32, 48]. The model follows from the ingestion-egestion model of Harris [58], which suggests that nonpersistent virus transmission results from the uptake of plant sap into the food canal of the aphid and its subsequent egestion into an uninfected plant. The ingestion-egestion model is supported by observations that aphids can take up substantial volumes of virus-laden fluid and subsequently release virions during the process of feeding [59, 60]. An important addition to the model comes from recent work of Fereres' group [61, 62]. Using electronic monitoring techniques, they observed that successful transmission events correlated with salivation and cell membrane puncture; the patterns were very similar for both CMV and a potyvirus. Since the salivary and food canals within the maxillary stylets meet at the tip of the stylet [63, 64], they hypothesized that the virions bound at sites near the stylet tip are those that are primarily responsible for transmission and that their release is mediated or enhanced by salivation. The ingestion-salivation and ingestion-egestion models of nonpersistent transmission have been discussed in detail in chapters 5 and 6, respectively. Several lines of evidence support a specific binding event during nonpersistent transmission. Aphids given a single access to a CMV-infected plant can transmit virus multiple times when successively transferred for brief (10-minute) periods to healthy plants [39]. When two different viruses are acquired by a single aphid from a doubly infected plant and the insect is successively transferred to uninfected plants, multiple transmissions either of one or of both viruses can occur. The latter suggests that the two viruses are retained at independent but not necessarily fundamentally different sites [39]. A discriminating binding interaction is consistent with the observation that small changes in CP sequence can alter transmission (discussed in section VB). In developing a more detailed mechanistic model, the availability of mutants or variants that are defective or very inefficient in transmission by the vector has been invaluable. The original mapping of the aphid transmission phenotype to the
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CP of CMV was made possible by such mutants [9, 10, 17]. Two classes of CP mutants have proved useful in this regard. B.
Spontaneous Mutants Defective in Aphid Transmission
The transmission phenotype of CMV appears to be quite stable, as spontaneous CMV mutants defective in transmission are relatively rare [65]. There are only four described mutants, CMV-M [17], CMV-2Al-MT-60x [65], CMV-C [66], and the Badami mutant [35]. The nature of the defect has been thoroughly characterized at the molecular level for only two of these strains. The CMV-M mutant was shown to differ from an efficiently transmitted wild type strain at eight CP amino acid positions [67, 68]. Of these, positions 129 and 162 were important in determining transmissibility by the aphid A. gossypii [11]. Interestingly, a restoration of CMV-M transmission by M persicae required additional amino acid modifications. Thus, molecular determinants for differential transmission by two species of aphids were established [12]. The CP of the second fully characterized mutant, CMV-2Al-MT-60x, differs from its parental virus at only a single amino acid position. Remarkably, this change was the same alanine-to-threonine change at position 162 as was observed in CMV-M, even though these mutants were isolated decades apart [69]. Virions of the two mutants share the additional feature of instability under the standard conditions of CMV purification, that is, in the presence of chloroform and 1.5 M sodium [17, 65]. It is hypothesized that the defect may be due in part to a lowered stability of virions and that the requirements for stability within the mouthparts of different aphid species may vary. Two lines of evidence are consistent with this hypothesis. The first is the results of experiments in which virions of the mutants and wild-type strains were subjected to an in vitro disruption assay. The wild-type strains are stable when incubated in 4 M urea, whereas the mutants CMV-M and CMV-2Al-MT-60x are completely disrupted in 1 M and 2 M urea, respectively [69]. Second, structural analyses reveal that the amino acid at position 162 is buried within the folded CP [54] and must exert its effect indirectly at the virion surface or through its influence on stability. There is a paradox in the isolation of this class of mutant. The parental CMV isolates were passaged plantto-plant mechanically, removing the selective pressure for vector transmissibility. The mutants were presumably isolated because of their competitive ability to either replicate or move systemically within the host. Thus, a relative instability appears to have conferred a competitive advantage within the plant, perhaps affecting coating-uncoating steps in the viral life cycle. C.
Engineered Mutants Defective in Aphid Transmission
Using the model for the folded CMV CP as a starting point, published CP gene sequences can be analyzed and the extent of amino acid conservation in different
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structural domains determined. As would be expected, sequences in the p strands are relatively conserved [70]. The converse, that intervening loop structures tend to be less conserved is also true, with one notable exception. Amino acids in the exposed pH-pi loop are remarkably well conserved; seven of the nine amino acids are invariant, not only in CMV, but among the other cucumoviruses as well (Fig. 2). This pH-pI loop is positioned on the surface of virions, and its exposure to solution has been confirmed by chemical labeling. The aspartic acid at position 192 (Fig. 2) was replaced with a cysteine residue, which was then labeled with a thiol-reactive reagent [70]. The pH-pI loop was also shown to be highly antigenic [70, 71]. Since six of the nine amino acids in the pH-pI loop are charged, this region is likely to affect electrostatic interactions among CP subunits and potentially between virus and vector surfaces. It is hypothesized that structural domains on the surface of virions will mediate interactions in the aphid vector, and that a conservation of surface amino acid residues is due, at least in part, to their functional role in virus transmission. This has been shown to be the case with the potyviruses, as a conserved DAG motif in the N-terminal region of the CP was demonstrated to be essential for vector transmissibility [72, 73]. Additionally, there are two conserved sequences in the helper protein (HC) required for transmissibility, the PTK and KITC motifs, further substantiating the role that a linear amino acid sequence can play in transmission [51, 52, 74]. Although sequences may be invariant in natural populations under selective pressure to maintain vector transmissibility, transmission-defective variants could arise or be engineered that are still competent in replication and systemic movement. This has been shown to be the case in a mutational analysis of the PHpl loop of the CMV CP Radical amino acid substitutions were made in five of the conserved positions: these changes were to alanine or to an amino acid of opposite charge. Changes at nearly all of these positions dramatically reduced or eliminated vector transmissibility while not affecting the accumulation or systemic movement of mutant virus in tobacco (Perry, unpublished data). Especially intriguing is the fact that spontaneous, intramolecular, second-site revertants arose, which were fully aphid-transmissible. The CP amino acid changes that restored aphid transmissibility were within or adjacent to the pH-pi loop or in adjoined surface loop structures (Perry, unpublished data). These results are consistent with the requirement of a surface structure or charge density that is essential for aphid transmission.
W. Concluding Remarks Vector transmission provides a strong selection pressure that will influence the evolution of viral CP, virion structure, and in some cases nonstructural proteins [48]. In the case of CMV, it has been shown that the insect vector can mediate the
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establishment of reassortants with mixed populations of RNAs [24]. Mixed populations of CMV have been observed in the field [75], as have naturally occurring reassortants [76]. Thus, CMV can be seen as a virus whose coevolution with aphid vectors has determined both its genomic and physical properties. Given the high degree of variability among RNA plant viruses and the error-prone nature of RNA replication [77], it is surprising how stable the aphid transmission phenotype is when virus is passaged in the absence of its natural vector [65]. This can be interpreted as resulting from the absence of a helper component and consequentially a greater selection pressure on the CP for maintenance of its frinctional roles in encapsidation, movement, and transmission by the vector. A more provocative interpretation is that there are essential virion-host plant factor interactions that mimic the interactions of the virus CP with components in the aphid vector. In this case, there would be a strong selection pressure for CP sequences or structures required for aphid transmission even when virus is mechanically passaged in the absence of the vector. An understanding of these and other issues will be furthered by ongoing structural analyses of the virus. It is hoped that the opportunity to directly visualize the static structure of CMV will shed light on more dynamic properties, such as its ability to bind in and be transmitted by an aphid vector. Authors note. After the preparation of this chapter, a 3.2 angstrom atomic structure for CMV was determined and reported (Smith, T.J., Chase, E., Schmidt, T, and Perry, K.L. 2000, The structure of cucumber mosaic virus and comparison to cowpea chlorotic mottle virus. J. VirolJ4, 7578-7586). The results of this work impact our interpretation of the transmission defective mutants and the model for the coat protein presented in Figure 2.
References Johnson, J. (1927). The classification of plant viruses. Wisconsin Agricultural Experiment Station Research Bulletin No. 76. Doolittle, S.P., and Walker, M.N. (1928). Aphis transmission of cucumber mosaic. Phytopathology 18, 143. Hoggan, LA. (1929). The peach aphid (Myzus persicae Sulz.) as an agent in virus transmission. Phytopathology 19, 109-123. Hoggan, LA. (1931). Further studies on aphid transmission of plant viruses. Phytopathology 21, 199-212. Francki, R.LB., Mossop, D.W., and Hatta, T. (1979). Cucumber mosaic virus. In "CMI/AAB Descriptions of Plant Viruses" (B.D. Harrison and A.F. Murant, eds.), No. 213. Commonwealth Agricultural Bureau/Association of Applied Biologists, Kew, England. Hayes, R.X, and Buck, K.W. (1990). Complete replication of a eukaryotic virus RNA in vitro by a purified RNA-dependent RNA polymerase. Cell 63, 363-368.
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7. Palukaitis, P., Roossinck, M.J., Dietzgen, R.G., and Francki, R.I.B. (1992). Cucumber mosaic virus. ^Jv. Virus Res. 41,281-348. 8. Ding, S., Li, W., and Symons, R.H. (1995). A novel naturally occurring hybrid gene encoded by a plant RNA virus facilitates long distance virus movement. EMBOJ. 14, 5762-5772. 9. Gera, A., Loebenstein, G., and Raccah, B. (1979). Protein coats of two strains of cucumber mosaic virus affect transmission by Aphis gossypii. Phytopathology 69, 396-399. 10. Chen, B., and Francki, R.I.B. (1990). Cucumovirus transmission by the aphid Myzus persicae is determined solely by the viral coat protein. J. Gen. Virol. 71, 939-944. 11. Perry, K.L., Zhang, L., Shintaku, M.H., and Palukaitis, P. (1994). Mapping determinants in cucumber mosaic virus for transmission by Aphis gossypii. Virology 205, 591-595. 12. Perry, K.L., Zhang, L., and Palukaitis, P. (1998). Amino acid changes in the coat protein of cucumber mosaic virus differentially affect transmission by the aphids Myzus persicae and Aphis gossypii. Virology 242, 204-210. 13. Roossinck, M.X, Zhang, L., and Hellwald, K. (1999). Rearrangements in the 5' nontranslated region and phylogenetic analyses of cucumber mosaic virus RNA 3 indicate radial evolution of three subgroups. J. Virol. 73, 6752-6758. 14. Quiot, J.B., Devergne, J.C, Cardin, L., Verbrugghe, M., Marchoux, G., and Labonne, G. (1979). Ecologie et epidemiologic du virus de la mosaique du concombre dans le sud-est de la France. VII. Repartition de deux types de populations virales dans des cultures sensibles. Ann. Phytopathol. 11, 359-373. 15. Haack, I. (1986). A contribution to the ecology of the cucumber mosaic virus. Acta Phytopathol. Entomol Hung 21,279-285. 16. Van Regenmortel, M.H.V (1999). "Virus Taxonomy: Seventh Report of the International Committee on the Taxonomy of Viruses." Academic Press, San Diego. 17. Mossop, D.W., and Francki, R.I.B. (1977). Association of RNA 3 with aphid transmission of cucumber mosaic virus. Virology M, 177-181. 18. Hebert, T.T. (1967). Epidemiology of the peanut stunt virus in North Carolina. Phytopathology 57,461. 19. Tolin, S., and Miller, ID. (1975). Peanut stunt virus in crownvetch. Phytopathology 65, 321-324. 20. Blencowe, J.W., and Caldwell, J. (1949). Aspermy—a new virus disease of the tomato. Ann. Appl. Biol. 36, 320-326. 21. Noordam, D. (1952). Virusziekten bij Chrysanten in Nederland. Tijdsch. Plantenziekten 58, 121-189. 22. Brierly, P., Smith, F.F., and Doolittle, S.P. (1955). Some hosts and vectors of tomato aspermy virus. Plant Dis. Rep. 39, 152-156. 23. HoUings, M. (1955). Investigation of chrysanthemum viruses I. Aspermy flower distortion. Ann. Appl. Biol. 43, 86-103. 24. Perry, K.L., and Francki, R.I.B. (1992). Insect-mediated transmission of mixed and reassorted cucumovirus genomic RNAs. J. Gen. Virol. 73, 2105-2114. 25. Hollings, M., and Stone, O.M. (1971). Tomato aspermy virus. In "CMI/AAB Descriptions of plant viruses" (A.J. Gibbs, B.D. Harrison, and A.F. Murant, eds.), No.79. Commonwealth Agricultural Bureau/Association of Applied Biologists, Kew, England. 26. Mink, G.I. (1972). Peanut stunt virus. In "CMI/AAB Descriptions of Plant Viruses" (A.J. Gibbs, B.D. Harrison, and A.F. Murant, eds.). No. 92, Commonwealth Agricultural Bureau/Association of Applied Biologists, Kew, England.
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27. Rizzo, T.M., and Palukaitis, P.(1990). Construction of full-length cDNA clones of cucumber mosaic virus RNAs 1, 2, and 3: Generation of infectious RNA transcripts. Mol. Gen. Genet. 222, 249-256. 28. Banik, M.T., and Zitter, T.A. (1990). Determination of cucumber mosaic virus titer in muskmelon by enzyme-linked immunosorbent assay and correlation with aphid transmission. Plant Dis. 74, 857-859. 29. Watson, M.A., and Roberts, KM. (1939). A comparative study of the transmission of Hyoscyamus virus 3, potato virus Y and cucumber virus 1 by the vectors Myzus persicae (Sulz), M. circumflexus (Buckton), and Macrosiphum get (Koch). Proc. R. Soc. Lond. [Biol] 111, 543-576. 30. Harris, K.F. (1990). Aphid Transmission of Plant Viruses. In "Plant Viruses" (C.L. Mandahar, ed.). Vol. 2, pp. 177-204. CRC Press, Boca Raton, FL. 31. Gray, S.M. (1996). Plant virus proteins involved in natural vector transmission. Trends Microbiol. 4, 259-264. 32. Pirone, T.P., and Perry, K.L. (2001). Aphids—nonpersistent transmission. Adv. Bot. Res. (hi press). 33. Simons, J.N. (1957). Three strains of cucumber mosaic virus affecting bell pepper in the everglades area of south Florida. Phytopathology 47, 145-150. 34. Bhargava, K.S. (1951). Some properties of four strains of cucumber mosaic virus. Ann. Appl. Biol. 38, 377-388. 35. Badami, R.S. (1958). Changes in the transmissibility by aphids of a strain of cucumber mosaic virus. Ann. Appl. Biol. 46, 554-562. 36. Kennedy, J.S., Day, M.F., and Eastop, VF. (1962). "A Conspectus of Aphids as Vectors of Plant Viruses." Commonwealth Agricultural Bureau, London. 37. Conti, M., Caciagli, P., and Casetta, A. (1979). Infection sources and aphid vectors in relation to the spread of cucumber mosaic virus in pepper crops. Phytopathol. Mediterr 18, 123-128. 38. Simons, J.N. (1959). Variation in efficiency of aphid transmission of southern cucumber mosaic virus and potato virus Y in pepper. Virology 9, 612-623. 39. Fujisawa, L (1985). Aphid transmission of turnip mosaic virus and cucumber mosaic virus 2. Transmission from virus mixtures. Ann. Phytopathol. Soc. Jpn. 51, 562-568. 40. Simons, J.N. (1955). Some plant-vector-virus relationships of southern cucumber mosaic virus. Phytopathology 45, 217-219. 41. Simons, J.N. (1958). Titers of three nonpersistent aphid-borne viruses affecting pepper in south Florida. Phytopathology 48, 265-268. 42. Stimmann, M.W., and Swenson, K.G. (1967). Aphid transmission of cucumber mosaic virus affected by temperature and age of infection in diseased plants. Phytopathology 57, 1074-1076. 43. Yamamoto, T., and Masayoshi, I. (1983). Aphid transmission from cucumber cultivars infected with watermelon mosaic virus and cucumber mosaic virus. Ann. Phytopathol. Soc. Jpn. 49, 508-513. 44. Jacquemond, M. (1982). L'ARN satellite du virus de la mosaique du concombre IV-Transmission experimentale de la maladie necrotique de la tomate par pucerons. Agronomic 2, 641-646. 45. Sayama, H., Sato, T., Kominato, M., Natsuaki, T., and Kaper, J.M. (1993). Field testing of a satellite-containing attenuated strain of cucumber mosaic virus for tomato protection in Japan. Phytopathology 83, 405^10. 46. Escriu, F, Perry, K., and Garcia-Arenal, F (2000). Aphid transmissibility of cucumber mosaic virus by Aphis gossypii correlates with viral accumulation and is affected by the presence of satellite RNA. P/j^topa^Ao/ogy 90:1068-1072.. 47. Pirone, T.P., and Megahed, E. (1966). Aphid transmissibility of some purified viruses and viral RNA's. Virology 30, 631-637.
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48. Pirone, T.P., and Blanc, S. (1996). Helper-dependent vector transmission of plant viruses. Annu. Rev. Phytopathol 34, 227-247. 49. Govier, D.A., and Kassanis, B. (1974). A virus-induced component of plant sap needed when aphids acquire potato virus Y from purified preparations. Virology 61, A20-A26. 50. Ammar, E.D., Jarlfors, U., and Pirone, T.P. (1994). Association of potyvirus helper component protein with virions and the cuticle lining the maxillary food canal and foregut of an aphid vector. Phytopathology^^, 1054-1060. 51. Wang, R.Y., Powell, G., Hardie, J., and Pirone, T.P (1998). Role of the helper component in vector-specific transmission of potyviruses. ^ Gen. Virol. 79, 1519-1524. 52. Blanc, S., Ammar, E.D., Garcia-Lampasona, S., Dolja, VV, Llave, C , Baker, J., and Pirone, T.P. (1998). Mutations in the potyvirus helper component protein: Effects on interactions with virions and aphid stylets. J. Gen. Virol. 79, 3119-3122. 53. Speir, J.A., Munshi, S., Wang, G., Baker, T.S., and Johnson, XE. (1995). Structures of the native and swollen forms of cowpea chlorotic mottle virus determined by X-ray crystallography and cryo-electron microscopy. Structure 3, 63-78. 54. Wikofif, W.R., Tsai, C.J., Wang, G., Baker, T.S., and Johnson, J.E. (1997). The structure of cucumber mosaic virus—cryoelectron microscopy, x-ray crystallography and sequence analysis. Virology 232, 91-91. 55. Bancroft, J.B., Hills, G.J., and Markham, R. (1967). A study of the self-assembly process in a small spherical virus: Formation of organized structures from protein subunits in vitro. Virology 31, 354-379. 56. Francki, R.I.B. (1968). Inactivation of cucumber mosaic virus (Q strain) nucleoprotein by pancreatic ribonuclease. Virology 34, 694-700. 57. Kaper, J.M. (1969). Reversible dissociation of cucumber mosaic virus (strain S). Virology 37, 134-139. 58. Harris, K.F. (1977). An ingestion-egestion hypothesis of noncirculative virus transmission. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 165-219. Academic Press, New York. 59. Harris, K.F, and Bath, J.E. (1973). Regurgitation by Myzus persicae during membrane feeding: Its likely function in transmission of nonpersistent plant viruses. Ann. Entomol. Soc. Am. 66, 793-796. 60. Garrett, R.G. (1973). Non-persistent aphid-borne viruses. In "Viruses and Invertebrates" (A.J. Gibbs, ed.), pp. 476^92. North Holland, Amsterdam. 61. Collar, J.L., Avilla, C., and Fereres, A. (1997). New correlations between aphid stylet paths and nonpersistent virus transmission. Environ. Entomol. 26, 537-544. 62. Martin, B., Collar, J.L., Tjallingii, WF, and Fereres, A. (1997). Intracellular ingestion and salivation by aphids may cause the acquisition and inoculation of nonpersistently transmitted plant viruses. J. Gen. Virol. 78, 2701-2705. 63. Forbes, A.R. (1969). The stylets of the green peach aphid, Myzus persicae (Homoptera: Aphididae). Can. Entomol. 101, 3 1 ^ 1 . 64. Forbes, A.R. (1977). The mouthparts and feeding mechanism of aphids. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 83-103. Academic Press, New York. 65. Ng, J., and Perry, K.L. (1999). Stability of the aphid transmission phenot)^e in cucumber mosaic virus. Plant Pathol 48, 388-394. 66. Rodriguez, R. (1993). Cucumber mosaic virus: Aphid nontransmissible isolates generated by passages and regions of the coat protein involved in transmission. Ph.D. thesis, Cornell University, Ithaca, NY.
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67. Owen, J., Shintaku, M., Aeschleman, P., Tahar, S.B., and Palukaitis, P (1990). Nucleotide sequence and evolutionary relationships of cucumber mosaic virus (CMV) strains: CMV RNA 3. J. Gen. Virol. 71, 2243-2249. 68. Shintaku, M, (1991). Coat protein gene sequences of two cucumber mosaic virus strains reveal a single amino acid change correlating with chlorosis induction. J. Gen. Virol. 11, 2587-2589. 69. Ng, J., and Perry, K.L. (2000). Cucumber mosaic virus mutants with altered physical properties and defective in aphid transmission. Virology 276:395^03. 70. Perry, K.L., Reiley, B., and He, X. (2001). A highly antigenic coat protein domain on the surface of cucumber mosaic virus. (Manuscript in preparation) 71. He, X., Liu, S., and Perry, K.L. (1998). Identification of epitopes in cucumber mosaic virus using a phage-displayed random peptide library. J. Gen. Virol. 79, 3145-3153. 72. Harrison, B.D., and Robinson, D.J. (1988). Molecular variation in vector-borne plant viruses: Epidemiological significance. Philos. Trans. R. Soc. Lond. Biol. 321, 447^62. 73. Atreya, CD., Raccah, B., and Pirone, T.P (1990). A point mutation in the coat protein abolishes aphid transmission of a potyvirus. Virology 178, 161-165. 74. Huet, H., Gal-on, A., Meir, E., Lecoq, H., and Raccah, B. (1994). Mutations in the helper component protease gene of zucchini yellow mosaic virus affect its ability to mediate aphid transmissihi\i\y. J. Gen. Virol. 75, 1407-1414. 75. Perry, K.L., Habili, N., and Dietzgen, R.G. (1993). A varied population of cucumber mosaic virus from peppers. Plant Pathol. 42, 806-810. 76. White, P.S., Morales, E, and Roossinck, M.J. (1995). Interspecific reassortment of genomic segments in the evolution of cucumoviruses. Virology 207, 334-7. 77. Roossinck, M.J. (1996). Mechanisms of plant virus evolution. Annu. Rev. Phytopathol. 35, 191-209.
CHAPTER 10
Potyviruses BENJAMIN RACCAH HERVE HUET STEPHANE BLANC
/. Introduction Potyviruses comprise the largest group of plant viruses. They are transmitted mostly by aphids in a nonpersistent manner. Many plant viruses cause severe disease in the host they infect. Therefore, survival of plant viruses in nature strongly depends on their capacity to spread among new hosts. Most plant viruses require vectors for their transmission. This is attributed to the impermeable cuticle that coats the plant epidermis and cellulose cell walls, preventing entrance of virus particles, in contrast to animal viruses, which enter through natural openings. The association of nonpersistent viruses (NPV) with the vectors is very brief Therefore, they were believed to be carried passively and were named stylet-borne, for almost half a century [1]. The wide use of mechanical inoculation led to the appearance of virus strains that were highly infective but not transmissible by aphids [2]. Complementation between aphid-transmissible (AT) and non-aphidtransmissible (NAT) strains led to the understanding that the process of transmission of potyviruses is much more complex than simply being carried on the external ridges of stylets, as suggested by van Hoof [3]. Many comprehensive reviews were written on the mechanisms of transmission in general [4, 5] and recently also on viral genes involved in the NPV transmission [6, 7]. The present chapter is intended to complement the former reviews and summarize some important biological, biochemical, and molecular findings that may resolve the complexity of potyvirus transmission by aphids.
Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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//. A.
Biology of Potyvirus Transmission
Vectors
The majority of plant pathogen vectors and the most important are found in the class Insecta, particularly in the order Homoptera. Homopterans pierce plants to suck sap. Aphids, whiteflies, and leafhoppers comprise the major vector families. The success of these insects as vectors is attributed to their delicate penetration of the plant cell where virus is deposited in the cytoplasm. Important vectors are also found among other organisms (e.g., mites, nematodes, or fungi). Virus spread can take place also by contact or through seed or infected propagative material or both. Potyviruses are transmitted almost exclusively by aphids. More than 4,000 aphid species have been identified to date. Almost 300 species have been tested, and more than 200 species have been found to transmit at least one virus [8]. The majority of vector species are found in the subfamily Aphidinae, including the genera Myzus, Aphis, and Macrosiphum. Successful vector species for potyviruses are polyphagous, for example, Myzus persicae, Aphis gossypii, Aphis craccivora, dind Aphis fabae. Many factors determine the potential of an aphid species to become a successful vector: (1) alternation between wingless and winged forms; (2) alternation between short- and long-distance fliers; (3) migration from primary and secondary hosts (for the sexual and parthenogenetic morphs, respectively); and (4) host plant specificity. Feeding behavior is an important factor for understanding the role of aphids as vectors of nonpersistent viruses. In the process of host selection, they probe the host. Probing is accomplished by insertion of the stylet into the epidermal layer for several seconds. Such a probe is sufficient to sample the cell sap or contents. Muller [9] reported that more probes are made on hosts than on nonhosts. The process of virus release from the stylet is not yet understood. An ingestion-egestion hypothesis [10] suggests that the virus is inoculated during intracellular egestion on a healthy plant. Martin et al [11] propose an alternative mechanism based on the fact that the virus is released during the first phase of the intracellular stylet puncture. This would favor salivation over egestion as a mechanism for virus release in target hosts. Readers are referred to chapters 5 and 6 for further reading on the ingestion-egestion and ingestion-salivation hypotheses and feeding behavior of aphids [12, 13]. B.
Mode of Transmission
Watson and Roberts [14] used potyviruses and aphids as case studies to formulate the nonpersistent mode of virus transmission. The original definition of nonpersistent transmission included the following parameters: • Brief acquisition and transmission (seconds to minutes)
1.
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P O T Y VIRUSES
Table I
Genera of the Family Potyviridae. Number of viruses
Genus Potyvirus Rymovirus Ipomovirus Bymovirus
Definite
Tentative
118 5 1 6
69 2 1
Vectors Aphids Mites Whiteflies Fungi
Source: http://biology.anu.edu.au/research-groups/MES/vide/ descr652.htm
• Short retention of virus in or on the vector's feeding apparatus (less that 1 hour) • Lack of a measurable latent period in the vector • Loss of virus when the vector molts • Low viral specificity for aphid species • Increased transmissibility following preacquisition fasting Most parameters of NPVs indicate a temporary association between virus and aphid mouthparts. For this reason, the term stylet-borne has been preferred to describe a simple and rather mechanistic association [1]. C.
Viruses
The family Potyviridae includes four genera transmitted by different vectors (Table 1). The present chapter deals mainly with the genus Potyvirus, the most important and most researched of the genera. However, some remarks about other genera will be stressed when relevant. The genus Potyvirus includes many economically important viruses, such as the type member, potato virus Y (PVY), tobacco etch virus (TEV), zucchini yellow mosaic virus (ZYMV) in annuals, and plum pox virus (PPV) in perennials. A comprehensive description of potyvirus groups is given elsewhere [15]. In brief, potyviruses are filamentous, 720 to 770 nm long, and contain a linear, unipartite, ssRNA of about 10 kb. The positive-sense RNA is translated to produce a polyprotein, which is subsequently processed by three viral proteases to yield eight proteins and two peptides. The two viral proteins that are involved in transmission, helper component (HC) and coat protein (CP), are encoded by their respective genes in the viral genome (Fig. 1, top). A characteristic feature of potyvirus infection is the presence of distinctive cytoplasmic inclusions (pinwheels) in the host cells. In certain hosts, the virus induces nuclear inclusions as well as cytoplasmic or amorphous inclusions.
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;:;i(:i;;}-(AAAA)i,
Cysteine-rich region
I CO Ktroig Aphid transmission
\r^^tS^
^ dccvnn Aphid transmission Long distance movement
Fig. 1 A schematic representation of the potyviral genome and processing of the derived polyprotein (top). Sites of relevance to aphids transmission in the HC-Pro (bottom).
D.
Nonmolecular Approaches to the Study of Potyvirus Coat Protein and Helper Component
Many important biological studies have contributed to the understanding of potyvirus transmission by aphids. Some of these provided the basis for the recent molecular studies. 1.
AVAILABILITY OF NON-APHID-TRANSMISSIBLE VIRUS STRAINS
Partial or complete loss of aphid transmissibility has been recorded in many studies [2, 16-23]. Loss of aphid transmissibility has been attributed to continuous mechanical inoculation [2]. 2.
LACK OF APHID TRANSMISSION RESULTING FROM VIRION DEFICIENCY
For nonenveloped plant viruses, the CP encapsidates the nucleic acid and is the most exterior part of the virion. Therefore, the CP can come in immediate contact with vector surfaces and organs. Intuitively, differences in transmissibility have been attributed to changes in the CP. The first proof for the role of the CP in transmission came from transcapsidation studies among barley yellow dwarf luteovirus isolates and the effect of transcapsidation on transmission by aphids [24]. These experiments inspired in vitro transcapsidation experiments with the nonpersistent cucumber mosaic virus [25]. In both cases, transmissibility characteristics were altered when the nucleic acid of an AT strain was encapsidated with the CP of a NAT strain and vice versa. Loss of transmissibility due to a deficiency in the CP is well documented among potyviruses [26] and will be detailed further.
10.
POTYVIRUSES 3.
185
DISCOVERY OF THE HELPER PHENOMENON
Watson [27] was the first to report on the transmission of the non-aphid-transmissible potato virus C (known then as PVC and now designated PVY^) from a plant also infected with potato virus Y (PVY). Later, Kassanis [28] reported aphid transmission of another nontransmissible virus (potato aucuba mosaic virus (PaMV), a potexvirus that was then a potyvirus) from plants also infected with PVY This assisted transmission prompted work that led Kassanis and Govier [29] to the discovery of the helper phenomenon in aphid transmission. In these early experiments, they showed in vivo complementation between PVY and the nontransmissible PaMV and PVY^. Unlike luteoviruses, the helper virus could assist the non-aphid-transmissible (NAT) viruses even if it was previously acquired from a separate host [30]. Later, Govier and Kassanis also showed that PaMV and PVY^ could become transmissible by aphids that first acquired a PVY extract through membranes [31] and that the PVY virus particle itself could not account for the HC activity [32]. This led to the designation of the term helper component (HC) for the proteinaceous fraction assisting transmission. The sequence of acquisition (HC first, then virions, but not the reverse) was found essential for successful transmission, and Govier and Kassanis concluded that HC might act as a "bridge" between aphids and virus particles. This hypothesis has recently been designated the bridge hypothesis [7]. 4.
IMPORTANCE OF IN VITRO ACQUISITION FOR DETERMINING DEFICIENCIES IN COAT PROTEIN OR HELPER COMPONENT
Deficiencies in aphid transmissibility were also found to possibly be caused by a deficient HC. Hence, preparation of heterologous mixtures of HC and virions of aphid-transmissible strains with their reciprocal nontransmissible strains (e.g., HC from a nontransmissible strain mixed with virions from a transmissible strain and vice versa) helped elucidate the cause of transmission deficiency [21, 26, 33-35].
///. A.
Role of Coot Protein in Potyvirus Transmission
Structure of Potyvirus Coat Protein
As indicated above, in many potyviruses loss of transmissibility has been attributed to the coat protein [36]. The potyviral genome is coated with nearly 2,000 subunits of coat protein. Sequence comparison and particle assembly properties suggest the presence of three regions in the CP molecule: a surface-exposed N-terminus (varying in length and sequence), a highly conserved core with 215-227 amino acids, and a surface-exposed C-terminus with 18-20 amino acids [37, 38] (Fig. 2).
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Amino acid sequence 218 amino adds N-terminal 30 amino acids
w.
C-terminal 19 amino acids
Sub unit fokjinq pattern
-CID-C
C
RifM CED
^
P
(I
Assembled vrus
7 to 8 subunits form a disk
Fig. 2
B.
A schematic description of the potyviral coat protein subunit (From Shukla and Ward [38]).
Importance of N-Terminal DAG Motif
In order to interact either with the HC or with a putative receptor in the aphid stylets, the CP motif involved in vector transmission must be located at the surface of the virus particle. Although the structure and conformation of the CP subunits have not been totally elucidated, a number of studies have been devoted to determining the topography of various CP motifs in the mature virus particle. Both biochemical [37] and immunological [39] analyses first suggested that the N-terminal extremity of the CP is located on the external surface of the tobacco etch virus (TEV) particle. Shukla et al [40] confirmed these findings for several other potyviruses and demonstrated that a short C-terminal region was also surface-exposed (Fig. 1). At the time of these studies, more and more sequences of potyviruses were becoming available (Riechmann et al.
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[41]). The alignment of CP genes from different potyviruses consistently revealed that the N-terminus was variable, whereas the C-terminus and core regions were highly conserved. Harrison and Robinson [42] proposed a detailed analysis of this variability, comparing the N-terminal sequences of the CP of aphid-transmissible potyviruses with those of a non-aphid-transmissible TEV isolate that produced a functional HC. These authors pointed out an aspartic acid-alanine-glycine (DAG) motif that was highly conserved in all aphid-transmitted isolates but not in the nontransmissible TEV. The suggested involvement of this DAG motif in aphid transmission proved accurate when tested 2 years later (see section IIIC). Altered DAG sequences associated with nontransmissibility were consistently described for many other viruses, a change from DAG to DAL in PPV [43], DAG to DTG in papaya ringspot virus (PRSV) [44] and ZYMV [45], and DAG to DAE in tobacco vein mottling virus (TVMV) [46] and kalanchoe mosaic virus (KMV) [47]. Indirect proof for the involvement of CP termini (containing the DAG) in transmission was achieved by partial digestion with trypsin. Reduction or total loss of aphid transmission was recorded for ZYMV and maize dwarf mosaic virus (MDMV) [48] as well as for TEV and TVMV [B. Raccah and T.P Pirone, unpublished results] when virions were subjected to proteolysis. However, the first direct evidence of the importance of the DAG motif in aphid transmission came from the availability of full-genome length infectious clones and mutagenesis experiments. An engineered DAE mutation in the CP coding region of an infectious clone of TVMV abolished the aphid transmissibility of virus progeny [46]. Gal-On et al. [49] reported the reverse experiment in which change of the DTG motif to DAG restored aphid transmissibility of an infectious clone of ZYMV. Another line of evidence for the key role of the DAG motif in aphid transmission of potyviruses came indirectly from experiments on a potexvirus. Potato virus X (PVX) is the type member of the genus Potexvirus and is not aphid-transmissible in any of the conditions tested. However, for a British isolate of potato aucuba mosaic virus (PaMV), which was ascribed to the Potexvirus genus, transmission was successful when the aphids were first fed on a plant extract containing the HC of PVY potyvirus [29]. The N-terminal region of the potexvirus CP is exposed at the surface of virus particles [50, 51]. Baulcombe et al [52] have reported the presence of a DAG motif between amino acid positions 14 and 16 in the coat protein sequence of the British isolate of PaMV The same authors transferred the corresponding N-terminal region to PVX and thereby produced a recombinant virus, which was transmitted by aphids previously fed on a source of functional HC to a PVY-infected plant. They concluded that the potyvirus-dependent aphid transmission of PaMV was directly attributable to the CP domain, including DAG, located at the N-terminus of the CP Taken together, these data definitely established the pivotal role of the DAG motif in the mechanism of potyvirus transmission by aphids. However, recent
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investigations have demonstrated that the situation may be a little more complicated than first thought. Observation of various potyvirus sequences [53] as well as mutagenesis in TVMV [54] demonstrated that some particular changes might occur in the DAG motif without necessarily inducing the loss of aphid transmission. Furthermore, substitution of residues either upstream or downstream of the DAG sequence can affect aphid transmission of both TVMV [54, 55] and TEV The latter indicates that the amino acid context flanking the DAG motif is also important [56]. C.
Roles of the N-Terminal and DAG Motif Involved in Aphid Transmission
According to the bridge hypothesis, Harrison and Robinson [42] suggested that the DAG motif might primarily be the site of HC attachment to the virus particle. Experimental evidence in support of this hypothesis was gradually obtained over the last decade. By using a protein blotting-protein overlay technique, Blanc et al [57] demonstrated direct attachment of the HC to the DAG motif of the CP of TVMV. The minimum CP domain required for this interaction was seven amino acids (DTVDAGK), including the DAG motif. A series of TVMV mutants in and around the DAG were used to demonstrate a very strong correlation between HC binding and aphid transmission. This fact is corroborated by similar findings for TEV [58], ZYMV [59], and turnip mosaic virus (TuMV) [60, 61]. Hence, the involvement of an interaction between DAG or corresponding motifs and HC-Pro is likely to be a general feature for members of the genus Potyvirus. As described earlier, the flanking context around the DAG is crucial, and variations found in various viruses certainly allow the DAG (or corresponding motives) to be properly exposed and readily accessible for interaction with HC. A different role for the CP N-terminus, and thus DAG, in aphid transmission was recently proposed by Salomon and Bemardi [62]. In this work, a sequence of 59 amino acids corresponding to the N-terminal region of the maize dwarf mosaic virus (MDMV) CP was expressed in bacteria as a C-terminal fusion to the maltose-binding protein. Additional treatment of the fiision protein with factor Xa allowed isolation of the N-terminal domain alone. Subsequent MDMV plant-toplant transmission was highly reduced when aphids were previously fed on solutions containing either the fusion protein or the N-terminal domain alone. These results were interpreted as indirect evidence for the existence of a direct interaction between the CP N-terminus and some putative receptors in the aphid stylets. According to that view, the bacterially expressed N-terminus of the MDMV CP may be capable of interacting with the aphid stylets, and when acquired by aphids, this polypeptide would interact and block the aphid receptors, thereby preventing any subsequent MDMV virion attachment necessary for transmission. Salomon and Bemardi [62] did not definitely conclude that the direct binding of
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the CP to the aphid stylets was operated via the DAG motif. However, they mentioned preliminary results showing that deletion of DAG in the bacterially expressed N-terminal domain of MDMV CP abolished the inhibitory effect of this peptide on aphid transmission. Hence DAG, as also suggested in a recent review [63], was suspected to be the motif responsible for the direct CP attachment to the aphid receptor. The mode of action proposed for the HC in their alternative model differs from the aforementioned bridge hypothesis and will be further discussed.
IV A.
Role of the Helper Component
Loss of Transmissibility by Purified Fotyvirus Virions
The use of artificial acquisition through Parafilm membranes was a useful tool in the study of potyvirus transmission. This made it possible to show that pot3^iruses and a number of other viruses lose their ability to be transmitted by aphids when they are purified [64]. However, acquisition of the supernatant of extracts of potyvirus-infected plants was able to restore the transmissibility to purified virions [31]. As mentioned above, the assisting component that was present in the supernatant was named the helper component (HC). B.
Purification and Characterization of the Helper Component
The first attempts to extract the HC were made in the 1970s by ultracentrifugal precipitation of the virus particles and collection of the HC present in the supernatant fraction [32, 65]. The proteinaceous nature of the HC was determined by techniques that were then available, namely gel filtration and ultrafiltration [65]. Pirone et al. [66] continued further characterization and purification. In these attempts, potyvirus HC was purified to an almost single band of protein by a combination of methods, which included ultracentrifugation to remove the virus particles, density-gradient centrifugation to separate the HC from plant proteins, and affinity chromatography to separate the HC from other, unrelated proteins [66]. The resulting product was analyzed by polyacrylamide-SDS electrophoretic (PAGE) migration. The subunit of the HC ranged in size from 53 kDa for TVMV to 58 kDa for PVY. The specific activity of purified HC was 200 times as active as the crude supernatant and was therefore used for preparation of specific antiserum. The viral nature of HC was discovered when it was established that TVMV HC and PVY HC are serologically distinct [67]. The most convincing evidence for the viral origin of the HC came from cell-free translation of PVY RNA or TVMV RNA, which resulted in a 75-kDa polypeptide
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BENJAMIN RACCAH, HERVE HUET, AND STEPHANE BLANC
[68]. This polypeptide reacted with the respective anti-HC sera, but not with antisera to other potyviral proteins (e.g., the coat protein, cyhndrical inclusion bodies, and nuclear inclusion bodies). Later it was established that cytoplasmic amorphous inclusions of PRSV also react with anti-PVY HC serum [69]. Cell-free translation was also useful to map the viral genes and to designate the gene encoding for HC as the second gene from the 5' end of the viral genome [70]. Nondissociated, biologically active HC seems to be a dimer or a trimer, with a molecular weight ranging between 100 and 150 kDa, as determined by high-pressure liquid chromatography [66] and gel filtration [61]. Consistent HC self-interaction via an N-terminal domain was demonstrated in the yeast two-hybrid system [71, 72]. Recently, a histidine-tag consisting of six histidine residues was fused to the Nterminus of the HC of TEV by using a full-length clone [73, 74], which allows purification of tagged HC employing a nickel-charged resin. In this work, the histidine tag did not alter the activity of HC, nor did the corresponding nickel-based purification method used to isolate the tagged HC. Using a similar approach Kadoury et al. also extracted the HC of ZYMV Surprisingly, in that case, no histidine tag was required, and the wild-type HC was demonstrated to have an intrinsic affinity for nickel-charged resin [60]. These authors showed that the HCs of PRSV, watermelon mosaic virus 2 (WMV2), and turnip mosaic virus (TuMV) potyviruses, but not those of PVY and TVMV, could also be purified by use of nickel-charged resins with no histidine tagging. This suggested that solanaceous hosts are not suitable for HC extraction, in contrast to cucurbits and crucifers. In order to rule out the role of the host, Chenopodium amaranticolor, a common local lesion host for both PVY and ZYMV, was used for the extraction of HC. By using nickel-charged resin, it was demonstrated that the HC of ZYMV but not that of PVY could be extracted [H. Huet and B. Raccah, unpublished results]. Isolation of non-tagged or tagged HC by employing nickel-charged resin enables the purification of large amounts of HC for antibody production, overlay assays, and other studies. The amount of HC required to attain a similar rate of transmission was much greater for the nickel-purified than for the ultracentrifuge-semipurified HC. This relative low activity of the nickel-based resin preparation may be attributed to formation of inactive aggregates due to condensation of the HC molecules in the process of binding to the resin. Another possibility is that part of the HC bound to the resin represents an inactive form of the helper present in the host, as do the amorphous inclusions [69]. Identincation of Functional Domains in the Helper Component
C. 1.
PROTEOLYTIC FUNCTION
A number of studies led to the characterization of the HC open reading frame (ORF) in the potyviral genome [75]. Unlike most other potyviral proteins, the HC
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undergoes a single autoprocessing event at its C-terminus [76]. In these studies, it also was established that the proteol3^ic function of the HC is confined to its Cterminus half; since then, the protein has been termed HC-Pro to connote both its transmission and proteolytic function. The proteolytic activity was demonstrated both in vitro and in planta, showing that it cleaves only in cis as an autocatalytic reaction [76, 77]. Mutagenesis and sequence comparison experiments indicate that it is a papain-like protease, with a cysteine residue instead of a serine in the active site [78]. Sequence similarity of several conserved motifs suggests that this type of protease activity may be shared with the "helperlike" proteins of other members of Potyviridae. This may imply an involvement of protease activity in the aphid transmission process; however, until now this involvement has not been supported by experimental evidence. The 35-kDa proteinase (PI) releases the HC-Pro N-terminus from the viral pol3^rotein. The 35-kDa proteinase is found immediately upstream in the viral polyprotein [79]. For TVMV it was shown that the cleavage site of the N-terminus of the HC-Pro is located between amino acid residues 256 (phenylalanine) and 257 (serine) [80]. In most other potyviruses, a putative cleavage site is present 25 residues downstream of a conserved motif (FIVRG), after a phenylalanine or tyrosine residue [15]. This cleavage site was also found in the ipomovirus sweet potato mild virus [81] and, to some extent, in the wheat streak mosaic rymovirus [82]. 2.
APHID TRANSMISSION FUNCTION
a. Mutations Proven to be Involved in Aphid Transmission, Search for domains that are responsible for helper activity were initiated when genomic information on potyviruses became available. The first comparison was made between PVY^ (a strain with a defective helper) and the wild-type PVY [83]. The comparison of the derived amino acid sequence of the HC-Pro of PVY'^ with that of the wild-type PVY and several other pot5rvAiruses revealed two amino acid changes within areas of conserved sequence. One change was from lysine to glutamic acid (within the KITC conserved box at amino acid positions 50 to 53 in PVY), and the other was from isoleucine to valine (position 225) [83]. The availability of a ftiU-length cDNA clone of TVMV from which infectious transcripts could be produced [84] allowed the introduction of mutations in TVMV HC-Pro that mimic the change seen in PVY^. The change of lysine to glutamic acid in the KITC box of the TVMV polyprotein resulted in loss of HC activity [85]. A natural mutation in this site was also found for the helper-deficient ZYMV (Ct and R5A) [86, 87]. Additional mutations in the KITC motif (glutamine and histidine instead of lysine) resulted in total loss of helper activity [88]. The KITC motif in the type member, PVY, and many other potyviruses may vary and appears with other amino acids with a similar function: these include KLSC in ZYMV [87, 89], bean common mosaic virus [90], peanut stripe virus [91], and soybean mosaic virus (SBMV) [92]; RITC in pea seed-borne mosaic virus [93], clover
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BENJAMIN RACCAH, HERVE HUET, AND STEPHANE BLANC
yellow vein virus [94], and bean yellow mosaic virus (BYMV) [95]; KVSC in peanut mottle virus [91]; KLTC in pepper mottle virus [96]; RTTC in sweet potato feathery mottle virus [97]; and QITC in yam mosaic virus [98]. The KITC motif is located within a conserved cysteine-rich region (Fig. 1, bottom), which is similar to zinc finger motifs in other organisms [99]. Zinc finger motifs were reported to be involved in nucleic acid binding, dimerization, and other protein-protein interactions [100]. Lecoq et al [35] reported an additional helper-deficient potyvirus, namely ZYMV-PAT (poorly aphid-transmissible). A comparison of the HC-Pro of wildtype ZYMV strains to that of ZYMV-PAT revealed an intact KITC box. However, a mutation was found in the central region of the HC in a conserved PTK (positions 309 to 311) box with a change from threonine to alanine [87, 89]. Furthermore, use of the full-length cDNA clone of ZYMV, from which infectious transcripts could be produced [101], enabled mutations to be introduced in the PTK conserved region. The change from threonine to alanine [89] and from proline to alanine [59] resulted in total loss of helper activity. On the other hand, replacement of the positively charged lysine by the negatively charged glutamic acid did not reduce the rate of transmission compared with that of wild type. A significant reduction in helper activity was recorded when threonine was replaced by valine or serine [59]. Interestingly, the PTK motif is conserved in the sweet potato mild mottle ipomovirus (SPMMV) transmitted by whiteflies [81]; however, in rymoviruses the motif was variable, for example, ATG, PTP, and ATP in ryegrass mosaic virus (RGMV), brome streak mosaic virus (BrSMV), and wheat streak mosaic virus (WSMV), respectively [82, 102, 103]. The low similarity among the rymoviruses suggests that the PTK motif may not be involved in transmission of mite-bome viruses. Furthermore, among the bymoviruses that are fungi-borne, the PTK motif is not present at all [104, 105]. b. Other Mutations in the HC-Pro with Potential Effect on Aphid Transmission. Mutations in the highly conserved CG motif region, upstream of the KITC box, produced an aphid transmission-defective HC-Pro in TVMV (CG>SG) [88] and in PVY (CG>CD) [106]. In the case of PVY, an additional mutation occurred in a nonconserved region. Furthermore, HC could be detected in the total extracts of infected plants, and it was shown that purified virus was readily transmitted when mixed with active HC. However, this mutated HC could not be purified and tested in membrane-feeding assays [106]. On the other hand, the HC of TVMV (with the CG mutation) could be extracted and was shown to be deficient in the ability to assist transmission even when concentrated [88]. It has recently been shown that a mutation, phenylalanine to leucine, at amino acid position 10 of the TEV HC totally abolished helper activity [58]. This phenylalanine residue is located near the beginning of the N-terminus, where aromatic amino acids are conserved among potyviruses.
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Finally, a number of other mutations have been reported to be associated with defective HCs in various potyviruses, but their direct involvement in HC inactivation remains to be proved. As an example, a defective helper was reported for TuMV [107]. In this case, six amino acid differences were found between the strains with active and inactive helper. However, none of these mutations were either in the KITC or PTK conserved region, which suggests that other domains may be important for HC activity. Mutations may have primary effects on active sites and secondary effects on structure. 3.
OTHER FUNCTIONS
The first function that was assigned to the HC-Pro gene was assisting aphid transmission (sections C.2.a and C.2.b). However, it is now well documented that the HC-Pro has many more functions and seems to be a key protein in the potyvirus life cycle. The following functions have been assigned to the HCPro: long-distance [108, 109] and cell-to-cell movement [110], RNA binding [75], genome amplification [85, 108, 109], symptom expression [85, 108, 109] synergism [111] and, more recently, silencing repressor [112-114]. Dealing with these functions is beyond the scope of this chapter; therefore, the reader should refer to the reviews of Maia et al [63] and Revers et al [115] for further details. D.
Structure-Function Relationship in HC-Pro 1.
HC-PRO IS REQUIRED FOR VIRUS RETENTION IN THE APHID'S STYLETS
The first experimental evidence for the role of the HC in virus retention in aphid stylets was obtained in 1986 when Berger and Pirone [116] showed that ^2^1-labeled virus was taken into aphid guts at same level in the presence or absence of HC. However, ^^^I-labeled TVMV was only found in the stylets of aphids that acquired the virus in the presence of helper. In the absence of HC, the label was found in the gut. Later, using ^^^I-labeled virions and gold labeling of HC, it was possible to see virions and HC-Pro both attached to the epicuticle of the aphid mouthparts, confirming that retention occurs only when an active helper has been used [117, 118]. These results are in total agreement with the bridge hypothesis [Section VB], which predicts that in order to "bridge between the two," the HC-Pro should be able to interact both with aphid and with virion. 2.
BINDING DOMAINS OF HC-PRO
A direct association between the helper and CP or purified virions was shown by Blanc et al [57], Peng et al [59], and Wang et al [61] (see section D 1). The availabiUty of helper mutants with mutations in the KITC motif and the PTK
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motif made it possible to determine which motif is involved in binding to coat protein (CP) and which in binding to aphid. a. Capsid-Binding Domain ofHC-Pro, The HC from virus strains with a mutation in the PTK motif were nonfunctional and failed to bind to dot-blotted virions, whereas nonmutated functional HC did bind [59]. This shows that the PTK motif is directly or indirectly involved in HC binding to the CP. As described earlier, the most drastic change in the charge of the PTK motif, when a negatively charged glutamic acid residue replaced a positively charged lysine residue, did not alter the helper activity. On the other hand, replacing proline by alanine and threonine by alanine resulted in almost total loss of helper activity [59]. This suggested the possibility that the change from polar to hydrophobic amino acids is the cause for the loss in helper activity. However, when the polar amino acid threonine replaced the also polar serine residue, helper activity was not retained and transmission was greatly reduced. Hence, it is not possible to attribute the activity of the PTK motif to changes in charge or polarity. Moreover, proline residues are "rigid" and therefore often very important for protein conformation. Thus, a structural interaction between the helper (PTK) and the virion may exist. Thus far, the PTK motif of the HC-Pro seems highly conserved among potyviruses. However, in order to generalize about the role of this motif in binding to potyviral virions, there is a need to repeat the experiments reported for ZYMV in additional virus systems. b. Aphid Binding Domain of HC-Pro. Though nonfunctional, HCs from viruses that are mutated in the KLSC (ZYMV-Ct) or in the KITC (PVY and TEV) motif did bind to the blotted virions [59] or to the CP [58]. These findings are consistent with those obtained for the HCs mutated in PTK and clearly suggest that the KITC motif is not functional in direct binding to virions. Furthermore, using immunogold labeling, Blanc et al [58] were able to detect the wild-type HC (with a KITC motif), but not the mutated HC (with an EITC motif), in the stylet. Therefore, it was proposed that the KITC motif might be involved in binding of the HC to the aphid's stylets [58, 59, 89]. In the KITC motif of TVMV, where the lysine is crucial for helper activity, changes to glutamine or histidine resulted in loss of transmissibility [88]. The position of the KITC motif within the zinc finger region of the HC-Pro [99] may lead to speculation that this region is involved in an association with charged "receptors" that are present in the epicuticle lining the aphid mouthparts. However, this cysteine-rich domain may also be involved in dimer formation. The yeast twohybrid system has not allowed clarification of this point so far. Indeed, it was shown that KITC is indispensable for dimerization [72] of lettuce mosaic virus HC-Pro but not for potato virus A (PVA) [71].
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V. Potyvirus Transmission by Aphids A.
Proposed Mechanisms
Several theories have been proposed for the mode of action of HC-Pro which were recently summarized by Pirone and Blanc [7] and Maia et al [63]. The possibility that HC-Pro acts by protecting virions in the aphids or helping the early infection process (after inoculation by aphids together with the virions) are not discussed here, because no supporting data are available. Two models, which are supported by experimental results, will now be considered: the bridge hypothesis (section VB) and the direct binding hypothesis (section VC). In the following sections, we will analyze, in light of all the available data presented above, which of these two models best describes the molecular mechanisms underlying the interaction of potyviruses with their vectors. A number of facts are now well established, thoroughly documented, and generally admitted by all scientists in the field. These facts will be briefly summarized and examined to see if they fit either of these two hypotheses. • HC-Pro must be acquired by aphids prior to or at the time of virus acquisition to mediate transmission. • HC-Pro clearly contains at least two ftmctional domains for aphid transmission, one (the KITC region) involved in HC-Pro retention in the aphid stylets and the other (the PTK region) involved in virion binding. • HC-Pro itself is retained in the aphid's stylets, as indicated both by sequence acquisition feedings and immunogold labeling of thin sections of aphids. • Virions are retained in the aphid's stylets solely in the presence of a functional HC-Pro. • The DAG motif in the N-terminus of the viral CP specifically interacts with the HC-Pro. As indicated above, conflicting data do obviously exist concerning the question of whether the DAG motif interacts with HC-Pro or directly with the aphid mouthparts (see section IIIC). Direct evidence for the attachment of HC-Pro to the DAG motif was published in three independent papers and has been reproduced using additional potyviruses, PVY and PPV (C. Llave and M. Ravelonandro, respectively, personal communications). This accumulation of results and their reproducibility has led us to conclude that if the N-terminal region contained a domain capable of direct attachment to the aphid stylets, this would be a domain other than the DAG or the corresponding motif in various potyviruses. B.
Bridge Hypothesis
We believe that recent findings taken together strongly favor the bridge hypothesis, which was originally formulated by Kassanis and Govier [30]. Indeed,
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confronting this hypothesis with available data results in the following model. The N-terminal region of HC-Pro (KITC domain or elsewhere) recognizes an unknown receptor in the aphid stylets. At the same time or subsequently, an HCPro downstream domain, presumably containing the PTK motif, undergoes a specific interaction with the DAG motif on the virus CP, thereby mediating retention of the virions at appropriate sites in the vector. C. Direct Binding Hypothesis According to the direct binding model and recently published data (see section IIIC), on binding of HC-Pro onto the DAG motif, the CP undergoes a conformational change, which allows direct attachment of virions to putative aphid receptors located in the stylets. We believe that this hypothesis involves several assumptions that are not consistent with the experimental data: 1. To avoid competition problems, this hypothesis implies the existence of two different receptors in the aphid, one specific for the HC and the other specific for the CP. Microscopy studies (see section IVDl) suggest that HC-Pro and virions colocalize in the aphids, a fact that favors a single receptor rather than two distinct ones. 2. The N-terminal CP domain involved in binding to the aphid is a domain other than DAG. Sequence comparison between viruses transmitted by the same aphids indicated that this region is highly variable except for the DAG motif, thus rendering the existence of such an additional domain very unlikely. Additionally, this other domain, which should also be present in the potato aucuba mosaic potexvirus (PaMV), has never been reported. 3. The binding of HC-Pro to the DAG would provoke a conformational change in the CP exposing the aphid-binding domain. This implies that all CPs from viruses that are transmitted with the help of a given HC-Pro have similar folding properties—and so is it for the CP of PaMV. However, considering the tremendous sequence variation found at this level, this possibility is highly unlikely. 4. If the virus were to interact directly with the aphid, one would expect the virus-vector specificity to depend on the origin of the CP. In contrast, the data presented by Wang et al [119] indicate that this specificity depends on the origin of the HC-Pro rather than that of the CP (see section VI). D.
Which Hypothesis Is the Most Likely?
The bridge hypothesis seems to match the overall data presented in this review. The only strong support for the direct binding hypothesis is the report by Salomon and Bemardi [62] on maize dwarf mosaic virus (MDMV) aphid transmission. Blanc et
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al [57] mentioned their failure to reproduce similar experiments on tobacco vein mottling virus (TVMV). It is reasonable to assume that the general mechanisms of the virus-vector interaction are probably common to most, if not all, members of the genus Potyvirus, as demonstrated by frequent compatibility in aphid transmission between the HC of one member and the virions of another. Therefore, unless complementary evidence for a direct CP-aphid interaction is presented for additional potyviruses, one must view the direct binding hypothesis as questionable.
VL Specificity of Potyvirus Transmission by Aphids A.
Types of Specificity
Specificity is a term intended to describe the likelihood of a certain virus to be transmitted by a certain aphid species. In comparison with propagative circulative viruses, potyviruses, as typical nonpersistent noncirculative viruses, were believed to have a low specificity for vector species. In the literature there is ample proof for vector species that do not colonize but serve as vectors on crops [1]. However, there are also reports of differential transmission efficiency of potyviruses by different aphid species. Failure of a given aphid species can derive from: • Specificity that depends on the aphid species used • Specificity that depends on the host from which the virus is acquired or inoculated • Specificity between virions and the helper component • Specificity among aphid species and helper components Three of these forms of specificity are discussed below. B.
Variation in Transmissibility by Different Aphid Species
Variable transmissibility of potyviruses by different aphid species feeding on several target hosts was reported by Sako [20]. Myzus persicae transmitted TuMV at high rate to Raphanus sativus, Spinacia olearaceae, and Physalis floridana but at a lower rate to Zinnia elegans and Chrysanthemum coronarium. A variation in the proportion of viruliferous species among airborne aphids [13] was also recorded. However, variation in specificity may derive from the attraction or repellence between aphid and host. Aphid species may also differ in their affinity for different viruses [120]. C.
Virus-Aphid Specificity That Depends on the HC-Pro Used
Sako et al [120] made use of the fact that the HC of watermelon mosaic virus (WMV) assists the transmission of TuMV to compare three species of aphids.
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Myzus persicae transmitted TuMV from membranes at about the same rate whether the HC was previously acquired from TuMV-infected turnips or from WMV-infected pumpkin. However, when Dactynotus gobonis was used, transmission of TuMV was efficient when the HC source was TuMV-infected turnips but inefficient when the HC source was WMV-infected pumpkin. In using infected plants as a source of HC, Sako et al [120] could not separate host-aphid effects from HC-aphid effects. In a recent study, Wang et al [119] provided convincing evidence that different aphids have different affinities for the HC used for transmission. Aphis gossypii transmitted TEV efficiently with the HC of potato virus Y (PVY) but inefficiently with the HC of TuMV More significantly, Lipaphis erysimi failed to transmit TEV or TuMV with the HC of PVY but successfully transmitted them with that of TuMV In both cases, the HCs and virions were acquired from membranes and inoculated to tobacco or mustard. Wang et al [119] suggest that aphid specificity for a particular HC may relate to the compatibility between their saliva and the HC of the virus involved. D.
Specificity Between HC-Pro and Virions
Potyviral HCs were found to assist transmission not only of their own virions but also virions of other potyviruses [35, 121, 122]. As an example, Lecoq and Pitrat [34] reported high heterologous compatibility between helpers and virions of papaya ringspot virus type W (PRS V-W) and WMV, but poor compatibility between zucchini yellow mosaic virus (ZYMV) and PRSV-W On the other hand, WMV assisted transmission of TuMV virions, but the helper of TuMV failed to assist the transmission of WMV virions [122]. Although there is no direct evidence, the CCC motif (at position 290 upstream from the PTK motif in PVY) may also be crucial for aphid transmission. Indeed some viruses with a modification in the CCC motif also have a change in the DAG motif of the CP: these are ASC and DAAA for peanut mottle potyvirus [91], and CSC and DAS in pea seed-borne mosaic potyvirus [56]. This could imply a coevolution of the two motifs, explaining specificity between the HC and CP. Furthermore, directed mutation of this motif (CCC > RPA) in TEV friU-length clones affects long-distance movement of the mutant, which may be in part explained by interference with particle binding [108].
W/. Concluding Remarks The key feature for understanding NPV transmission is the ability of aphids to retain the acquired virus until it is released by egestion in the target host. With no retention, the virus particles will be flushed into the aphid's digestive system and excreted with the honey dew. The inability of aphids to transmit purified virions is attributed to their failure to retain the virus particles [115]. As stated above (section IVDl), the presence of HC is essential to effect retention; this was demon-
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Stylet Virion
stylet
Virion Stylet
HC-Pro
Virion
CP subunit Fig. 3 Models depicting possible interaction between the HC-Pro, the aphid's stylet, and the pot3rviral coat protein. (A) Position of the virion particle close to the apical section of the food canal. (B) A model assuming an association between two molecules of HC-Pro. Note that one molecule of the HC-Pro is bound to a "receptor" on the stylet whereas the second HC-Pro molecule is bound to the coat protein subunit. (C) A model assuming that a dimer is needed to bind to the receptor on the stylet. Both HC-Pro molecules are linked to coat protein subunits. (D) A proposed structural binding between the PTK motif of the HC-Pro and the DAG motif found on the N-terminus of the coat protein subunit.
strated by transmission studies and by electron microscopy. Two hypotheses that explain how virus attaches to the stylets have been discussed in detail, the bridge hypothesis and the direct binding hypothesis. We have also explained the reasons for favoring the first over the second. However, in order to narrow gaps of knowledge, it will be necessary to determine the specific sites (receptors?) in the aphid's cuticular proteins that may or may not bind to the KITC domain. As indicated above (section IVB), the functionally active helper appears as a dimer (or possibly a trimer). However, we still do not know how the HC dimer functions in aphid transmission. In the absence of experimental data, we would like to speculate on a possible role of the dimer in the process of virus release from the aphid's stylet (Fig. 3). We propose two alternative models. The first (Fig. 3B) assumes that two molecules of the HC are coassociated. The first molecule of the HC dimer is attached on one side to a "receptor" on the stylet, and the other side is linked to the second molecule of HC, which is bound to the CP subunit coating the virion. The second model assumes that dimer formation is essential for binding to a receptor on the aphid stylet; the two molecules bind to the DAG motif of two adjacent CP subunits (Fig. 3C).
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In section IVD2a we detailed the reason for suggesting that the association between the DAG domain and the HC-Pro is based on structure rather than on charge. If this is true, the weakest Hnk in the stylet-HC-virion complex may exist between the two HC molecules. The mechanism of virus release from aphid stylets may then be based on the dissociation of the HC-HC link triggered by the aphid's salivation (see section IIA) or by certain host components. As a result of such dissociation, in the first model, a single HC molecule will remain bound to the virion when released by the aphid in the plant cell in the process of probing. In the second model, the dissociation between the two HC molecules will free the HC from the receptor, allowing the release of the HC-virion complex in the plant cell. In view of the movement functions of the HC-Pro, the HC-virion complex may have certain advantages for translocation of the virion within the infected tissue (Fig. 3D). Future work will be needed to ascertain the role of the HC in the release of virus, in the early infection events, or in both.
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73. Dolja, VV, Peremyslov, VV, Keller, K.E., Martin, R.R., and Hong J. (1998). Isolation and stability of histidine-tagged proteins produced in plants via potyvirus gene vectors. Virology 252, 269-274. 74. Blanc S., Dolja VV, Llave C , and Pirone T.P. (1999). Histidine tagging and purification of tobacco etch potyvirus helper component protein. J. Virol. Methods 77, 11-15. 75. Maia, I.G., and Bemardi, F. (1996). Nucleic acid binding properties of a bacterially expressed potato virus Y helper component-proteinase. J. Gen. Virol. 11, 869-877. 76. Carrington, J.C., Gary, S.M., Parks, T.D., and Dougherty, W.G. (1989). A second proteinase encoded by a plant potyvirus genome. EMBO J. 8, 365-370. 77. Kasshau, K.D., and Carrington, J.C. (1995). Requirement for HC-Pro processing during genome amplification of tobacco etch potyvirus. Virology 209, 268-273. 78. Oh, C.-S., and Carrington, J.C. (1989). Identification of essential residues in potyvirus proteinase HC-Pro by site directed mutagenesis. Virology 173, 692-699. 79. Verchot, J., Hemdon, K., and Carrington, J.C. (1991). Mutational analysis of the tobacco etch potyviral 35-kDa proteinase: Identification of essential residues and requirements for autoproteolysis. Virology 190, 298-306. 80. Mavankal, G., and Rhoads, R.E. (1991). In vitro cleavage at or near the N-terminus of the helper component protein in the tobacco vein mottling virus polyprotein. Virology 185, 721-731. 81. Colinet, D., Kummert, X, and Lepoivre, P. (1998). The nucleotide sequence and genome organization of the whitefly transmitted sweet potato mild mottle virus: A close relationship with members of the family Potyviridae. Virus Res. 53, 187-196. 82. Stenger, D C , Hall, J.S., Choi, I.-R., and French, R. (1998). Phylogenetic relationships within the family Potyviridae: Wheat streak mosaic virus and brome streak mosaic virus are not members of the genus Rymovirus. Phytopathology 88, 782-787. 83. Thombury, D.W., Patterson, C.A., Dessens, J.T., and Pirone, T.P. (1990). Comparative sequence of the helper component (HC) region of potato virus Y and a HC-defective strain, potato virus C. Virology 178, 573-578. 84. Domier, L.L., Franklin, K.M., Hunt, A.G., Rhoads, R.R., and Shaw, J.G. (1989). Infectious in vitro transcripts from cloned cDNA of a potyvirus, tobacco vein mottling virus. Proc. Natl. Acad. Sci. U.S.A. 86, 3509-3513. 85. Atreya, CD., Atreya, PL. Thombury, D.W., and Pirone, T.P (1992). Site directed mutations in the potyvirus HC-Pro gene affect helper component activity, virus accumulation and symptom expression in infected tobacco plants. Virology 191, 106-11. 86. Grumet, R., Bada, R., and Hammar, S. (1992). Analysis of the zucchini yellow mosaic virus (ZYMV) potyviral helper component, possible identification of an aphid-interaction domain. (Abstract). Phytopathology 82, 1176. 87. Granier, F, Durand-Tardiff, M., Casse-Delbart, F, Lecoq, H., and Robaglia, C (1993). Mutation in the zucchini yellow mosaic virus helper component associated with loss of aphid transmissibility. J. Gen. Virol. 74, 2737-2742. 88. Atreya, C D , and Pirone, T.P. (1993). Mutational analysis of the helper component proteinase gene of a potyvirus: Effects of amino acid substitutions, deletions, and gene replacement on virulence and aphid transmissibility. Proc. Natl. Acad. Sci. U.S.A. 90, 11919-19123. 89. Huet, H., Gal-On, A., Meir, E. Lecoq, H., and Raccah, B. (1994). Mutations in the helper component (HC) gene of zucchini yellow mosaic virus (ZYMV) affect aphid transmissibility. J. Gen. Virol. 75, 1407-1414. 90. Fang, G.W., Allison, R.E, Zambolim, E.M., Maxwell, D P , and Gilbertson, R.L. (1995). The complete nucleotide sequence and genome organization of bean common mosaic virus (NL3 strain). Virus Res. 39, 13-23.
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91. Flasinski, S., and Cassidi, B.G. (1998). Potyvirus aphid transmission requires helper component and homologous coat protein for maximal efficiency. Arch. Virol. 143,2159-2172. 92. Jayaram, C , Hill, J.H., and Miller, W.A. (1992). Complete nucleotide sequences of two soybean mosaic virus strains differentiated by response of soybean containing the Rsv resistance gene. J. Gen. Virol. 73,2067-2077 93. Johansen, E., Rasmussen, O.F., Heide, M., and Borkhardt, B. (1991). The complete nucleotide sequence of pea seed-borne mosaic virus RNA. J. Gen. Virol. 72,2625-2632. 94. Takahashi, Y., Takahashi, T., and Uyeda, I. (1997). A cDNA clone to clover yellow vein potyvirus genome is highly infectious. Virus Genes 14, 235-243. 95. Guyatt, K.J., Proll, D.E, Menssen, A., and Davidson, A.D. (1996). The complete nucleotide sequence of bean yellow mosaic potyvirus RNA. Arch. Virol. 141,1231-1246. 96. Vance, VB., Moore, D., Turpen, T.H., Bracker, A., and Hollowell, VC. (1992). The complete nucleotide sequence of pepper mottle virus genomic RNA: Comparison of the encoded polyprotein with those of other sequenced potyviruses. Virology 191, 19-30. 97. Sakai, X, Mori, M., Morishita, T, Tanaka, M., Hanada, K., Usugi, T, and Nishiguchi, M. (1977). Complete nucleotide sequence and genome organization of sweet potato feathery mottle virus (S strain) genomic RNA: The large coding region of the PI gene. Arch. Virol. 142,1553-1562. 98. Aleman, M.E., Marcos, J.E, Brugidou, C , Beachy, R.N., and Fauquet, C. (1996). The complete nucleotide sequence of yam mosaic virus (Ivory Coast isolate) genomic RNA. Arch. Virol. 141, 1259-1278. 99. Robaglia, C , Durand-Tardiff, M., Tronchet, M. Boudazin, G., Astier-Manifacier, S., and CasseDelbart, F. (1989). Nucleotide sequence of potato virus Y (N strain) genomic RNA. J. Gen. Virol. 70, 935-947. 100. Berg, J.M. (1990). Zinc fingers and other metal-binding domains. J. Biol. Chem. 265,6513-6516. 101. Gal-On, A., Antignus, Y, Rosner, A., and Raccah, B. (1991). Infectious in vitro RNA transcripts derived from cloned cDNA of cucurbit potyvirus, zucchini yellow mosaic virus. J. Gen. Virol. 72,2639-2643. 102. Gotz, R., and Maiss, E. (1995). The complete nucleotide sequence and genome organization of the mite transmitted brome streak mosaic rymovirus in comparison with those of potyviruses. J. Gen. Virol. 76,2035-2042. 103. Gibbs, A.J., Mackenzie, A.M., and Keese, P. (1998). The complete nucleotide sequence of an Australian isolate of ryegrass mosaic virus. Unpublished accession number AAC25028. 104. Kashiwazaki, S., Minobe, Y, and Hibino, H. (1991). Nucleotide sequence of barley yellow mosaic virus RNA2. J. Gen. Virol. 72, 989-993. 105. Namba, S., Kashiwazaki, S., Lu, X., Tamura, M., and Tsuchizaki, T (1998). Complete nucleotide sequence of wheat yellow mosaic bymovirus genomic RNAs. Arch. Virol. 143, 631-643. 106. Canto, T, Lopez-Moya, J.J., Serra-Yoldi, M.T., Diaz-Ruiz, J.R., and Lopez-Abella, D. (1995). Different helper component mutations associated with lack of aphid transmissibility in two isolates of potato virus Y Phytopathology 85, 1519-1525. 107. Nakashima, H., Sako, N., and Hori, K. (1993). Nucleotide sequences of the helper componentproteinase genes of aphid transmissible and non-transmissible isolates of turnip mosaic virus. Arch. Virol. 131, 17-27. 108. Cronin, S., Verchot, X, Haldeman-Cahill, R., Schaad, M.C., and Carrington, XC. (1995). Longdistance movement factor: A transport function of the potyvirus helper component proteinase. Plant Cell 7, 549-559. 109. Kasschau, K., Cronin, S., and Carrington, XC. (1997). Genome amplification and long-distance movement functions associated with the central domain of tobacco etch potjrvirus helper component-proteinase. Virology 228, 251-262.
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110. Rojas, M.R., Zerbini, KM., Allison, R.E Gilbertson, R.L., and Lucas, WJ. (1997). Capsid protein and helper component-proteinase function as potyvirus cell-to-cell movement proteins. F/ra/ogy 237, 283-295. 111. Shi, X.M., Miller, H., Verchot, J., Carrington, J.C, and Vance, VB. (1997). Mutations in the region encoding the central domain of helper component-proteinase (HC-Pro) eliminate potato virus X/potyviral synergism. Virology 231, 3 5 ^ 2 . 112. Anandalakshmi, R., Pruss, G.L., Ge, X., Marathe, R., Mallory, A.C., Smith, T.H., and Vance, VB. (1998). A viral supressor of gene silencing in plants. Proc. Natl. Acad. Sci. U.S.A. 95, 13079-13084. 113. Brigneti, G., Voinnet, O., Li, W.-X., Ji, L.-H., Ding, S.-H., and Baulcombe, D.C. (1998). Viral pathogenicity determinants are suppressors of transgene silencing in Nicotiana benthamiana. EMBO J. 11,6739-6746. 114. Kasshau, K.D., and Carrington, J.C. (1998). A counterdefensive strategy of plant viruses: Suppression of posttranscriptional gene silencing. Cell 95,461^70. 115. Revers, R, LeGall, O., Candresse, T., and Maule A.J. (1999). New advances in understanding the molecular biology of plant/potyvirus interactions. Mol. Plant Microbe Interact. 12, 367-376. 116. Berger, PH., and Pirone, T.P (1986). The effect of helper component on the uptake and localization of potyviruses in Myzuspersicae. Virology 153, 256-261. 117. Ammar, E.D., Jarfors, U, and Pirone, T.P. (1994). Association of potyvirus helper component protein and the cuticle lining the maxillary food canal and foregut of an aphid vector. Phytopathology ^4, 1054-1060. 118. Wang, R.Y., Ammar, E.D., Thombury, D.W., Lopez-Moya, J.J., and Pirone, T.P (1996). Loss of potyvirus transmissibility and helper component activity correlates with nonretention of virions in aphid stylets. J. Gen. Virol. 11, 861-867. 119. Wang, R.Y. Powell, G., Hardie, J., and Pirone, T.P (1998). Role of the helper component in vector-specific transmission of potyviruses. J. Gen Virol. 79, 1519-1524. 120. Sako, N. Yoshioka, K., and Eguchi, K. (1984). Mediation of helper component in aphid transmission of some potyviruses. Ann. Phytopathol. Soc. Jpn. 50, 515-521. 121. Pirone, T.P. (1981). Efficiency and selectivity of the helper-component-mediated aphid transmission of purified potyviruses. Phytopathology 71, 922-924. 122. Sako, N., and Ogata, K. (1981). Different helper factors associated with aphid transmission of some potyviruses. Virology 112, 762-765.
CHAPTER 11
Viral Determinants Involved in Luteovirus-Aphid Interactions VERONIQUE BRAULT VERONIQUE ZIEGLER-GRAFF K.E. RICHARDS
/. Introduction The genome of viruses in the family Luteoviridae consists of a single positivesense linear RNA molecule of about 5.7 kilobases (kb) packaged in small icosahedral virions measuring 25 nm in diameter [1-3]. Typical symptoms on host plants are dwarfing, reddening, or yellowing of leaves as well as leaf deformations, including rolling and stiffening. Virus infestations can be pernicious to major crop species [4-6]. Family members display two important and related biological characteristics that are of relevance here: (1) strong tissue tropism, with the virus multiplying and residing exclusively in the phloem tissue of infected host plants; and (2) obligate transmission in a persistent, circulative manner by phloem-feeding aphids (Homoptera: Aphididae). The family Luteoviridae is now divided into three genera: Luteovirus, type species barley yellow dwarf virus-PAV (BYDV-R\V); Polerovirus, type species potato leafroll virus (PLRV), and Enamovirus, type species pea enation mosaic virus-1 (PEMV-1) [7]. The genomic RNAs of all three genera contain five or six main open reading frames (ORFs) arranged in two clusters (Fig. 1). In the 5'-proximal cluster, ORFS 1 and 2 encode the viral RNA-dependent RNA polymerase and display distinct taxonomic affinities. Luteovirus polymerase is Carmovirus-like, whereas Polerovirus and Enamovirus polymerases are sobemovirus-like [2]. Poleroviruses and PEMV-1 possess a 5'-terminal genome-linked protein (VPg) derived from ORE 1 [8, 9], whereas luteoviruses do not [2]. Poleroviruses and PEMV-1 also contain a 5'-proximal ORE 0 of unknownfrinctionwhich is not present in luteoviruses. Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS
Genus Luteovirus
(BYDv-PAV)
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Gtrms Polerovirus (PLRV)
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Fig. 1 Genome organizations of the type species of three genera in the family Luteoviridae: Luteovirus, barley yellow dwarf virus-PAV (BYDV-PAV); Polerovirus, potato leaf roll virus (PLRV); and Enamovirus, pea enation mosaic virus-1 (PEMV-1). Numbered rectangles correspond to main open reading frames (ORFs). Diagonal arrows represent the approximate sites of translational frameshifts by which the product of ORF 2 is fused to that of ORF 1. Horizontal arrows denote the positions of the termination codon of ORF 3 and the major coat protein (CP) cistron, which undergoes translational suppression to produce the readthrough protein in which the product of ORF 5, the readthrough domain, is fused to the C-terminus of the major CP.
The genes in the 3'-proximal cluster are translated from a subgenomic RNA. Open reading frame 3 encodes the 22 kDa major viral coat protein (CP). The translation-termination codon of ORF 3 is suppressed by a translating ribosome, thus extending translation into the downstream ORF 5. The resulting 75-kDa readthrough (RT) protein has a 54-kDa readthrough domain (RTD), encoded by ORF 5, fused to the C-terminal end of the CP [10-13]. The RT protein is a minor component of the virion capsid [14-20]. In the luteoviruses and poleroviruses but not PEMV-1, an ORF 4 is embedded in the CP cistron in another reading frame. The ORF 4 protein is probably a viral movement protein [21, 22]. Its absence from PEMV-1 can be accounted for by the fact that in nature PEMV exists as a complex between PEMV-1 and a defective umbravirus, PEMV-2. The latter is believed to provide the missing movement fimctions [20]. In the following discussion, the term luteovirus is used to refer to members of that genus alone and occasionally, in a general sense, to members of all three genera.
//. A.
Viral Passage through the Aphid
Acquisition
The circuit that a luteovirus traces within its vector during transmission is complex. The aphid is an active partner in transmission and most, if not all, of
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•^mw^m Fig. 2 Model for transport of a luteovirus across a gut epithelial cell. Virions are recognized at the gut cell apical plasmalemma (APL) and bind to the membrane (1), initiating virus invagination (2) into the coated pits (CP) that bud off the APL as coated vesicles (CV) (3). Fusion of the CV to larger smooth vesicles (L) called receptosomes (RS) acts to concentrate the virus (4). Tubular vesicles containing virions form on the receptosomes (4) and transport the virus (5) to the basal plasmalemma (BPL). Fusion of the tubular vesicles with the BPL (6) releases viral particles to diffuse through the basal lamina (BL) to the hemocoel (7). MT-microtubules. (Reproduced with permission fromGildow[23].)
the steps in viral passage involve "hijacking" of normal cellular mechanisms in the vector. Thus, our expanding knowledge of intra- and intercellular trafficking in animals should eventually shed light on luteovirus transmission, and vice versa. Most of our current knowledge of luteovirus-aphid interactions has been gleaned from the biological and ultrastructural observations of viruliferous aphids initiated by Rochow and colleagues [5, 6] and carried on by Gildow [23]. These researchers mainly employed BYDV and cereal aphids, but the basic conclusions drawn from their experiments appear to apply to other virus-aphid combinations as well. The process involves three major steps. After ingestion by the aphid during feeding in the phloem of infected plants, virions move through the alimentary canal to the hindgut or, in the case of poleroviruses, the posterior midgut [24, 25; C. Reinbold, personal communication]. Here, some virions associate with the epithelial cell apical plasmalemma and form coated pits by a mechanism reminiscent of receptor-mediated endocytosis (Fig. 2). Once taken up, individual virions are observed in small coated vesicles and in clusters in larger vesicles known as receptosomes within the epithelial cell cytoplasm. Virions in linear arrays are also frequently observed in smooth, tubelike vesicles, which are believed to bud off from the receptosomes. The virion-loaded smooth tubules can occasionally be observed in fusion with the basal plasmalemma, which suggests
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VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS
ntmmti
Fig. 3 Passage of a luteovirus through an accessory saHvary gland (ASG) cell. Luteoviruses suspended in the hemolymph first encounter the ASG's extracellular basal lamina (BL). Depending on the luteovirus, virions may be prevented from penetrating the BL (A) or diffuse through it (B, C) to the basal plasmalemma (BPL) and into plasmalemma invaginations (PLI). Luteoviruses not recognized at the BPL remain outside the cell in the pericellular space (B). Luteoviruses (C) that move into PLI and are recognized by putative receptors on the BPL are endocytosed (1) by coated pits (2) and accumulated in tubular vesicles (TV) in the cytoplasm (3). Coated vesicles containing individual virions (CV) bud off from the tubular vesicles (4, 5) and ftise to the apical plasmalemma (APL) (6), forming coated pits (CP). Virions are released via exocytosis into the central salivary canal lumen (C), transported in salivary secretions through the salivary duct (SD) system to the salivary canal of the interlocked maxillary stylets, and finally inoculated inplanta. (Reproduced with permission from Gildow and Gray [30].)
that they represent the principal vehicle for delivery of the virus to their site of exocytosis into the hemocoel. B.
Inoculation
Following acquisition and entry to the hemocoel, virions diffuse through the hemolymph until they encounter an accessory salivary gland (ASG). There is indirect evidence that virus in the hemocoel does not remain in isolated form but instead exists as a complex with symbionin, a GroEL homologue produced by the primary aphid endosymbiont [26]. Symbionin is abundant in the hemolymph, and its interaction with the virion may protect it from destruction when released into the hemocoel (see section IV). The final act of the virus acquisition-inoculation circuit occurs at one of the aphid's pair of ASGs (Fig. 3). These small organs consist of only four secretory cells each and are separated from the hemocoel by an extracellular basal lamina. Virions must penetrate this latter barrier before coming into contact with the
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VIRAL DETERMINANTS INVOLVED IN LUTEOVIRUS-APHID INTERACTIONS
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secretory cell's plasmalemma. The basal plasmalemma is highly invaginated, forming a multitude of membrane-lined canals that reach into the cytoplasm. Virions accumulate within these invaginations where they can undergo endocytosis into coated vesicles. Tube-shaped smooth vesicles containing linear arrays of virions also occur in the cytoplasm, but it is not known whether they represent fusion products of smaller vesicles or arise de novo by membrane invagination. Such tube-shaped vesicles are frequently seen near the microvilli-lined apical plasmalemma of the secretory cells in the process of budding off coated vesicles containing a single virion. These small coated vesicles appear to be the vehicle for delivery of virions to the apical plasmalemma, where they are released into the salivary duct by exocytosis. The reader will note that the viral endocytosis-exocytosis cycle in the ASG involves many of the same types of vesicular structures observed during virus movement through gut epithelial cells but that the process runs in reverse. C.
Sites Governing Virus-Vector Specificity
Luteovirus transmission displays considerable vector specificity; the nature of the selective interactions between virus and vector is of much current interest. Almost all early research on specificity was carried out with a series of five BYDV "strains." These strains are preferentially transmitted by different aphid species, with each BYDV strain being identified by an acronym referring to its most efficient vector(s): BYDV-PAV (transmitted by Rhopalosiphum padi and Sitobion avenae\ BYDV-MAV (transmitted by S. [Macrosiphum] avenae), BYDV-RPV (now CYDV-RPy transmitted by R, padi), BYDV-RMV (transmitted by Rhopalosiphum maidis) and BYDV-SGV (transmitted by Schizaphis graminum) [6]. Studies with different virus-aphid combinations revealed that, as a general rule, specificity is associated with inoculation of virus (i.e., movement of the virus through the ASGs), rather than acquisition (i.e., uptake of the virus by the gut epithelial cells and its delivery into the hemocoel). An aphid species that is a nonvector for a given BYDV strain is generally able to accumulate virus in the hemocoel on feeding on infected plants but is unable to transmit it. The latter is true even if purified virus is injected at high concentrations directly into the hemocoel to bypass the hindgut barrier [27-29]. One exception to this rule was noted: CYDV-RPV was never detected in the hemolymph of the nonvector Metopolophium dirhodum, even though it was readily detectable in that of other nonvector aphids tested [29]. The virus was observed in the gut lumen of M dirhodum but never in close association with the apical plasmalemma, within coated pits, or inside the various vesicular structures observed in hindgut epithelial cells. These observations suggest that M dirhodum hindgut cells lack the receptor(s) on their apical plasmalemma necessary for recognition of this particular luteovirus. The foregoing observations point, by process of elimination, to the ASG as the principal site at which recognition between a given luteovirus and aphid vector
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VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS
occurs. So far, two selective ASG barriers have been identified, the extracellular basal lamina that separates the hemocoel from the ASG and the underlying basal plasmalemma. After acquisition (either by feeding or microinjection) virions are rarely seenfi*eein the hemocoel but are particularly abundant within the ASG's extracellular basal lamina [30]. This suggests that the basal lamina either binds luteoviruses so avidly as to sweep the hemolymph fi"ee of virions, represents a bottleneck in the transmission circuit, or both. No association between viral particles and the basal lamina of other organs, such as the principal salivary glands, was observed, indicating that the ASG basal lamina has special properties conducive to its interaction with virus. Experiments with different BYDV-aphid combinations revealed different types of virus interaction with the ASG basal lamina [30, 31] (Fig. 3). When CYDV-RPV was injected into the hemocoel of the efficient vector R. padi, the inefficient vector S. graminum, and three nonvector species, S. avenue, M. dirhodum, and R. maidis, virions became embedded within the ASG basal lamina of all but one of the species, R. maidis, which suggests that the basal lamina of R. maidis lacks critical recognition elements for this particular virus. In the case of M. dirhodum, virions were observed in the ASG basal lamina but only at low densities and rarely more than midway into the basal lamina. This indicates that the virus is recognized at the basal lamina but not efficiently transported across the structure. In the efficient vector species, R. padi, virions were frequently observed embedded in the ASG basal lamina, in invaginations of the basal plasmalemma, in various endocytotic vesicles in the cytoplasm of the ASG cells, and in salivary ducts. A similar situation prevailed for the inefficient vector species, S. graminum, but virions associated with the various structures in lower numbers. The basal lamina is a complex three-dimensional matrix composed principally of collagen and laminin along with various minor components that may significantly influence its properties in different organs and species [32, 33]. The particle-size exclusion limit of the aphid ASG basal lamina is reportedly 20-30 nm, which could permit free passage of the 25-nm luteovirus virions. Nevertheless, the previously mentioned observations indicate that not all luteoviruses can penetrate the matrix, and Peiffer et al. [31] have suggested that specific protein sequences present in the luteovirus capsid protein(s) can transiently and locally loosen the matrix cross-links so that virions can pass through. The CYDV-RPV virions associated in large numbers with the ASG basal lamina of the nonvector species S. avenae. Virions were also present in the space between the basal lamina and the plasmalemma but not in the cytoplasm of salivary cells or in the salivary duct. This suggests that CYDV-RPV virions can traverse the basal lamina filter of *S. avenae but that the aphid species does not have the appropriate receptors to trigger endocytotic uptake of the virus at the ASG basal plasmalemma.
11.
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213
Identifying Virai Proteins invoived in Transmission
Virus-vector specificity implies that a luteovirus must possess structural determinants that are recognized at various vector barriers to permit uptake and passage of virus. Recent developments may help to identify these viral determinants. Sitedirected mutagenesis is a powerful tool for mapping functions on the viral genome. Full-length cDNA corresponding to several luteoviruses is available and can be used to generate infectious viral RNA by runoff transcription in vitro [34-36]. The major obstacle in using such transcripts to identify viral motifs involved in transmission stems from the fact that luteovirus RNA cannot be transmitted to plants by mechanical inoculation. A means of overcoming this difficulty is provided by agroinfection. Agroinfection (or agroinoculation) takes advantage of the ability of Agrobacterium tumefaciens harboring the Ti plasmid to efficiently transfer a portion of this plasmid, the T-DNA, to the chromosome of a plant cell in the vicinity of a wound [37]. The viral cDNA is placed within the TDNA behind a constitutive plant promoter such as the cauliflower mosaic virus 35S promoter. Optionally, a transcription termination sequence, a ribozyme sequence, or both may be placed at the 3'-terminus of the cDNA. Once the T-DNA arrives in the nucleus of a plant cell, the viral cDNA is transcribed to produce fulllength viral RNA, which can then be translated and replicated autonomously. Successful agroinfection has been described for three poleroviruses, beet western yellows virus (BWYV) [38], PLRV [39], and cucurbit aphid-borne yellows virus (CABYV) [36]. Most of the remaining discussion will center on work with the BWYV system, for which the most information is available. The role of the various viral gene products in transmission has been examined by site-directed mutagenesis using the infectious BWYV cDNA clone. Mutations were engineered into each of the major ORFs, and full-length cDNA carrying each mutation was generated. First, full-length RNA transcripts carrying the various mutations were inoculated to protoplasts to determine if the mutation affected replication of the virus. If replication was successful, extracts of the infected protoplasts were sometimes used as the virus source for acquisition by aphids through membranes. More commonly, however, mutants that passed the protoplast replication test were inoculated to plants by agroinfection, and transmissibility was tested from this infected material. Generally, both standard (8 aphids per test plant) and high (30 aphids per test plant) inoculum pressures were used; test plants were assayed by enzyme-linked immunosorbent assay (ELISA) for virus infection 3 to 4 weeks later. It is evident that results of transmission tests from agroinfected plants will be difficult to interpret if a mutation significantly lowers virus titer in the plant tissue relative to that observed for wild-type infections. Consequently, virus was also routinely purified from agroinfected plants, and transmission tests were carried out by membrane feeding of aphids on virus solutions adjusted to an appropriate concentration. Finally, to examine the role of viral proteins in steps in the trans-
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mission process subsequent to movement across the gut-hemocoel barrier, purified virions were also microinjected directly into the hemocoel of nonviruliferous Myzus persicae nymphs, which were then allowed to feed on test plants. Because RNA viruses are known to have high mutation rates [40], the appearance of revertants or second-site mutations that restore the function targeted by a particular point mutation is a concern. Consequently, the stability of the various mutations, both in planta and following successful aphid transmission, was routinely checked by sequence analysis of the mutant progeny RNA. A.
Viral Nonstructural Proteins Are Dispensable
In a first screen, null mutations were introduced into the major ORFs encoding viral nonstructural proteins (i.e., ORFs 0, 1,2, and 4). Except for mutations in ORFs 1 and 2, which inhibit viral repHcation in protoplasts, mutants were tested for their ability to agroinfect whole plants and to be aphid-transmitted. Expression of ORF 0 was blocked by introducing a point mutation in the initiation codon and by creating a small deletion in ORF 0 upstream of the ORF 1 initiation codon. The two resulting mutations were combined in a third mutant. The three mutants accumulated in agroinfected plants but only at 10 to 20% of wild-type levels [41]. Aphids were able to acquire the mutants from agroinfected plants and efficiently transmit them to test plants. The original mutations were conserved in viral progeny in both agroinfected and aphid-infected plants. Furthermore, no significant second-site mutations occurred in the vicinity of the primary mutations. To eliminate expression of ORF 4, the initiation codon was modified and two in-frame stop codons were introduced near the middle of the ORF in a way that did not alter the sequence of the ORF-3 protein. The resulting mutant, BW5.1845, accumulated efficiently in agroinfected Nicotiana clevelandii. This mutant was also efficiently transmitted by the vector, and the mutations were conserved in its progeny [41]. We conclude that neither ORF-0 nor ORF-4 proteins intervene in aphid transmission. B.
Both Viral Structural Proteins Are Required for Transmission 1. THE Major Coat Protein
Mutation of the initiation codon of the cistron for CP (mutant BW4.2) did not interfere with viral RNA replication in protoplasts but dramatically inhibited virus accumulation after agroinfection of A^. clevelandii [41]. There was some evidence for amplification of the mutant in the vicinity of the agroinoculation sites; however, the CP-defective virus is apparently incapable of moving from these sites. The AUG null mutation in BW4.2 eliminates translation not only of CP but also of the RT protein. To rule out the possibility that the absence of the RT protein, rather than the CP, was responsible for the failure of the BW4.2 to
11.
VIRAL DETERMINANTS INVOLVED IN LUTEOVIRUS-APHID INTERACTIONS
215
accumulate following agroinfection, a second mutant (BW6.26) was produced, in which the CP termination codon was replaced by a sense codon. The resulting mutant multiplied well in protoplasts and produced RT protein but no CP. Like BW4.2, this mutant too was incapable of accumulating in A^. clevelandii following agroinfection [41]. We conclude that the CP and presumably virus assembly are required for efficient proliferation of BWYV throughout an agroinfected plant. This conclusion is consistent with the observation that viruslike particles are observed in the plasmodesmata connecting nucleate phloem cells to the sieve elements of virus-infected plants [42-44], suggesting that virions are an important, if not the only, infectious entity involved in longdistance transport through the vascular system. In preliminary experiments, point mutations were introduced into the major CP sequence without altering the sequence of the ORF-4 protein. These mutants then were assessed for aphid transmissibility. So far, three classes of mutation have been detected: (1) mutants that provoke a defect in virion morphogenesis and, hence, are nontransmissible; (2) mutants that have no effect on virus morphogenesis and aphid transmission; and (3) mutations that do not interfere with virus assembly but do inhibit transmission ([44]; unpublished observations). Thus the major CP certainly carries aphid transmission determinants sensu stricto. Future research will focus on characterization of these determinants and analysis of their site(s) of action in the aphid. 2.
THE RT PROTEIN
As noted earlier (section I), the polypeptide sequence encoded by ORF 5 is expressed by occasional translational suppression of the stop codon of ORF 3. The resulting RT protein is a fusion protein, with the major CP at its N-terminus and the readthrough domain (RTD) derived from ORF 5 at its C-terminus. The RT protein is a minor component of the viral capsid. The RTD in virions is accessible to specific antibodies; this suggests that the RT protein is anchored in the virion by the CP moiety, with the RTD exposed on the particle surface [18]. The RTD is not essential for viral morphogenesis [16] but is required for efficient viral accumulation in plants. It also could play a role in systemic movement of virus in planta [18, 45]. Deletion of the entire RTD (mutant BW6.4) dramatically reduces but does not eliminate virus accumulation in agroinfected plants [18]. Sequence comparisons among different luteoviruses and poleroviruses have revealed considerable homology among their RTD sequences [1, 46]. The region immediately downstream of the major CP stop codon is a proline-rich tract of 16 to 30 amino acid units, which may serve as a tether joining the CP moiety to the rest of the RTD. This proline tract, also referred to as the proline hinge [46], is followed by a region of 210 amino acids, which displays a high degree of homology among all the poleroviruses and luteoviruses. We shall refer to this region as the conserved domain. The C-terminal half (the variable region) of the RTD, on the
216
VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS RT
Mutants in the conserved domain of the RTD
Mutants in the variable domain of the RTD
/'BW6.51 _ BW 6.106BW 6.104 _ BW 6.AMTBW 6.ATB BW 6.50 •
r
BW6.1 BW6.AE1
^
V
Punctual mutants in the conserved domain of the RTD
1
(
i l l
^
^
<^/
Readthrou^h Domain
11
<^/ ^
BW 6.40 BW6.41 BW6.MHD 1
f V <^/ ^
x5j
Fig. 4 Genetic map of BWYV and structure of the readthrough domain mutants discussed in this chapter. Open reading frames (ORFs) are symbolized by numbered rectangles and the viral structural proteins, coat protein (CP), and readthrough protein (RT) by arrows. The conserved domain of the readthrough domain (RTD) is stippled. Sequences deleted in the conserved-domain and variabledomain RTD mutants are represented by horizontal lines. Frameshifts introduced in mutants BW6.43 and BW6.1 are symbolized by dotted lines. The mutation of the Myzus persicae homology domain (MHD) is represented by a black square. Point modifications introduced in the conserved domain of the RTD are indicated by vertical arrow heads.
Other hand, is not well conserved except for a stretch of 45 amino acids, which is very similar among poleroviruses transmitted by M persicae (CABYV, PLRV and BWYV). This sequence has been referred to as the M persicae homology domain (MHD) (Fig. 4). An unexpected finding concerns the nature of the RT protein present in purified BWYV particles. In BWYV virions, the RT protein is present in a truncated form (P74*) of about 54 kDa, produced by loss of C-terminal sequences [18]. Analogues of P74* have also been observed in purified preparations of members of the Luteovirus and Polerovirus genera [11, 14, 17, 19] which suggests that cleavage of the RT protein is not due to accidental degradation but reflects a conserved, presumably biologically significant processing event. Based on the appar-
11.
217
VIRAL DETERMINANTS INVOLVED IN LUTEOVIRUS-APHID INTERACTIONS
Table I Aphid Transmission of Beet Western Yellows Virus (BWYV) Mutants in the Variable Domain of the Readthrough (RT) Protein" Aphid transmission from agro-infected plants^ BWYV mutants BW6.4 BW6.43 BW6.1 BW6.AE1 BW6.40 BW6.41 BW6.MHD BW-WT
Aphid transmission from purified viruses^
RT incorporation into virions
8 Aphids/pl.
30^ Aphids/pl.
8 Aphids/pl.
30 Aphids/pl.
— —
0% 0% 0% 0% 0% 0% 60% 73%
0% 0% 20%/ 22% 71% 8% nd 97%
0% nd^ 20%^ 60% 97% 100% 100% 95%
0% nd 100%/ 100% nd nd nd 100%
+ +++ +++ +++ +++ +++
" Results are presented as the percentage of infected plants/plants tested. Results of transmission of two mutants which do not synthesized the RT protein are presented in the table in shaded boxes. ^ The virus source was N. clevelandii leaves 3 to 6 weeks after agro-inoculation. After a 24-h-virus acquisition access period, aphids (8 or 30) were transferred to test plants for an inoculation feeding period of 4 days. '^ Purified virus at 25 |ig/ml was supplied to aphids by membrane-feeding. ^ "High inoculum pressure" was applied: 16 to 100 (but usually 30) aphids were transferred to each test plant. ^ nd = not determined. ^Transmission events related to pseudo-revertants.
ent molecular weight of P74*, the site of cleavage to produce the truncated product of the RT protein is predicted to He near the boundary between the conserved and variable domains. In view of its presence as a minor component in virions, the question naturally arises as to whether the RT protein and more particularly, the RTD contain essential aphid transmission determinants. To test this hypothesis, expression of the RTD was abolished in two mutants. In BW6.4, the entire ORF 5 was deleted. In BW6.43, the suppressible UAG stop codon of the major CP was reinforced by introducing two additional stop codons (UAA and UGA) directly downstream. A +1 frameshift was introduced during creation of the two extra stop codons (Fig. 4). Whatever the viral source (agroinfected plants, protoplasts, extracts, or purified virus) and the inoculum pressure, these ORF 5 mutants could not be transmitted by the vector (Table 1). When virions of BW6.4 were microinjected directly into the hemocoel, no transmission was observed [18]. This was the first evidence for an essential role of the RTD in aphid transmission of BWYV Evidence for a role of the RTD in luteovirus transmission has been provided by Chay et al. [21], who showed that the RT protein was required for transmission of
218
VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K.E. RICHARDS
BYDV-PAV by R. padi. These researchers [21] used a complementation system in which protoplasts were coinfected with an RTD mutant and a movement protein mutant (ORF 4 mutated). The RNA corresponding to the RTD mutant was heterologously encapsidated in the wild-type CP and RT protein and was transmitted by aphids to plants, where it replicated and spread. However, this RTD mutant could not be further transmitted by the vector, which demonstrated the requirement of the RT protein in the transmission process. Finally, work with PEMV-1 has shown that the presence of the RTD is mandatory for aphid transmission in this system as well [20]. a. The Variable Region of the RTD, As noted above, purified wild-type luteoviruses typically lack the variable portion of the RTD. Purified virus can nonetheless still be acquired and transmitted by aphids, which provides a first indication that the variable region contains no indispensable transmission determinants. McGrath et al [47] used an immunoblocking method to map the capsid domains important in the transmission of CYDV-RPV by its vector, R. padi. Their findings also indicate that the C-terminal part of the RT protein is not necessary for transmission. On the other hand, sequence comparison of two PLRV isolates that differ in their efficiency of transmission by M persicae identified three amino acid changes in the C-terminal part of the RTD as being critical [48]. Thus, even if the C-terminal variable region of the polerovirus RTD is not strictly necessary, signals governing the efficiency of aphid transmission could be located in this region. In the case of PEMV-1, reciprocal exchanges were created between aphidtransmissible and aphid-nontransmissible isolates in an effort to identify the regions of these RNAs that are critical for aphid transmission. A single nucleotide substitution located in the truncated variable domain of the RTD rendered the aphid-transmissible parental isolates aphid-nontransmissible [20]. It should be noted that for PEM V-1, there is no evidence of processing of the RT protein during virus purification. In this case, it can be assumed that the variable domain of the RT protein is present in the PEMV-1 particle. In order to characterize more precisely the sequences in the BWYV RTD involved in the transmission process, small deletions and point mutations have been introduced into the infectious clone. These mutants are shown in Figure 4. In mutant BW6.1, a frameshift (+4 nucleotides) was introduced 20 amino acids downstream of the conserved region. The results obtained with this mutant have been fully described by Brault et al [18]. Three deletion mutants were also created: BW6.AE1, BW6.40, and BW6.41 have 85, 128, and 79 amino acids, respectively, eliminated from the variable region. Finally, BW6.MHD is a point mutant in which four conserved amino acids of the MHD sequence were replaced by alanine residues. Bruyere et al. [49] analyzed these mutants and the effects of their deletions or point mutations on virus accumulation and aphid transmission. They found that BW6.MHD behaved like the wild-type virus, whereas plants agroin-
11.
VIRAL DETERMINANTS INVOLVED IN LUTEOVIRUS-APHID INTERACTIONS
219
fected with BW6.AE1, BW6.40, and BW6.41 accumulated only half to a third as much virus as did the wild-type. Accumulation of mutant BW6.1 was initially very low, but virus titers increased dramatically 5 to 7 weeks after agroinfection to approach wild-type levels. An RT protein of the expected size was detected in plants agroinfected with the different mutants, except for BW6.1, in which no truncated RT protein could be detected early after agroinfection. Later on, however, full-length RT protein could be detected in the agroinfected plants. Analysis of progeny virus at different times in plants agroinfected with BW6.1 revealed that the high levels of virus accumulation occurring at later stages of infection were associated with pseudoreversion (deletion of a single nucleotide) that restored the original reading frame. The pseudoreversions resulted in local changes in the amino acid sequence in the progeny, illustrating that the RTD can tolerate minor sequence alterations and still remain functional. Purified virions were prepared from plants agroinfected with the above mutants, and the protein content of the viral particle was examined by Western blot analysis. In addition to the major CP, P74* was present in levels comparable with those in the wild-type virus in mutants BW6.AE1, BW6.40, BW6.41, and BW6.MHD. When BW6.1 was purified at early stages after agroinfection (before the pseudoreverted forms has become abundant), only low amounts of P74* were associated with the viral particle. Possibly, the truncated RT protein resulting from the frameshift is less stable in planta, is not efficiently incorporated into the viral particle, or both. When agroinfected plants were used as viral sources for aphids, transmission rarely occurred, even at high inoculum pressure, for any of the variable-domain mutants except mutant BW6.MHD. The latter was transmitted by aphids as efficiently as the wild-type virus (Table 1). The significance, if any, of the so-called M persicae homology domain thus remains to be discovered. When purified viruses of BW6.AE1, BW6.40, and BW6.41 were membrane-fed to aphids, transmission occurred at a rate identical to that observed with wild-type virus. The only exception was mutant BW6. 1 purified at early intervals after agroinfection, for which only rare transmission occurred, even at a high inoculum pressure (Table 1). Nonviruliferous nymphs that are hemocoelically injected with purified BW6.1 still only transmit the mutant at a low rate. Progeny analysis of the few plants that are aphid-infected by BW6.1 in this way reveals that successftxl transmission is invariably associated with the appearance of the aforesaid pseudorevertants. The low transmissibility of this mutant may well be related to a low level of incorporation of the truncated form of the RT protein into virions rather than to lack of a transmission signdX per se. With the three deletion mutants BW6. AEl, BW6.40, and BW6.41, there was a pronounced difference in the transmission efficiency depending on whether the virus source was agroinfected plants or purified virus [49]. The poor transmissibility of these deletion mutants from leaves may be related to their lower concentration in the plants compared with wild-type virus (see above), to an altered distribution of virus in the plant that reduces virus accessibility to aphids, or to
220
VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K.E. RICHARDS
both. Alternatively, the low transmissibility of the variable domain mutants could be related to alterations in the three-dimensional structure of the RTD that hamper recognition by aphid receptor(s) of sites located elsewhere on the RTD (e.g., the conserved region), CP, or both. Perhaps the large deletion in mutant BW6.40 provokes less misfolding of the altered RT protein than do the smaller deletions in BW6.AE1 and BW6.41. This could account for the fact that BW6.40 is more efficiently transmitted than the other mutants (Table 1). b. The Conserved Region of the RTD, In an attempt to better characterize aphid transmission determinants in the conserved part of the RT protein, a set of point mutations and deletions in this region was produced (Fig. 4). Point mutations were generated at or near five positions that are conserved in all sequenced poleroviruses [50]. The mutations were mostly alanine substitutions and were generally targeted to amino acids with charged side chains, as such residues are more likely to be exposed on the surface of the native protein. The deletions in BW6.106, BW6.104, BW6.AMT, and BW6.ATB eliminated 6, 15, 15, and 33 amino acids, respectively, from the conserved region [49]. In mutant BW6.50, the deletion of 33 amino acids removed a sequence overlapping the boundary between the conserved and variable regions of the RTD. In BW6.51, the deletion eliminated the proline tract and three amino acids immediately downstream. Cell-free translation experiments have shown that sequences in the corresponding locations of ORF 5 of BYDV-PAV regulate in cis the rate of translational readthrough of the major CP termination codon [51]. Similar translation experiments with a BWYV readthough protein transcript containing the BW6.50 and BW6.51 mutations indicated that synthesis of the RT protein but not the CP was reduced as compared with the wild-type construct [44]. To determine if the deletions in BW6.50 and BW6.51 have a similar effect in vivo, the relative amount of major CP and RT protein produced during protoplast infections was estimated. For both mutants, the rate of RT protein synthesis was diminished at least ten-fold, suggesting that cw-acting readthrough enhancer elements are similarly positioned in the BWYV RTD [49]. None of the other mutations introduced into the RTD conserved region significantly lowered the efficiency of readthrough translation. In agroinfected plants, progeny virus accumulation for the various conserveddomain deletion mutants was always very low compared with wild-type virus. The ELISA values in such plants were lower by a factor of at least 10 than in plants agroinfected with the wild-type construct but similar to those in plants agroinfected with BW6.4 in which the RTD has been completely eliminated [49]. Progeny of BW5.127 also accumulated poorly early after agroinoculation but attained wild-type levels later on. Sequence analysis of the progeny showed conservation of the original mutation at 4 weeks postinoculation, but later (7 weeks postinoculation) the original mutation had undergone modifications (Fig. 5). When assayed by ELISA, the other point mutations had no effect on the accumulation of viral progeny in planta. The original mutations were maintained in mutants BW5.121 and BW5.123, but surpris-
11.
221
VIRAL DETERMINANTS INVOLVED IN LUTEOVIRUS-APHID INTERACTIONS
Secondary mutants in agroinfected A'^ clevelandii
Original mutant
_^^_Aphid Transniission
J
+._ti
Viral progeny in M perfoliata
Viral progeny in M. perfoliata .,.„.„..,.„.,.,,.„.,.,
Aphid TrMismission
li
± -^
J-*- . i
(59-60)
u.
BW5.123L{'
i
(200-201)
^ xmchanged primary mutation
WT transmission efficiencies
Y modified primary mutation • 0 mutations in the PVT patch
c^^#
Fig, 5 Distribution of the major mutations in the viral progeny of point-mutants BW5.121, BW5.123, and BW5.127 following agroinfection and aphid transmission. For each mutant, the horizontal line indicates the portion of the genome at which the primary mutation was introduced. Numbers in brackets indicate the position of the primary mutation along the amino acid sequence of the readthrough domain (RTD). Arrows represent aphid transmission after acquisition of virus by membrane feeding (black arrows) and microinjection (gray arrows). The thickness of the arrow is proportional to the rate of transmission. The gut epithelium and the two barriers of the accessory salivary gland (basal lamina and basal plasmalemma) are represented by separate vertical lines.
ingly, occasional second-site mutations appeared in an amino acid triplet prolinevaline-threonine (PVT), the "PVT patch." The PVT patch is located 7 amino acids downstream of the BW5.121 mutation and 26 amino acids upstream of the BW5.123 mutation (Fig. 5). In the second-site mutations, either the proline residue was replaced by leucine or the threonine residue was replaced by isoleucine. The possible significance of the PVT patch will be discussed below. Mutants BW5.125 and BW5.129 retained the primary mutations and no other modifications were noted. For the conserved-domain mutants, RT protein of the expected size was reproducibly detectable in crude protein extracts from all agroinfected plants except those infected with mutants BW6.50 and BW6.51. Deletion of sequences required for the readthrough phenomenon (see above) in these two mutants is probably responsible for the low amount of RT protein produced in these plants. For the other deletions in the conserved region, the RT protein accumulated to readily detectable levels in agroinfected plants, although those levels were somewhat lower than in plants agroinfected with wild-type virus.
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VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS
Vims could be purified from the plants agroinfected with the conserved-region deletion mutants [49], although the yields were low. Analysis of the protein content of purified virions revealed that the deletions in the conserved region interfered with the packaging of the RT protein into virions. Thus, no truncated RT protein (P74*) was detectable in virions of BW6.51, BW6.106, BW6.104, BW6.AMT, or BW6.ATB; P74*, was present but dramatically reduced for BW6.50. In the case of BW6.51 and BW6.50, poor incorporation of RT protein into virions was expected in view of the effect of these mutations on RT protein synthesis. The lack of RT protein incorporation into BW6.106, BW6.104, BW6.AMT, and BW6.ATB virions cannot, however, be explained in this manner. Rather, it appears that the latter deletion mutants have altered the structure of the RT protein in such a way that the deleted versions are packaging-deficient or are diverted away from sites of virus assembly, for example, into aggregates. In contrast to deletion mutants, none of the point mutants in the conserved region significantly interfered with incorporation of RT protein into virions [50]. When young, fully expanded leaves of agroinfected plants or purified virions were used as the viral source in aphid transmission tests, none of the conservedregion deletion mutants were transmitted, even at high inoculum pressure (Table 2). In contrast, all point mutants were transmitted at various rates from both inoculum sources [50]. Aphid transmission was very efficient for BW5.125 and BW5.129, whereas mutants BW5.121 and BW5.127 were transmitted at an intermediate efficiency and BW5.123 was transmitted at a very low rate, even when a high inoculum pressure was used (Table 2). Low accumulation in the agroinfected plants of the conserved-domain deletion mutants discussed above cannot account entirely for their nontransmissibility from leaves by M persicae. The mutant BW5.127 accumulated soon after agroinfection of plants to levels comparable with those for the deletion mutants, and it is reasonably well transmitted at high inoculum pressure. Instead, the absence (or near absence in the case of mutant BW6.50) of the RT protein from the virions is sufficient to account for the failure of conserved-region deletion mutants to be aphid-transmissible. However, it is quite possible that the deletions have also removed sequence motifs that are essential for transmission per se. In contrast, since the point mutants incorporated normal amounts of RT proteins into virions, the poor transmission of these mutants can be directly attributed to the introduced mutations. Microinjection of purified BW5.121, BW5.123, and BW5.127 into the hemocoel of nonviruliferous nymphs yielded transmission efficiencies similar to those for wild-type virus. The latter results suggest that these particular RTD mutations interfere with the initial step of transmission, that is, movement of virions through the intestinal cell barrier [50]. In the viral progeny analyzed following aphid infection with BW5.121, secondsite mutations were almost always present in the PVT patch mentioned above (Fig. 5). This finding suggests that strong selective pressure for appearance of the PVT patch mutations is exerted during aphid transmission of BW5.121. To verify
11.
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223
Table II Aphid Transmission of Beet Western Yellows Virus (BWYV) Mutants in the Conserved Domain of the Readthrough (RT) Protein" Aphid transmission from agro-infected plants^ BWYV mutants BW6.4 BW6.51 BW6.106 BW6.104 BW6.AMT BW6.ATB BW6.50 BW5.121 BW5.123 BW5.125 BW5.127 BW5.129 BW-WT
RT incorporation into virions
— — — — .— —
+ +++ +++ +++ +++ +++ +++
8 Aphids/pl. 0% 0% 0% 0% nd^ nd 0% nd nd 100% nd 100% 90%
30 Aphids/pl. 0% 0% 0% 0% 0% 0% 0% 71%^ 13%^ 100% 50%^ 100% 98%
Aphid transmission from purified viruses^ 8 Aphids/pl. 0% 0% nd nd nd 0% 0% nd nd nd nd 100% 97%
30 Aphids/pl. 0% 0% 0% 0% 0% 0% 0% 35%^ 6%^ nd 30%^ 100% 99%
" Results are presented as the percentage of infected plants/plants tested. ^ The virus source was N. clevelandii leaves 3 to 6 weeks after agro-inoculation. After a 24-h-virus acquisition access period, aphids (8 or 30) were transferred to test plant for an inoculation feeding period of 4 days. ^ Purified virus at 25 |ig/ml was supplied to aphids by membrane-feeding. ^ nd = not determined. ^ Transmission events associated with secondary mutants or pseudo-revertants.
this hypothesis, one of the PVT patch mutations was introduced directly into BW5.121. The resulting mutant (BW5.121L) was readily transmitted from agroinfected leaves, and both mutations introduced into the construct were maintained in progeny virions (Fig. 5). Thus, the engineered second-site mutation favors aphid transmission of the virus in the BW5.121 background. The nature of the viral progeny following aphid transmission of BW5.123 depended on the viral acquisition method. When purified virions were acquired by membrane feeding, the viral progeny carried a modification in the original mutation, which fully restored transmissibility from aphid-infected plants. When purified virions were microinjected into the hemocoel, on the other hand, the primary mutation was conserved and second-site mutations appeared in the PVT patch (Fig. 5). The fact that PVT patch mutations were only observed in plants infected by microinjected aphids indicates that, at least in the BW5.123 background, the sequence signals permitting virus movement through the ASG are distinct from those that permit transit from the intestine into the hemocoel. To test this point directly, a new mutant.
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VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS
BW5.123L, carrying one of the observed second-site mutations in the PVT patch was created. This mutant was not transmitted when acquired from agroinfected plants but was efficiently transmitted when delivered to aphids by microinjection. We conclude that the modifications observed in the primary mutation permit movement of the virus across both the intestine and ASG barriers but that in the BW5.123 background, the PVT patch operates at the second barrier. The mechanism by which PVT-patch second-site mutations restore transmissibility of BW5.121 and BW5.123 is not known. Possibly, the folding of the RTD brings the sites targeted by the primary mutations into proximity, and the same second-site mutations somehow correct misordering of this region brought about by the primary mutations. An alternate possibility is that the PVT-patch mutations act as "global stabilizers" [52]. Global stabilizers are mutations that increase the overall stability of a protein and can counteract, in a nonspecific manner, the action of other mutations that interfere with the proper and stable folding of the polypeptide. A somewhat similar type of secondary mutation has been characterized in the progeny of a poliovirus mutant, which had adapted to growth on cells expressing mutant forms of poliovirus receptor. The secondary mutation has been suggested to modulate the flexibility of the capsid to allow it to accommodate the mutant receptor [53]. Further experimentation, ultimately including structural studies on the luteovirus capsid and identification of receptors, will no doubt be required to distinguish among these possibilities. For mutant BW5.127, none of the progeny clones analyzed after aphid transmission retained the primary mutation; instead, progeny exhibited various pseudoreversions (Fig. 5). The most common of these modifications was introduced into BW5.127 to create BW5.127F. This mutant accumulated to near wildtype levels in plants and was efficiently transmitted by the vector. This shows that the modification observed in the primary mutation is beneficial for viral accumulation in plants and for virus transmission in the BW5.127 context [50]. Mutant BW5.129 was readily transmitted when supplied to aphids as agroinfected leaves or virions purified from such leaves, but was poorly transmitted when the virus was supplied as an extract of infected protoplasts. We expected that this difference in transmissibility would be due to significant amounts of revertant or compensatory second-site mutations or both during the relatively long period (4-6 weeks) of an agroinfection experiment. Surprisingly, however, the viral progeny of BW5.129 analyzed in aphid-infected plants was highly homogenous in the sequenced region; the original mutations were uniformly conserved, and no obvious second-site mutations were present. Thus, the high transmissibility of this mutant from plants as compared with protoplasts cannot be explained by the presence of compensatory mutations near the primary mutation site. Possibly, a compensatory mutation or mutations are present elsewhere in the genome, but ftirther experiments will be necessary to confirm this hypothesis. Finally, for BW5.125, the original mutation was conserved in all the clones analyzed, and we conclude that the motif targeted in this particular mutant is not necessary for transmission [50].
11.
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225
To sum up, the results obtained with the conserved-domain deletion mutants suggest that packaging of the RT protein in the particle is essential for the virus to be transmitted. Most of the deletions introduced in the conserved region inhibited incorporation of the RT protein in the particle, and none of these mutants were aphid-transmissible. Mutants BW6.1 and BW6.50, which incorporated very low amounts of RT protein into particles, were also transmission-defective. It should be mentioned, however, that the preceding remarks do not rule out the possibility that the regions eliminated by the various deletions may contain motifs important for transmission in their own right. Nevertheless, it is entirely possible that the viral receptors in the aphid that recognize the virus must interact with several copies of the RT protein concurrently to start the process of endocytosis. Thus, mutations that lower the efficiency of RT protein synthesis or its incorporation into virions or both could inhibit aphid transmission independently of a direct effect of the mutation on a putative aphid recognition motif. Point mutations were originally introduced in the conserved region of the RTD in an attempt to uncouple possible effects of RT protein synthesis and incorporation into virions from effects on aphid transmission. This proved to be possible for mutants BW5.121, BW5.123, and BW5.127, which incorporate normal amounts of RT protein into virions and accumulated to high titers inplanta but are inefficiently transmitted by M. persicae when acquired from agroinfected plants or from purified virions [50]. The identification of precise RTD sequences as frmctional motifs in the different steps of the transmission circuit is rendered difficult by the appearance of secondary mutations or pseudoreversions in viral progeny. This observation suggests that there is a considerable structural redundancy or flexibility in the RTD and that this protein can tolerate modifications even at highly conserved residues. Overall, however, we can conclude that (1) an exact RTD sequence is not necessary for transmission, because at least one mutant (BW5.125) was without effect; (2) the RTD appears to have considerable functional plasticity, because mutants such as BW5.121, BW5.123 and BW5.127, which are apparently defective in virus transmission, are subject to compensatory mutations that can reverse this effect; and (3) the transport of virus across the gut barrier and through the basal lamina and plasmalemma of the ASG involves distinct RTD motifs. This is illustrated by the behavior of the double mutant BW5.123L, which seems unable to cross the gut epithelium but can be transported efficiently through the ASG.
IV. Virus-Symbionin Interactions An unexpected turn in the story of luteovirus-aphid interactions has been the discovery that, in addition to the virus and the aphid, a third organism is involved in virus transmission [54]. Aphids contain gram-negative endosymbiotic bacteria of the genus Buchnera in specialized vesicles called mycetocytes in the hemocoel
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VERONIQUE BRAULT, VERONIQUE ZIEGLER-GRAFF, AND K . E . RICHARDS
[55]. In an attempt to detect luteovirus receptor proteins of the aphid, van den Heuvel et ah [26] carried out virus binding assays in which blots of aphid proteins (which had been separated by two-dimensional polyacrylamide gel electrophoresis) were allowed to interact with potato leafroU virus (PLRV) in an overlay solution. The most important protein that displayed affinity for the virus proved to be symbionin [56, 57], a GroEL-like chaperonin produced by the primary endosymbiont of the aphid. Immune electron microscopy revealed that symbionin was not only present in the endosymbionts but also in free form in the hemocoel [26]. Antibiotic treatment of aphids dramatically lowered symbionin levels in the hemolymph, and this was accompanied by a decrease in the efficiency of virus transmission. This correlation suggests that interaction between symbionin and the luteovirus stabilizes the virus in the hemolymph or affects transmission in some other way [26]. In vitro interactions have also been detected between BYDV-PAV and symbionins from S. avenae and R. padi [58]. Finally, symbionin has recently been implicated in persistence of tomato yellow leaf curl geminivirus within the hemolymph of the whitefly Bemisia tabaci [59]. The available data suggest that the conserved portion of the RTD is the principal determinant on the viral capsid that interacts with symbionin. Thus, purified virions of BWYV mutants that do not incorporate the RTD, such as BW6.4 and BW6.50 (Fig. 4), do not bind symbionin, whereas mutants such as BW6.AE1 (Fig. 4) with a deletion in the variable region bind symbionin with affinity comparable to that observed with wild-type virus [60]. Furthermore, microinjection experiments with the aforesaid mutants revealed that BW6.4 and BW6.50 virions are rapidly cleared from the hemocoel of M persicae, whereas BW6.AE1 and wild-type virions persist [60]. Even though the evidence is strong for a role of symbionin in luteovirus persistance, it is important to note that symbionin-luteovirus interactions cannot account for the specificity of luteovirus transmission, because luteoviruses bind with comparable affinity to symbionin from both vector and nonvector aphid species [26, 60]. It appears, rather, that symbionin acts in a nonspecific manner to facilitate the transmission process. The exact mechanism by which this takes place remains unclear. Among the possiblities (which are not mutually exclusive) are (1) symbionin protects the virus from attack by proteases or recognition by the aphid immune system; (2) symbionin prevents aggregation or disassembly of the virus in the hemocoel; and (3) symbionin participates in passage of the virus into the ASG by "presenting" putative receptor attachment sites on the RTD in proper conformation.
V. Concluding Remarks Although study of vector recognition motifs on the major viral CP is still in its early stages, there is now evidence of a role for the minor capsid protein RTD in
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efficient transmission of two luteoviruses, BWYV and BYDV-PAV Thus, Chay et al [21] reported that an RTD deletion mutant of BYDV-PAV could be acquired into the hemocoel but was not aphid-transmissible. Recent electron microscopic observations made using BWYV mutant BW6.4 showed viral particles, mostly in aggregates, between the basal lamina and basal plasmalemma of the gut cells (C. Reinbold, personal communication). This suggests that acquisition of both BYDV-PAV and BWYV particles is independent of the RTD and that the transmission step or steps requiring RTD are postacquisition events. Virus uptake and transport through the ASG may be the RTD-dependent step. Moreover, through its interaction with symbionin, the RTD probably also plays a role in protecting virus from degradation in the hemocoel. So far, analysis of RTD mutants has not allowed us to draw unequivocal correlations between RTD sequences and various functions in transmission. It appears that RT protein in both full-length and truncated (P74*) forms can be recognized by aphid receptors and promote virus transmission by the vector. Whereas the conserved part of the RTD is crucial for transmission, the variable domain, although not strictly essential, may nevertheless modulate transmission efficiency, especially if mutations introduced in this domain significantly modify the secondary or tertiary structure of the RT protein. These structural modifications could mask important determinants on the conserved portion of the RTD or on the CP involved in binding virus with aphid receptors. Furthermore, aphid receptors involved in virus endocytosis might in fact be multiple receptors, particularly those at the apical plasmalemma of gut cells, where virus particles must be captured from nutriment. Thus, a primary receptor may be responsible for capturing virions while a second receptor intervenes in the slower process of endocytosis [61]. A striking feature of our observations is that most of the point mutations scattered along the conserved region of the RTD have an effect on the transmission rate but none completely eliminate transmission. In part, this is a consequence of the high mutability of the luteovirus RNA genome, which allows second-site gainof-function mutations to be selected. The ability of second-site mutations in the RTD to restore function indicates that there is a certain amount of flexibility built into the structure. Our observations are also consistent with the idea that complex secondary or tertiary structural features of the capsid rather than simple linear sequence motifs are involved in virus recognition by aphid receptors. For poliovirus type 1 and rhinovirus type 14, crystallographic studies have revealed a conserved cleft in the virion, called the canyon, which extends from the surface of the virion to the inner region of the capsid [62, 63]. This cleft has been proposed to be the site on the virion that binds to the cell receptor [62]. Possibly, viral recognition sites for aphid receptors will be associated with a similar type of structure. Even though the weight of the evidence clearly favors a role for the RTD in aphid transmission of BWYV, recent experiments with PLRV [23] appear to tell a different story. Viruslike particles (VLP) of PLRV were produced in a baculovirus system by expressing the CP gene sequence modified by the addition of a histi-
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dine tag at the 5' end [64]. Ultrastructural studies reveal that the virusHke particles, which lack the RT protein, are able to enter the hemocoel of the vector and traverse the ASG to the lumina of the salivary canals and salivary duct [65]. Transmission of viruslike particles to plants could not be verified since these particles, which do not incorporate viral RNA, are noninfectious. At present we have no good explanation for the apparent contradiction between the above findings and the observations with BWYV Possibly PLRV viruslike particles behave differently from "normal" virions during their passage through the aphid, perhaps because of the additional N-terminal sequence. On the other hand, the aphid transmission experiments with BWYV have generally been limited to characterization of the starting material, virus in the agroinfected plant, and the final resulting transmission to a test plant. This "black box" approach could lead to serious misinterpretations of events within the aphid. Therefore, future experiments will be aimed at tracing the passage of various BWYV RTD mutants through the aphid by reverse transcriptase-polymerase chain reaction testing of the hemocoel contents and by ultrastructural techniques.
Acknowledgments We thank our colleagues, particularly Etienne Herrbach, for their comments on the manuscript.
References 1. Mayo, M.A., and Ziegler-Graff, V (1996). Molecular biology of luteoviruses. Adv. Virus Res. 46, 416-460. 2. Miller, W.A., and Rasochova, L. (1997). Barley yellow dwarf viruses. Annu. Rev. Phytopathol 35, 167-190. 3. Harrison, B.D. (1999). Steps in the development of luteovirology. In "The Luteoviridae" (H.G. Smith and H. Barker, eds.), pp. 1-14. CAB International, Oxford, U.K. 4. Rochow, W.F. (1961). The barley yellow dwarf virus disease of small grain. Adv. Agron. 13, 217-248. 5. Duffus, J.E. (1977). Aphids, viruses and the yellow plague. In "Aphids as Virus Vectors" (K.F. Harris and K. Maramorosch, eds.), pp. 361-383, Academic Press. New York. 6. Rochow, W.F., and Duffus, J.E. (1981). Luteoviruses and yellows diseases. In "Handbook of Plant Virus Infections and Comparative Diagnosis" (E. Kurstak, ed.), pp. 147-170, Elsevier/North Holland Biomedical Press, Amsterdam. 7. Mayo, M.A., and D'Arcy, C.J. (1999). Family Luteoviridae: A reclassification of luteoviruses. In "The Luteoviridae" (H.G. Smith and H. Barker, eds.), pp. 15-22, CAB International, Oxford, U.K. 8. Van der Wilk, R, Verbeek, M., Dullemans, A.M., and van den Heuvel, J.FJ.M. (1997). The genome-linked protein of potato leafroll virus is located downstream of the putative protease domain of the ORFl product. Virology 234, 300-303.
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9. Wobus, C.E., Skaf, J., S., Schultz, M.H., and de Zoeten G.A. (1998) Sequencing, genomic localization and initial characterization of the VPg of pea enation mosaic enamovirus. J. Gen. Virol. 79, 2023-2025. 10. Veidt, I., Lot, H., Leiser, M., Scheidecker, D., Guilley, H., Richards, K., and Jonard, G. (1988). Nucleotide sequence of beet western yellows virus RNA. Nud. Acids Res. 16, 9917-9932. 11. Martin, R.R., Keese, RK., Young, M.J., Waterhouse, RM., and Gerlach, W.L. (1990). Evolution and molecular biology of luteoviruses. Annu. Rev. Phytopathol. 28, 341-363. 12. Tacke, E., Prufer, D., Salamini, R, and Rohde, W. (1990). Characterization of a potato leafroll luteovirus subgenomic RNA: Differential expression by internal translation initiation and AUG suppression. J. Gen. Virol. 71,116S-1211. 13. Dinesh-Kumar, S.R, Brault V, and Miller W.A. . (1992). Precise mapping and in vitro translation of a trifiinctional subgenomic RNA of barley yellow dwarf virus. Virology 18, 711-722. 14. Bahner, I., Lamb, J., Mayo, M.A., and Hay, R.T. (1990) Expression of the genome of potato leafroll virus: Readthrough of the coat protein termination codon in vivo. J. Gen. Virol. 71, 2251-2256. 15. Vincent, J.R., Lister, R.M., and Larkins, B.A. (1991). Nucleotide sequence analysis and genomic organization of the NY-RPV isolate of barley yellow dwarf virus. J. Gen. Virol. 72, 2347-2355. 16. Reutenauer, A., Ziegler-Graff, V, Lot, H., Scheidecker, D., Guilley, H., Richards, K., and Jonard, G. (1993). Identification of beet western yellows luteovirus genes implicated in viral replication and particle morphogenesis. Virology 195, 692-699. 17. Filichkin, S.A., Lister, R.M., McGrath, PR, and Young, M.J. (1994). In vivo expression and mutational analysis of the barley yellow dwarf virus readthrough gene. Virology 205,290-299. 18. Brault, V, van den Heuvel, J.EJ.M., Verbeek, M., Ziegler-Graff, V, Reutenauer, A., Herrbach, E., Garaud, J.C., Guilley, H., Richards, K., and Jonard, G. (1995). Aphid transmission of beet western yellows virus requires the minor capsid read-through protein P74. EMBO J. 14, 650-659. 19. Wang, J.Y., Chay C , Gildow RE., and Gray S.M. (1995). Readthrough protein associated with virions of barley yellow dwarf luteovirus and its potential role in regulating the efficiency of aphid transmission. Virology 206, 954-962. 20. Demler, S.A., Rucker-Feeney, D.G., Skaf, J.S., and de Zoeten, G.A. (1997). Expression and suppression of circulative aphid transmission in pea enation mosaic virus. J. Gen. Virol. 72, 511-523. 21. Chay, C.A., Gunasinghe U.B., Dinesh-Kumar S.P, Miller W.A., and Gray S.M. (1996). Aphid transmission and systemic plant infection determinants of barley yellow dwarf luteovirus-PAV are contained in the coat protein readthrough domain and 17 kDa protein, respectively. Virology 219, 57-65. 22. Schmitz, X, Stussi-Garaud, C , Tacke, E., Prufer D., Rohde W., and Rohfritsch O. (1997). In situ localisation of the putative movement protein (prl7) from potato leafroll luteovirus (PLRV) in infected and transgenic potato plants. Virology 235, 311-322. 23. Gildow, F. (1999). Vector-virus interactions. In "The Luteoviridae" (H.G. Smith and H. Barker, eds), pp. 88-112. CAB International, Oxford, U.K. 24. Garret, A., Kerlan, C , and Thomas, D. (1993). The intestine is a site of passage for potato leafroll virus from the gut lumen into the haemocoel in the aphid vector Myzus persicae Sulz. Arch. Virol. 131, 377-392. 25. Garret, A., Kerlan, C , and Thomas, D. (1996). Ultrastructural study of acquisition and retention of potato leafroll luteovirus in the alimentary canal of its aphid vector, Myzus persicae Sulz. Arch. Virol. 141, 1279-1292. 26. Van den Heuvel, J.F.J.M., M. Verbeek, and F. van der Wilk. (1994). Endosymbiotic bacteria associated with circulative transmission of potato leafroll virus by Myzus persicae. J. Gen. Virol. 75, 2559-2565.
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27. Rochow, W.F., and Pang, E. (1961). Aphids can acquire strains of barley yellow dwarf vims they do not transmit. Virology 15, 382-384. 28. Rochow, W.F. (1969) Biological properties of four isolates of barley yellow dwarf virus. Phytopathology 5% 1580-1589. 29. Gildow, RE. (1993). Evidence for receptor-mediated endocytosis regulating luteovirus acquisition by aphids. Phytopathology 83, 270-277. 30. Gildow, F.E., and Gray S.M. (1993). The aphid salivary gland basal lamina as a selective barrier associated with vector-specific transmission of barley yellow dwarf luteovirus. Phytopathology 83,1293-1302. 31. Peififer, M.L., Gildow, F.E., and Gray, S.M. (1997). Two distinct mechanisms regulate luteovirus transmission efficiency and specificity at the aphid salivary gland. J. Gen. Virol. 78,495-503. 32. Yurchenco, P.D., and Schittny, J.C. (1990). Molecular architecture of basement membranes. FASEB J. 4, \511-1590. 33. Pedersen, K.J. (1991). Structure and composition of basement membranes and other basal lamina matrix systems in selected invertebrates. Acta Zool. 72, 181-201. 34. Veidt, I., Bouzoubaa, S.E., Ziegler-Graff, V, Leiser, R.M., Guilley, H., Jonard, G., and Richards, K. (1992). Synthesis of full-length transcripts of beet western yellows virus RNA: Messenger properties and biological activity in protoplasts. Virology 186, 192-200. 35. Mohan, B.R., Dinesh-Kumar, S.P., and Miller, W.A. (1995) Genes and cis-acting sequences involved in replication of barley yellow dwarf virus-PAV RNA. Virology 111, 186-195. 36. Prufer, D., Wipf-Scheibel, C , Richards, K., Guilley, H., Lecoq, H., and Jonard, G. (1995). Synthesis of a full-length infectious clone of cucurbit aphid-borne yellows virus and its use in gene exchange experiments with structural proteins from other luteoviruses. Virology 214, 150-158. 37. Grimsley, N.H. (1990) Agroinfection. Physiol. Plant 79, 147-153. 38. Leiser, R.M., Ziegler-Graff V, Reutenauer A., Herrbach E., Lemaire O., Guilley H., Richards K., and Jonard G. (1992). Agroinfection as an alternative to insects for infecting plants with beet western yellows virus. Proc. Natl. Acad. Sci. U.S.A. 89, 9136-9140. 39. Commandeur, U, and Martin, R. (1993). Investigations into the molecular biology of potato leafroll luteovirus by means of agroinfection. (Abstr.) Phytopathology 83, 1426. 40. Domingo, E., and Holland J.J. (1997). RNA virus mutations and fitness for survival. Annu. Rev. Microbiol. 51, 151-178. 41. Ziegler-Graff, V, Brauh V, Mutterer J.D., Simonis M.T, Herrbach E., Guilley H., Richards K.E., and Jonard G. (1996). The coat protein of beet western yellows luteovirus is essential for systemic infection but the viral gene products P29 and PI9 are dispensable for systemic infection and aphid transmission. Mol. Plant-Microbe Interact. 9, 501-510. 42. Esau, K., and Hoefert, L.L. (1972). Development of infection with beet western yellows luteovirus in the sugarbeet. Virology 48, 724-738. 43. D'Arcy, C.J., and de Zoeten G.A. (1979). Beet western yellows virus in phloem tissue of Thlaspi arvenae. Phytopathology 69, 1194-1198. 44. Mutterer, J.D. (1998). Etude des determinants viraux impliques dans le mouvement et la transmission du virus des jaunisses occidentals de la betterave ou BWYV Ph.D. dissertation, Universite Louis Pasteur, Strasbourg, France. 45. Mutterer, ID., Stussi-Garaud C , Michler P, Richards K.E., Jonard G., and Ziegler-Graff V (1999). Role of the beet western yellows virus readthrough protein in viral movement in Nicotiana clevelandii. J. Gen. Virol. 80, 211\-111%. 46. Guilley H., Wipf-Scheibel, C , Richards, K., Lecoq, H., and Jonard, G. (1994). Nucleotide sequence of cucurbit aphid-borne yellows luteovirus. Virology 202, 1012-1017.
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47. McGrath, RE, Lister, R.M., and Hunter, B.G. (1996). A domain of the readthrough protein of barley yellow dwarf virus (NY-RPV isolate) is essential for aphid transmission. Eur J. Plant Pathol. 102, 671-679. 48. Jolly, C.A., and Mayo, M.A. (1994). Changes in the amino acid sequence of the coat protein readthrough domain of potato leafroU luteovirus affect the formation of an epitope and aphid transmission. Virology 201, 182-185. 49. Bruyere, A., Brauk V, Ziegler-GraflfV, Simonis M.T., van den Heuvel J.F.J.M., Richards K., Guilley H., Jonard G., and Herrbach E. (1997). Effects of mutation in the beet western yellows virus readthrough protein on its expression and packaging, and on virus accumulation, symptoms and aphid transmission. Virology 230, 323-334. 50. Brauh, V, Mutterer, J., Scheidecker, D., Simonis, M.T., Herrbach, E., Richards, K., and ZieglerGrafif, V (2000). Effects of point mutations in the readthrough domain of the beet western yellows virus minor capsid protein on virus accumulation in planta and aphid transmission. J. Virol. 74, 1140-1148. 51. Brown, CM., Dinesh-Kumar S.P., and Miller W.A. (1996). Local and distant sequences are required for efficient readthrough of the barley yellow dwarf PAV coat protein gene stop codon. J. Virol. 70, 5884-5892. 52. Matthews, B.W. 1995. Studies on protein stability with T4 lysozyme. Adv. Protein Chem. 46, 249-278. 53. Racaniello, VR. (1996). Early events in poliovirus infection: Virus-receptor interactions. Proc. Natl. Acad. Sci. U.S.A. 93, 11378-11381. 54. Van den Heuvel, J.F.J.M., Hogenhout, S.A., and van der Wilk, F. (1999). Recognition and receptors in virus transmission by arthropods. Trends Microbiol. 7, 71-76. 55. Buchner, P. (1965). "Endosymbiosis of Animals with Plant Microorganisms." Interscience, New York. 56. Ishikawa, H. (1982). Host-symbiont interactions in protein synthesis in the pea aphid. Insect Biochem. 12, 613-622. 57. Hara, E., Fukatsu, T, Kakeda, K., Kengaku, M., Ohtaka, C , and Ishikawa, H. (1990). The predominant protein in an aphid endosymbiont is homologous to an E. coli heat shock protein. Symbiosis %, 27 \-2%?>. 58. Filichkin, S.A., Brumfield S., Filichkin T.P, and Young M.J.. (1997). In vitro interactions of the aphid endosymbiotic SymL chaperonin with barley yellow dwarf virus. J. Virol. 71, 569-577. 59. Morin, S., Ghanim, M., Zeidan, M., Czosnek, H., Verbeek, M., and van den Heuvel, J.F.J.M. (1999). A GroEL homologue from endosymbiotic bacteria of the whitefly Bemisia tabaci is implicated in the circulative transmission of tomato yellow leaf curl virus. Virology 256, 75-84. 60. Van den Heuvel, J.FJ.M., Bruyere, A., Hogenhout, S.A., Ziegler-Grafif, V, Brauh, V, Verbeek, M., van der Wilk, F, and Richards, K. (1997). The N-terminal region of the luteovirus readthrough domain determines virus binding to Buchnera GroEL, and is essential for virus persistence in the aphid. J. Virol 71, 7258-7265. 61. Haywood, A.M. (1994). Virus receptors: Binding, adhesion strengthening, and changes in viral structure. J. Virol. 68, 1-5. 62. Rossmann, M.G., Arnold, E., Erickson, J.W., Frankenberger, E.A., Griffith, J.P, Hecht, H.J., Johnson, J.E., and Kamer, G. (1985). Structure of a human common cold virus and functional relationship to other picomaviruses. Nature 317, 145-153. 63. Hogle, J.M., Chow, M., and Filmann, D.J. (1985). The three-dimensional structure of poliovirus at 2.9 A resolution. Science 229, 1358-1365.
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64. Lamb, J.W., Duncan, G.H., Reavy, B., Gildow, F.E., Mayo, M.A., and Hay, R.T. (1996). Assembly of virus-like particles in insect cells infected with a baculovirus containing a modified coat protein gene of potato leafroll virus. J. Gen. Virol. 11^ 1349-1358. 65. Gildow, R, Reavy, B., Mayo, M.A., Woodford, T. and Duncan, G. (1997). Potato leafroll virus-like particles lacking readthrough protein are transmitted by Myzus persicae (Abstr.) Phytopathology 87, S33.
CHAPTER 12
Approaches to Genetic Engineering of Potato for Resistance to Potato Leafroll Virus CHARLES R. BROWN ONEY P. SMITH
/- Introduction Potatoes are hosts for a variety of insect pests and plant pathogens. The most important pests of potatoes are aphids [1]. This is mainly attributable to damage from aphid-borne viral diseases but also to damage from direct feeding [1,2]. The major RNA viruses infecting potato are potato viruses A, M, S, X, and Y and potato leafroll virus (PLRV) [3]; the last two are the two most damaging viruses in terms of ubiquity of inoculum pressure and economic loss [4]. Potato leafroll virus is a member of the family Luteoviridae (genus Polerovirus) [5] with worldwide distribution and is transmitted by aphids in a circulative or persistent manner [6]. Oortwijn-Botjes [7] is credited in 1920 with discovering that PLRV is transmitted by the green peach aphid, Myzus persicae Sulz. There are now more than ten aphid species known to transmit PLRV [8]; Myzus persicae is considered the most efficient [6]. Restricted to the phloem, PLRV occurs mainly in sieve elements but also in companion cells and plasmodesmata connecting the two cell types [9]. Disruption and blockage of phloem is the most noticeable symptom in infected plants. Symptom expression differs markedly in potato varieties but usually includes mild to severe stunting and thickening of stems and leaves and an upward rolling of the older leaves due to starch accumulation [6,10]. Diseased plants have significantly reduced tuber yields [11, 12]. Current-season infection is usually less severe and damaging than secondary infection from virus-contaminated seed potato plantings [10]. Certain cultivars also develop an internal tuber disorder, net necrosis, Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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caused by death of tuber phloem cells [3, 13]. Tuber net necrosis develops during the growth period and in postharvest storage and is characterized by the appearance of dark internal vascular discoloration, which is pronounced at the stem end of the tuber. This discoloration causes serious problems for table stock and processor growers because the affected tubers do not pass grading standards and cannot be processed into French fries or potato chips. Net necrosis occurs in tubers from plants with primary, secondary, or tertiary infections [10]. Highly susceptible cultivars in the United States, such as Russet Burbank, can have high postharvest losses attributable to net necrosis. Field-resistant cultivars, such as Katahdin, are immune to this disorder. Potato leafroll virus in potato is best managed by developing clean seed stocks and producing certified seed potatoes with a disease incidence of less than 1%. Aphids require long phloem-feeding probes to acquire and inoculate PLRV Once acquired, virions require additional time, a latent period, to circulate to and through the accessory salivary glands in an infectious titer. These delays in the transmission cycle make vector control by insecticides a viable control measure. Probably PLRV would not survive in cropping systems that did not include potato. In the Columbia River basin of the Pacific Northwest, infected seed and diseased overwintering volunteers are reportedly the sole sources of PLRV [14]. Therefore, PLRV can be controlled by using hygienic seed production practices [15], provided virus in the volunteer potato reservoir is not allowed to spread to the current-season crop. Many seed programs rely on absolutely virus-free stock derived from tissue-cultured plants as a starting point for a "flush-through" system. Nonetheless, PLRV remains a serious problem on a worldwide basis, especially where hygienic seed programs are poorly monitored and controlled. Even in regions where well developed and managed seed programs are in force, PLRV outbreaks are serious in some years and produce residual infection problems lasting several years [16]. Enhancement of host resistance to PLRV could play a significant role in seed and commercial potato production. There are a number of breeding materials with usefiil levels of resistance to PLRV A hypersensitive reaction to the virus, resulting in plant death, has been studied and incorporated into certain varieties [17]. Resistance in cultivated materials includes resistance to infection as well as resistance to titer buildup [18-20]. The percentage of plants infected is significantly less in resistant clones than in susceptible ones, but the mean levels of infection are dependent on inoculation intensity. Clones possessing high levels of this relative resistance can be completely infected, given a sufficiently high infection pressure. Inheritance of relative resistance in advanced cultivated clones appears to be polygenic. A high level of resistance combined with superior horticultural traits, such as processing quality and high yield, is difficult to recover in breeding populations [21-23]. Extreme resistance, the inability to infect with intense inoculation pressure, including graft inoculation, has been noted in wild species or certain cultivated diploid breeding clones with wild ancestors [24, 25].
12.
GENETICALLY ENGINEERED POTATOES
0
1
235
2
ORF 0
3
4
5
6kb
ORF 2 I
t
I
ORF3
I
rir: ORF1
11—
ORF5
=^
ORF 4 Genomic RNA ^
Subgenomic RNA
Fig. 7 Genomic organization of potato leafroll virus (PLRV). The single-stranded RNA approximates 6 kilobases (kb) and is organized into at least six open reading frames (ORFs). The viral structural proteins include coat protein (ORF 3) and readthrough protein (ORF 5). The latter is a minor component of the virion and is expressed by readthrough translation of ORF 3. The viral nonstructural proteins include a movement protein (ORF 4) and replicase. The viral replicase is produced by ribosomal frameshifting between the 3' end of ORF 1 and the 5' end of ORF 2 to produce a fusion protein which functions in RNA replication [26]. ORFs 3, 4, and 5 are expressed by translation of a subgenomic RNA produced in virus-infected plants [27].
In heterozygous, clonally propagated crops, such as potato, the addition of traits during the breeding process cannot be accompanied by exact recovery of the recurrent phenotype. Transformation occupies a special place in potato breeding strategies because it can result in an exact replica of the original variety plus one or more new traits, an achievement that is not possible with sexual breeding. Consequently, the introduction of high-level PLRV resistance by gene transfer to an agronomically acceptable cultivar (e.g.. Russet Burbank) is an appealing solution to a formidable problem in potato cultivar development.
//,
The Virus
The genome of PLRV is a single-stranded, 6-kb, positive-sense RNA containing at least six open reading frames (ORFs) [26] (Fig. 1). Infected plants produce a 3' subgenomic RNA for the translation of downstream ORFs, including coat protein and the readthrough protein [26, 27]. The 23-kDa coat protein (CP) of PLRV is encoded by ORF 3, whereas readthrough protein is encoded by ORF 5. The latter, a minor component of PLRV particles, is expressed as a fusion protein by periodic readthrough translation of ORF 3 [28]. The product of ORFs 1 and 2, a fusion protein expressed by ribosomal frameshifting, functions as an RNA replicase [26]. The translational product of ORF 4 probably functions as a movement protein [29, 30]. Although no function has been attributed to ORF 0, roles in host recognition [26] and symptom expression [31] have been suggested. Two addi-
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tional ORFs, ORF 7 and ORF 8, have been reported for the 3' end of the genome, but their functions are not known [32].
///. Approaches to Pathogen-Derived Resistance Pathogen-derived resistance [33] is the expression of a pathogen's gene in a manner that interferes with the pathogen's Hfe cycle and disease expression. The wrong gene is expressed or a gene is expressed at the wrong time in the sequence of the pathogen's gene expression events. Pathogen-derived resistance is, therefore, the conceptual basis for inserting various ORFs of the PLRV genome into the potato genome for expression in planta and host-resistance production. A.
Coat Protein Strategy
The majority of published studies on transgenic resistance in potato involve potato transformed by insertion of the PLRV CP gene in its sense orientation. In all the studies discussed below, the viral CP cDNA was placed in a binary vector under transcriptional control of the 35S cauliflower mosaic virus promoter. In addition, the binary vector employed in these studies contained the npt-II gene, which is regulated by the nopaline synthase promoter, thus allowing the selection of plant transformants by kanamycin resistance. Kawchuk et al [34] inserted the CP gene into the cultivar Russet Burbank, reducing virus titer in transformed plants by 90% as compared with nontransformed control plants. In a second study [35], virus titer in transformed plants was reduced by 86 to 96% compared with control aphids (M. persicae) acquired virus from transgenic virus source plants 0 to 43%) less efficiently than from nontransformed virus source plants [35]. In this study, disease symptoms did not develop in transformed plants, and extensive field testing revealed that one transformed clone had virus titer reductions of 29 to 68%) compared with nontransformed control plants. However, 24 to 45%) of these transformants had higher virus titers than the nontransformed plants. Analysis of eight field evaluations revealed consistent reductions in the virus titers of transformants [35]. In multiple field experiments, primary and secondary symptoms reportedly developed in the foliage of transformed clones but were less severe than in control plants. Transformed clones showed reductions in the incidence and severity of tuber net necrosis of 88 and 92%), respectively [36]. A somaclonal variant derived from callus in tissue culture and intended to be a susceptible control also was resistant, showing reduced virus titer and lowered net necrosis incidence and severity. Brown et al. [37] measured the PLRV titer in primary and secondary infections of Russet Burbank and Ranger Russet cultivars transformed with the PLRV CP gene. For Russet Burbank clones in which virus buildup was most inhibited.
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reductions in primary and secondary infections were 77% and 85%, respectively, compared with nontransformed control plants. Similarly, virus titer reductions were 64 and 69%) for primary and secondary infections of Ranger Russet clones, respectively. However, some transformants of both cultivars had a virus titer equal to that of nontransformed control cultivars. Presting et al [38] examined two CP constructs with different lengths of the untranslated sequence upstream (5') of the coding region. Russet Burbank and Ranger Russet were transformed with these constructs, as well as a vector lacking the CP gene (empty-vector control). The goal of this work [38] was to produce a resistance phenotype based on increased translational initiation of the CP gene. However, differences relative to virus titer buildup and construct type were not observed [38]. The most resistant Russet Burbank and Ranger Russet transformants showed virus titer reductions of 92 and 96%), respectively, compared with nontransformed controls. Interestingly, three empty-vector transformants of Russet Burbank displayed 54%) virus titer reduction compared with nontransformed controls. In addition, one Ranger Russet empty-vector transformant showed a 75%) reduction in virus titer compared with the nontransformed plants. There was a strong correlation between virus titer in individual infected transformants and percentage incidence of infection for both cultivars (r = 0.94, P < .001). Van der Wilk et al. [39] examined the resistance of cultivar Desiree transformed with the PLRV CP gene. Assuming colinearity of enzyme-linked immunosorbent assay (ELISA) optical density values and virus in primaryinfected plants, virus titer reductions in resistant transformants were 74 to 64%o of titers in nontransformed, tissue-culture regenerated controls. Correspondingly, daughter tubers from these plants showed virus titer reductions of 33 to 65%) compared with nontransformed controls that were similarly regenerated. Thomas et al. [40] examined the incidence of infection, foliar symptom severity, and incidence and severity of net necrosis in a number of Russet Burbank PLRV CP transformants during two years of field trials. Viruliferous aphids were manually placed on each plant, providing a uniform inoculum that was earlier and more intense than normal inoculum pressure in commercial field production. The incidence of infection, delayed in certain transformants, ranged from 60 to 100%) in the first-year trial, but all clones were 100% infected in the second-year trial. Symptom severity was reduced in certain clones over both years. Virus titer ranged from 26 to 59%) that of nontransformed controls. In addition, the statistical reduction in the incidence of net necrosis to one-fourth that of the susceptible controls is economically significant. Net necrosis severity in symptomatic tubers of the most resistant transgenic clones was about half that for nontransformed controls. The level of resistance in the most resistant transgenic clones was clearly economically useful under commercial growing practices in the Columbia River basin of the Pacific Northwest of the United States. However, Monsanto Chemical Co., the intellectual property holder, did not pursue commercialization of these lines in the United States.
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Barker et al [41] transformed the cultivars Desiree and Pentland Squire and noted virus titer reductions ranging from 15 to 82% of the titer of nontransformed controls in graft-inoculated plants. Attempts to detect transgene-encoded CP in uninfected plants were successftil in some immunoblotting tests but not in others. Virus titers in resistant breeding clones from traditional breeding programs were comparable with those in transgenics. These researchers [41] found partial resistance to infection and resistance to virus accumulation. In other studies, overall resistance was enhanced by combining breeding clone, host, and transgenic clone resistance. The best transformants had virus titers that were 1% of the titer in susceptible controls [42, 43]. Thus, it is possible to approach extreme resistance by transforming traditionally bred resistant clones with PLRV CP genes. Nevertheless, given sufficient inoculum pressure, all of the resulting transformants could be infected. The most resistant of the infected transformants had lower virus titers than nontransformed, breeding clone controls. Derrick and Barker [44] studied PLRV movement through grafted stem sections of distinct potato genotypes having either transgenic or host resistance. Virus moved across graft unions in about 14 days in susceptible, transgenic-resistant and host-resistant genotypes. Both transgenic and host-resistant genotypes differed markedly from susceptible materials in that they lacked virus in the external phloem bundles and companion cells, whereas susceptible genotypes showed substantial presence of virus in these tissues [44]. Palucha et al [45] transformed the Polish cultivar Bzura, which has a moderate level of host resistance to PLRV, with the viral CP gene. This transformation produced transgenic plants, which when infected showed an 80% reduction in virus titer compared with PLRV-infected, nontransformed controls. In addition, secondarily infected plants from daughter tubers had virus titers 87% lower than controls. After graft inoculation, the virus titer reduction in this particular transformant was 41% as compared with infected, nontransformed control plants. Murray et al [46] transformed the cultivar Late Harvest with the viral CP gene and were unable to detect PLRV CP in the resulting transformants. B.
Coat Protein Antisense Strategy
A CP gene inserted in reverse, 5' - 3' orientation is described as antisense. In practical terms, an RNA product of an antisense, cDNA, nuclear transgene would be exactly complementary to a specific portion of the sequence of a positivestrand RNA virus. Theoretically, the RNA product of the CP antisense gene transcribed in the nucleus would be transported to the cytoplasm, where it would hybridize with the sense viral RNA and block translation of the CP encoded by the infecting virus. Kawchuk et al [35] transformed Russet Burbank with an antisense PLRV CP gene. Two transformants showed virus titer reductions of 93% and 95%, respectively, compared with nontransformed control plants. Aphid transmission efficiency from one of these clones was reduced 72%, compared with aphids acquiring virus from PLRV-infected, nontransformed controls.
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Palucha et al [45] transformed the cultivar Bzura with antisense PLRV CR One PLRV-infected transformant was characterized by a virus titer that was 21% of that observed in infected, nontransformed controls. Secondary infection in plants derived from daughter tubers resulted in titers only 13% of those in control plants. When challenged by graft inoculation, this particular clone showed a virus titer 10%) that of the control. C. Replicase Strategy In 1990, Golemboski et al [47] discovered that a portion of the replicase gene of tobacco mosaic virus (TMV) conferred extreme resistance to TMV when expressed in tobacco. In PLRV, ORFs 1 and 2 contribute to the viral replicase (Fig. 1). Open reading frame 2 is translated by means of a readthrough frameshift near the end of ORF 1, which terminates at the 3' end of ORF 2 to produce a 118kDa protein [26]. Kaniewski et al. [48] and Thomas et al [49] investigated resistance obtained by transforming Russet Burbank with PLRV replicase constructs that represent fulllength (ORF 1 and ORF 2) or truncated (ORF 2) versions of the gene. Populations of transformants harboring either full-length or truncated replicase gave rise to extremely resistant phenotypes. However, the full-length construct yielded transformants at a higher frequency. The inability to detect virus was accompanied by an absence of foliar symptoms and net necrosis. Moreover, virus also was undetectable in indexed plants regrown from tubers of inoculated plants [48]. Mitsky et al [50] reported (in a patent assigned to Monsanto) that both full-length PLRV replicase (ORF 1 and ORF 2) and a truncated replicase (ORF 2) produced phenotypes approaching extreme resistance to virus infection. The results of the replicase strategy were clearly superior to CP or antisense-CP in terms of the level of resistance. In 1998, Monsanto released this strategy to the marketplace as Newleaf Plus, a Russet Burbank transformant with replicase-mediated resistance to PLRV and resistance to Colorado potato beetle mediated by Bacillus thuringiensis toxin. D.
Putative Movement Protein Strategy
The ORF 4 is internal to ORF 3, the CP gene, and encodes a 17-kDa protein. This protein has nucleic acid binding properties suggestive of movement protein functioning [29]. Cultivar Desiree potato plants transformed with mutant forms of this gene were resistant to PLRV, whether the putative movement protein was expressed or not. Protein-producing plants also were resistant to PVX and PVY. All infected "resistant" plants had virus titers; hence, the definition applied, in this case, was that of moderate titer reduction (29 to 12% of titers in nontransformed controls) [51]. The role of the 17-kDa protein product of ORF 4 in virus movement was confirmed when immunolabeling revealed the pro-
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tein's preferential association with plasmodesmata interconnecting sieve elements and companion cells [30]. Constitutive expression of ORF 4 in tobacco, even in the absence of virus, resulted in limited resistance to PVY and PLRVlike disease symptoms. The latter suggests a viral movement function, which to some degree might affect unrelated viruses as well [52]. This further confirms the phloem as the primary site of action and indicates that the alteration in carbohydrate metabolism characteristic of PLRV (i.e., starch buildup in leaves, thickening of leaves and leaf rolling) is mainly due to the action of the 17-kDa gene product. E.
Ribozyme Strategy
Ribozymes are RNA molecules that cleave RNAs containing GUC, GUU, and GUA sequences. Lamb and Hay [53] showed that RNA molecules generated in vitro could cleave RNA derived from PLRV cDNA or particles. In vitro assays showed that cleavage occurred immediately downstream of the requisite GUC sequences in the replicase gene (ORF 2) and CP gene (ORF 4). This strategy has been discussed as a PLRV control methodology, but to date no studies have been published.
IV. Resistance Mechanisms After one and one half decades of transgenic research involving potatoes and PLRV, there is little information in the literature on the mechanisms of either transgenic or natural host resistance. At least one study suggests that patterns of PLRV accumulation in transgenic and naturally resistant clones are identical [44]. Hypotheses of the mechanism(s) of transgenic resistance, which are as unsupported now as they were at their conception, fall into three categories (Fig. 2). The first is that transgenic resistance resembles cross-protection. Hypothetically, cross-protection works because the presence of viral gene product results in feedback inhibition of viral replication. The presence of viral CP hypothetically impedes viral particle uncoating or interferes with viral particle assembly. As with cross-protection, only strains of the same virus are inhibited [54, 55]. Interest in replicase genes (a gene encoding a nonstructural component of virus particles) was intensified with the near immunity discovered with TMV replicase trangenes in tobacco [47]. A second hypothesis, mainly based on research from potyviruses, proposes that transgenic-produced viral RNA interacts with viral RNA to disrupt the life cycle of the virus. Transgenic RNA presumably interacts directly with invading RNA to short-circuit the infection sequence. The highest levels of resistance to PVY or tobacco etch virus (TEV) occur in transgenic plants carrying nontranslatable versions of the PVY or TEV CP gene [56-58]. Possibly positive-
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Fig. 2 Hypothetical mechanisms for the development of transgenic virus resistance. (A) Interference of product of a transgene with the gene product of an invading viral genome [54, 55]. (B) RNA of transgene interacts with RNA of invading virus and obstructs normal expression or replication [56]. (C) Transgenic RNA triggers a host mechanism, which identifies foreign RNA in a sequence-specific manner and degrades it [59].
Strand RNA of the transgene, unable to complex with ribosomes, forms a double-stranded complex with the negative-strand RNA of the invading virus. It is hypothesized that this "unnatural" RNA hybridization sequesters viral RNA template needed to produce positive-strand RNA for progeny virus particles [56]. Nontranslatable transgenes of PLRV have not yet been employed in transgenic PLRV resistance research. A third hypothesis suggests that viral RNA production by the host's transformed genome predisposes the host cell to activate a cellular process that degrades invading foreign RNA [59]. This predispostion by the host to recognize and destroy viral RNA inhibits virus replication and disease development. This host cell immune response may destroy the transient, double-stranded RNA formed during viral replication. Studies involving mutant, nontranslatable PLRV transgenes have not been reported. However, equivocal results involving detection of CP or replicase gene product(s) does not rule out such a resistance mechanism [59].
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Replicase transgenes apparently provide a viable solution to controlling PLRV, arguably the most damaging virus of potato. The virtual absence of public activity in this area of research is due perhaps to the perception that the approach is patented and unavailable [50]. However, the patent for use of replicase for viral resistance, wherein the readthrough of the replicase gene is deleted, has been issued to an independent party [60]. Therefore, replicase strategy research may be more of an option than previously considered.
V. Concluding Remarks PLRV titer reduction can be achieved with plants carrying the CP gene in sense or antisense orientation. It is unclear whether CP synthesis is an essential step in titer reduction. All published reports indicate that numerous transformants with demonstrable PLRV CP or antisense CP are not highly resistant to virus. Furthermore, in the case of CP sense and antisense transformants, complete or extreme resistance to infection has not been reported. It is conceivable, however, that a useful level of resistance, one worthy of commercial exploitation, could be obtained with either approach. Transformants carrying the 17-kDa movement protein gene also show reduced PLRV titers, although this gene seems less effective than the CP gene in this regard. Without a doubt, the replicase strategy produces the highest level of resistance; high to extreme resistance occurs frequently in transformed populations. No virus was detected in most resistant transformants of Russet Burbank, even under field conditions with intense inoculation pressure [48]. The mechanism of transgenic resistance to PLRV is not understood. It would be worthwhile to determine if the "nontranslatable" strategy that works so well with pot3rv^iruses also will produce resistance to PLRV Protocols that can localize PLRV particles and viral gene products in plant hosts will facilitate research on resistance arising from transgene expression. The discovery that symbiotic bacteria are involved in aphid acquisition and transmission of PLRV presents interesting possibilities for novel approaches to virus control [61] (see also chapter 11]. Studies have shown that interfering with symbiont production of symbionin affects virus transmission by the green peach aphid, M persicae. Control strategies based on affecting the biology of this symbiont or its production of symbionin or both could prove potent remedies to the PLRV problem in commercial potato production. Administration of environmentally friendly chemicals that would mainly act to disrupt interactions among virus, aphid, and plant may be possible [61]. The possibilities of controlling PLRV by application of aphid antifeedants such as the limonoid azadirachtin from the Neem plant, Azadirachta indica A. Juss., also are being studied [62, 63]. Because PLRV requires potato culture to be a sustained presence in agriculture, the virus would essentially disappear if potato were disguised to prevent aphids from recognizing it as a suitable host.
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Finally, making replicase strategy generally available would facilitate potato production worldwide and be particularly helpful in capital-limited countries.
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20. Valkonen, J.P.T. (1994). Natural genes and mechanisms for resistance to viruses in cultivated and wild potato species (Solanum spp.). Plant Breed. Ill, 1-16. 21. Davidson, T.M.V (1980). Breeding for resistance to virus disease of the potato (Solanum tuberosum) at the Scottish Plant Breeding Station. In "59th Annual Report of the Scottish Plant Breeding Station 1979-1980," pp. 100-108. 22. Ross, H. (1986). Potato breeding—^problems and perspectives. Adv. Plant Breed., Supplement 13. Paul Parey, Berlin and Hamburg. 23. Brown, C.R., Corsini, D., Pavek, J., and Thomas, R (1997). Heritability of field resistance to potato leafroll virus in cultivated tetraploid potato. Plant Breed. 116, 585-588. 24. Swiezynski, K.M., Dziewondska, M.A., and Ostrowska, K. (1989). Resistance to potato leafroll virus (PLRV) in diploid potatoes. Plant Breed. 103, 221-227. 25. Brown, C.R., and Thomas, RE. (1994). Resistance to potato leafroll virus derived from Solanum chacoense: Characterization and inheritance. Euphytica 74, 51-57. 26. Mayo, M.A., and Ziegler-Graft, V (1996). Molecular biology of luteoviruses. Adv. Virus Res. 46, 413-^60. 27. Smith, O.P., and Harris, K.R (1990). Potato leafroll virus V genome organization: Sequence of the coat protein gene and identification of a viral subgenomic RNA. Phytopathology 80, 609-614. 28. Bahner, I., Lamb, J., Mayo, M.A., and Hay, R.T. (1990). Expression of the genome potato leafroll virus: Readthrough of the coat protein termination codon in vivo. J. Gen. Virol. 71, 2251-2256. 29. Tacke, E., Prufer, D., Schmitz, J., and Rohde, W. (1991). The potato leafroll luteovirus 17k protein is a single stranded nucleic-acid binding protein. J. Gen. Virol. 72, 2035-2038. 30. Schmitz, J., Stussi-Garaud, C , Tacke, E., Prufer, D., Rohde, W, and Rohfrtisch, O. (1997). In situ localization of the putative movement protein (prl7) of potato leafroll virus (PLRV) in infected and transgenic potato plants. Virology 235, 311-322. 31. Van der Wilk, K, Hourterman, P., Molthofif, X, Hans, R, Dekker, B., Van den Heuvel, J., Huttinga, H., and Goldbach, R. (1997). Expression of the potato leafroll virus ORFO induces viral-diseaselike symptoms in transgenic potato plants. Mol. Plant Microbe Interact. 10, 153-159. 32. Ashoub, A., Rohde, W., and Prufer, D. (1998). Inplanta description of a second subgenomic RNA increases the complexity of the subgroup 2 luteovirus genome. Nucleic Acids Res. 26, 420^26. 33. Sanford, J.C, and Johnston, S.A. (1985). The concept of parasite-derived resistance: Deriving resistance genes from the parasite's own genome. J. Theor Biol. 113, 395^05. 34. Kawchuk, L.M., Martin, R.R, and McPherson, J. (1990). Resistance in transgenic potato expressing the potato leafroll virus coat protein gene. Mol. Plant Microbe Interact. 3, 301-307. 35. Kawchuk, L.M., Martin, R.R., and McPherson, J. (1991). Sense and antisense RNA-mediated resistance to potato leafroll virus in Russet Burbank plants. Mol. Plant Microbe Interact. 4, 247-253. 36. Kawchuk, L.M., Lynch, D.R., Martin, R.R., Kozub, G.C., and Parries, B. (1997). Field resistance to potato leafroll luteovirus in transgenic and somaclone potato plants reduces tuber disease symptoms. Can. J. Plant Pathol. 19, 260-266. 37. Brown, C.R., Smith, O.P, Damsteegt, VD, Yang, C.-P, Fox, L., and Thomas, RE. (1995). Suppression of PLRV titer in transgenic Russet Burbank and Ranger Russet. Am. Pot. J. 11, 589-598. 38. Presting, G.G., Smith, O.P, and Brown, C.R. (1995). Resistance to potato leafroll virus in potato plants transformed with the coat protein gene or with vector control constructs. Phytopathology 85,436-442. 39. Van der Wilk, R, Willink, D.R-L, Huisman, M.J., Huttinga, H., and Goldbach, R. (1991). Expression of the potato leafroll luteovirus coat protein gene in transgenic potato plants inhibits viral infection. Plant Mol. Biol. 17, 431^39.
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40. Thomas, P.E., Kaniewski, W.K., and Lawson, E.G. (1997). Reduced field spread of potato leafroll vims in potatoes transformed with the potato leafroll virus coat protein gene. Plant Dis. 81, 1447-1453. 41. Barker, H., Reavy, B., Kumar, A., Webster, K.D., and Mayo, M.A. (1992). Restricted virus multiplication in potatoes transformed with the coat protein gene of potato leafroll luteovirus: Similarities with a type of host gene-mediated resistance. Ann. Appl. Biol. 120, 55-64. 42. Barker, H., Reavy, B., Arif, M., Webster, K.D., and Mayo, M.A. (1994). Towards immunity to potato leafroll virus and potato mop-top virus by using transgenic and host gene-mediated forms of XQsisXdincQ. Aspects Appl. Biol. 39, 189-194. 43. Barker, H., Webster, K.D., Jolly, G.A., Reavy, B., Kumar, A., and Mayo, M.A. (1994). Enhancement of resistance to potato leafroll virus multiplication in potato by combining the effects of host genes and transgenes. Mol. Plant Microbe Interact. 7, 528-530. 44. Derrick, P.M., and Barker, H. (1997). Short and long distance spread of potato leafroll virus luteovirus: Effects of host genes and transgenes conferring resistance to virus accumulation in potato. J. Gen. Virol. 78, 243-251. 45. Palucha, A., Zagorski, W, Ghrzanowska, M., and Hulanicka, D. (1998). An antisense coat protein gene confers immunity to potato leafroll virus in a genetically engineered potato. Eur J. Plant Pathol. 104,287-293. 46. Murray, S.L., Burger, XT, Oelfse, D., Gress, W.A., Staden, J.V, and Berger, D.K. (1998). Transformation of potato (cv. Late Harvest) with the potato leafroll virus coat protein gene and molecular analysis of transgenic lines. S. Afr J. Sci. 94,263-268. 47. Golemboski, D.B., Lomonossoff, G.P., and Zaitlin, M. (1990). Plants transformed with a tobacco virus nonstructural gene sequence are resistant to the virus. Proc. Natl. Acad. Sci. U.S.A. 87, 6311-6315. 48. Kaniewski, W, Lawson, G., Loveless, J., Thomas, P., Mowry, T, Reed, G., Mitsky, T, Zalewski, J., and Muskopf, Y. (1995). Expression of potato leafroll virus (PLRV) replicase genes in Russet Burbank potatoes provide field immunity to PLRV In "Environmental Biotic Factors in Integrated Plant Disease Gontrol" (M. Manka, ed.), pp. 289-292. Polish Phytopathological Society, Poznan, Poland. 49. Thomas, P.E., Kaniewski, W.K., Reed, G.L., and Lawson, E.G. (1995). Transgenic resistance to potato leafroll virus in Russet Burbank potatoes. In "Environmental Biotic Factors in Integrated Plant Disease Gontrol" (M. Manka, ed.), pp. 551-554. Polish Phytopathological Society, Poznan, Poland. 50. Mitsky, T.A., Hemenway, G.L., and Tumer, N.E. (1996). Plants resistant to infection by PLRV U.S. Patent No. 5,510,253. To Monsanto Go. http://www.uspto.gov/patft/ 51. Tacke, E., Salamini, F, and Rohde, W (1996). Genetic engineering of potato for broad-spectrum protection against virus infection. Nature Biotechnol. 14, 1597-1601. 52. Herbers, K., Tacke, E., Hazirezaei, Krause, L.-P, Melzer, M., Rohde, W, and Sonnewald, U. (1997). Expression of a luteoviral movement protein in transgenic plants leads to carbohydrate accumulation and reduced photosynthetic capacity in source leaves. Plant J. 12, 1045-1056. 53. Lamb, J.W, and Hay, R.T (1990). Ribozymes that cleave potato leafroll virus RNA within the coat protein and polymerase genes. J. Gen. Virol. 71, 2257-2264. 54. Powell-Abel, P, Nelson, R.S., De, B., Hoffman, N., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1986). Delay of disease development in transgenic plants that express the tobacco virus coat protein gene. Science 232, 738-743. 55. Reimann-Phillip, U. (1998). Mechanisms of resistance: Expression of coat protein. In "Plant Virology Protocols" (G.D. Foster and S.G. Taylor, eds.), pp. 521-532. Humana Press, Totowa, NX 56. Lindbo, XA., Silva-Rosales, L., and Dougherty, W.G. (1993). Pathogen-derived resistance to potyviruses: Working, but why? Semin. Virol. 4, 357-361.
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57. Smith, H.A., Powers, H., Swaney, S., Brown, C , and Dougherty, W.G. (1995). Transgenic potato virus Y resistance in potato: Evidence for an RNA-mediated cellular response. Phytopathology 85,864-870. 58. Dougherty, W.G., and Lindbo, XA. (1996). Production of virus resistant plants. U.S. Patent No. 5,583,021. To the State of Oregon, Acting by and through the State Board of Higher Education on Behalf of Oregon State University, http://www.uspto.gov/patft/ 59. Haan, PD. (1998). Mechanisms of RNA-mediated resistance to plant viruses. In "Plant Virology Protocols" (G.D. Foster and S.C. Taylor, eds.), pp. 533-556. Humana Press, Totowa, NJ. 60. Zaitlin, M., and Palukaitis, P. (1999). Implanting disease resistance to plants with viral replicase DNA molecules which do not have a read-through portion. U.S. Patent No. 5,945,581. To Cornell Research Foundation, http://www.uspto.gov/patft/ 61. Van den Heuvel, J.F.J.M., Verbeek, M., and Van der Wilk, F. (1994). Endosymbiotic bacteria associated with circulative transmission of potato leafi*oll virus by Myzus persicae. J. Gen. Virol 75, 2559-2565. 62. Nesbit, A.J., Woodford, J.A.T., and Strang, R.H.C. (1996). The effects of azadirachitin on the acquisition and inoculation of potato leafi-oU virus by Myzus persicae. Crop Protect. 15, 9-14. 63. Van den Heuvel, J.EJ.M., Hogenhout, S.A., Verbeek, M., and Van der Wilk, E (1998). Azaduracta indica metabolites interfere with the host-endosymbiont relationship and inhibit the transmission of potato leafi-oll virus by Myzus persicae. Entomol. Exp. Appl. 86, 253-260.
CHAPTER 13
Bemisia: Pest Status, Economics, Biology, and Population Dynamics TJ. HENNEBERRY S J. CASTLE
/. Introduction In recent years, Bemisia tabaci (Gennadius) (Strain B = 5. argentifolii Bellows and Perring) has risen in status as a major pest of world agriculture as destructive crop infestations and viral diseases have proliferated. Increased globalization of agriculture and floriculture and international transport of plant material have contributed to the introduction of variant forms of both B. tabaci and viruses to regions where they were previously unknown. Economic losses have been estimated in the hundreds of millions of dollars in some affected areas [1, 2]. Increased whitefly management efforts have been generally successful, but at the cost of greater inputs and increased vigilance to avoid outbreaks of the magnitude observed in the early 1990s in certain regions of the southwestern United States [2-A]. Crop losses due to B. ^(2Z?ac/-transmitted viruses recently introduced into new areas [5] or characterized and described for the first time [6] have also contributed to the problem. Establishment of virulent whitefly biotypes and viruses continues in new geographical areas and emphasizes the need for greater focus on B. tabaci as a serious crop pest and virus vector. Various biological traits expressed by B. tabaci, including a broad host range, contribute to rapid population expansion during favorable seasons. In addition to climatic factors such as temperature, humidity, and rainfall, the quality and quantity of food resources also influence population growth. The types of crops grown and their relative acreages and phenologies during the annual crop cycle, together with wild and ornamental hosts, form the resource base available to B. tabaci. Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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T.J. HENNEBERRY AND S.J. CASTLE
Dispersal between fields or relocation to preferred hosts works in concert with its polyphagous nature to increase population densities in a multiple crop environment. High fecundity levels, coupled with fast developmental rates, help define an organism with a high intrinsic rate of increase and the capability to rapidly colonize a potential food source. Bemisia tabaci is broadly tolerant of harsh environmental conditions, it routinely survives and sometimes flourishes in desert climates that may range between -2° and 50°C. It is also highly adaptable to extreme stress factors such as insecticides, as evidenced by the multiple mechanisms of resistance that have evolved in populations around the world [7, 8]. How these traits are expressed in various environments strongly influences population growth and infestation levels in crops. Expansion within a particular environment also depends on natural and applied population-regulating factors. Bemisia tabaci has often been considered as an "upset" pest, induced to damage crop levels through misuse of insecticides [9, 10]. There are indeed many well-documented cases of insecticide resistance that argue for integrated management and nonchemical measures to suppress populations. However, some crop environments may represent such an optimal balance of conditions that natural control factors may be overwhelmed, thus leaving little choice but to counter with chemical measures. The key to successfixl management of any agricultural pest is understanding its intrinsic potential for increase in various environments while also knowing the capacity of countermeasures to suppress population growth. More information on the biology and population dynamics of B. tabaci is becoming available as it gains importance in additional regions. In this review, we examine basic life history traits oiB. tabaci and how they have contributed to an organism now widely recognized for its explosive populations and elevated pest status.
//-
Economic Impact and Pest Status
Crop damage results from direct feeding and associated yield reductions, contamination of produce and cotton (Gossypium spp.) lint with honeydew, and transmission of plant-pathogenic viruses. The 1986 outbreaks in Florida resulted in estimated losses of $140 million in the tomato {Lycopersicon esculentum Mill.) industry [ 11 ]. In 1991, losses for cotton and vegetables in Texas were estimated to be $24 and $29 million, respectively [12]. In southern California, crop losses of over $100 million a year and a reduction in 3,000 agricultural jobs annually have been reported since 1992 [13]. Bemisia tabaci has also been a serious problem in greenhouse culture throughout the world, with 1991 losses for ornamentals in the United States alone reported to have exceeded $23 million [14]. Yield reductions associated with B. tabaci-mducQA physiological plant disorders have occurred on squash {Cucurbita pepo L.), lettuce (Lactuca sativa L.), broccoli
13.
BEMISIA: PEST STATUS, ECONOMICS, BIOLOGY, AND POPULATION DYNAMICS
249
(Brassica oleracea van botrytis L.), and tomato [11]. Various other symptoms of leaf and plant disorders have been reported for numerous crops [15,16]. Cotton lint stickiness from honeydew contamination is a major issue in the textile industry [17]. Price reduction differentials greater than 10% as compared with nonsticky cottons may be imposed and gin turnout may be reduced up to 25% [18]. Cotton leaf crumple virus (CLCV) in California resulted in cotton yield reductions of 41 to 81%) [19] in the mid-1960s, and major epidemics of CLCV occurred in Arizona during 1981 to 1984 [20, 21]. Crop losses in Arizona and California in 1981 exceeded $100 million from the B.toZ?(2c/-transmittedsquash leaf curl virus (SLCV) and lettuce infectious yellows virus (LIYV) [22]. Further losses in melons (Cucumis melo) and lettuce due to viral diseases were reported in Arizona in 1982 [21]. For the Near Eastern countries of Bahrain, Cyprus, Egypt, Iran, Iraq, Jordan, Kuwait, Lebanon, Libya, Malta, Saudi Arabia, and Turkey, viral diseases of vegetables have resulted in losses ranging from 5 to 80% [23], in most cases from tomato leaf curl virus, cucumber vein yellowing virus, or cucumber yellows virus. Cotton leaf curl virus is estimated to result in 20 to 25% crop value losses annually in Pakistan [23]. Bemisia tabaci-bomQ geminiviruses are also prevalent in Central and South America, Africa, Japan, and Australia [24]. Geminiviruses in the Americas and the Caribbean basin cause as many as 31 plant viral diseases [25]. A wide range of legume crops, cotton, cucurbits, tomatoes, peppers, potatoes, and numerous weeds are involved in the ecology of both vector and virus worldwide. With the exception of specific viral diseases spread by B. tabaci and the particular crops that are infested in a given area, the general characteristics of recent B. tabaci infestations are similar to those reported as early as the 1920s in India [26-28] and the 1930s in Israel [29] and the Sudan [30, 31]. Severe problems with viral diseases were also encountered during the 1930s [32-34]. Development of large infestations on multiple crops and wild hosts resulting in direct and indirect damage has marked B. tabaci as a serious pest for many decades. The perception ofB. tabaci as a more serious pest during the 1990s began in the 1980s with the onset of increasingly virulent infestations in North America. The 1981 outbreak in the Imperial Valley of California and in Yuma, Arizona were preceded by outbreaks in Turkey in 1974 [35], the Sudan during the 1970s [9, 36], and Israel between 1976 and 1988 [37]. Cotton was the principal crop affected by severe infestations in each of those outbreaks. At the end of the season, mass dispersal out of cotton and into surrounding fields sometimes severely impacted other crops. For example, large numbers of dispersing whiteflies out of cotton led to the epidemic of LIYV in the Imperial Valley in 1981 [22]. Most of the direct damage was concentrated in cotton, with the indirect damage due to LIYV occurring in lettuce and melons. By the late 1980s in North America, the types of crops infested by B. tabaci became less predictable, and it was no longer only an agricultural pest. Heavy infestations in some of the floricultural crops grown in greenhouses were occurring in Florida and soon followed in other states. The
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T.J. HENNEBERRY AND S.J. CASTLE
focus of attention then shifted back to agriculture, as more dynamic infestations occurred in CaHfornia, Arizona, Texas, and Florida on other crops beside cotton.
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Taxonomy Flux
Bemisia tabaci was first described in 1889 from tobacco as Aleurodes tabaci (Gennadius) and has since been variously referred to as the tobacco, cassava, sweet potato, or cotton whitefly Its center of origin was suggested as the Indian subcontinent [38], but tropical Africa has also been considered to be a plausible center of origin based on combined information from the fossil record, current geographic distributions, and molecular phylogenetic analysis of nucleotide sequences of 18S rDNAs [39]. Numerous other descriptions of this species were madefi-omdiverse locations in the world, including the United States [40], Brazil [41], India [26], and Nigeria [42]. Several of these species were synonymized as B. tabaci [43, 44]. By consolidating earlier descriptions ofB. tabaci, it was recognized that differences in the appearance of the fourth nymphal instar, the key stage used to separate genera and species in the Aleyrodidae, were strongly correlated to morphological characteristics of the rearing host [45] and not representative of species-specific differences. Bemisia infestations during the 1950s and 1960s were widely accepted as B. tabaci even though several common names were retained. At that time, B. tabaci was known to occur in Europe, Asia, Russia, Africa, United States, Central America, South America, Australia and the Caroline and Mariana Islands, Fuji, Papua, and New Guinea [43, 46]. The taxonomic picture became more complex beginning in 1986 with the proliferation of B. tabaci on poinsettia {Euphorbia pulcherrima Willdenow) and other floricultural crops in Florida, which previously had not commonly been infested by B. tabaci. At about the same time, a disorder of squash that produced silvering of foliage also appeared in Florida. Similar phenomena began to occur in other states (Arizona, California, Florida, Hawaii, New York, Ohio, Tennessee, and Texas), the U.S. Territory of Puerto Rico, and Canada [47]. The strain found on poinsettia was suspected to be a genetically different whitefly, but no clear supporting evidence occurred until different esterase banding patterns were identified [48]. The esterase electromorphs were designated as A and B types from cotton and poinsettia, respectively. Only the B-type whiteflies could induce squash silverleaf (SSL) symptoms [48]. Further examination of differences between A and B types showed biochemical [49, 50] and molecular [51] variation, reproductive isolation [51], and morphological differences in the fourth instar nymphal stage [52]. It was proposed that type B was sufficiently different from type A to warrant species separation and, subsequently, it was described as Bemisia argentifolii Bellows & Perring [51, 52]. Further studies have revealed some ambiguity concerning this species separation.
13.
BEMISIA: PEST STATUS, ECONOMICS, BIOLOGY, AND POPULATION DYNAMICS
251
At the molecular level, only two base-pair substitutions out of an average length of 1,048 base pairs in 185 rDNAs were observed between B. tabaci and B. argentifolii. In contrast, as is similar to interspecific distances between congeners of other whitefly genera, 16 base-pair substitutions were observed in the comparison of B. berbericola (Cockerell) with either B. tabaci or B. argentifolii [39]. However, molecular data obtained via application of rapidly amplified polymorphic DNA (RAPD) techniques showed genetic difference between B. tabaci types A and B to be as large as those for two other whitefly species, Parabemisia myricae Kuwana and Trialeurodes abutilonea Haldeman [53]. Limited matings between A and non-B laboratory strains [10] and wild-type crosses in the field between type B and a non-B type indigenous to Australia [54] have resulted in female progeny production, thus indicating some degree of reproductive compatibility between B and non-B types. Insofar as differences in morphological characters between B. tabaci and B. argentifolii are concerned, the issue of host plant influences on variability in thoracic tracheal folds was addressed by the observation that they are "usually absent in the specimens from cassava, long and narrow in those from tobacco, and most clearly developed in form from Dolichos, in which they are short and broad" [45]. This trait and another that involved whether the anterior submarginal seta 4 (ASMS 4) was present or absent on fourth instar nymphs were noted as two morphological differences between B. argentifolii and B. tabaci in the argument for separating these two putative species [52]. The ASMS 4 character appears less dependable than the thoracic tracheal fold. Two specimens of B. argentifolii were observed with the ASMS 4 present, whereas it was absent in the holotype specimen [52]. The absence of ASMS 4 also was reported for specimens from non-B populations from the Old World, just as it was for B-type populations [55]. The debate over the proper taxonomic designation has opened a much wider investigation into the relationships ofB. tabaci populations worldwide. Thus far, variability among many geographic populations has been reported at the same level as that revealed in the comparison between B. tabaci types A and B. Assessment of variability in esterases among populations of B. tabaci has extended the list of esterase electromorphs beyond the initial A and B designees [56]. Although RAPD analyses showed continuous variation among the majority of B. tabaci populations tested from around the world [57], it is unclear at this time how variability at the molecular level should be treated taxonomically [58]. In spite of the ambiguity among particular characters used to separate B. tabaci and B. argentifolii, it appears valid to acknowledge differences among B, tabaci populations where they exist as clearly as between B. tabaci and B. argentifolii, which are also known as B. tabaci type A and type B, respectively. No other biotype has demonstrated as great a capability to induce SSL symptoms or as large a number of other plant disorders as does the B biotype. It also appears to be more capable of colonizing a wider number of plants and crops as compared with other B. tabaci biotypes; hence, its greater pest potential.
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T.J. HENNEBERRY AND S.J. CASTLE
The B biotype, based on esterase banding and the silverleaf symptom, has been identified in areas of the Caribbean region, United States, Mexico, Central and South America, Canada, Africa, the Arabian Peninsula, Japan, the Mediterranean basin, Britain [47, 50, 59-68], and more recently Australia [54] and Pakistan [69]. The origin of B. tabaci type B is uncertain, but the chronology associated with reports of type B suggested [70] either an origin in the Dominican Republic or Puerto Rico or a Central or South America origin based on greater esterase variation among B, tabaci populations in these as compared to other regions [68]. However, these suggestions are questionable in view of observations from Israel of a B. tabaci colony, collected and maintained since 1960, being able to induce SSL and also conforming to the B-type esterase pattern [71]. Further studies of squash silverleaf symptoms in Israel [72], which originally presumed a physiological disorder of squash, in retrospect suggest the undetected presence of Btype B. tabaci as the probable inducer of "leafsilvering" of squash. Our policy in this review will be to retain B. tabaci as the specific designation. Biotypes A and B will be designated as described by the original authors in papers published since 1986. For publications prior to 1986, biotype designations based on esterase typing will not be considered, but host plant associations will be given when they suggest possible biotype differences.
IV. Population Dynamics A.
Biological Traits 1.
REPRODUCTION
Adult emergence generally occurs between 0800 and 1200 hours [28, 73, 74] (Table I). Both sexes are sexually immature at emergence; little activity occurs for up to 10 hours, while wax is transferred from an abdominal gland to the wings [75]. Females accept courting males 10 to 24 hours after emergence when males are at least 10 hours old. The preoviposition period varies with temperature and may range from 1 to 22 days under field and insectary conditions [29] and 2 to 5 days under laboratory conditions of constant temperatures ranging from 16° to 28°C [73, 76]. Following the preoviposition period, maximum egg production is reached quickly in about 5 days at 25° and 28°C and about 10 days at 19° and 22°C for females ovipositing on tobacco for 25 to 27 days and 30 to 40 days, respectively [76]. Thereafter, numbers of eggs per day decline. Parthenogenetic reproduction occurs with unfertilized eggs producing males [28]. Sex ratios are variable and may range from 0.5 to 0.8 female [73], and in most cases females have been reported to predominate in insectary and field populations [34, 77-81]. In a laboratory study, the proportion of female progeny increased with temperature, from 0.60 to 0.63, 0.69, and 0.76 females at temperatures of 19°, 22°, 25°,
13.
BEMISIA: PEST STATUS, ECONOMICS, BIOLOGY, AND POPULATION DYNAMICS
Table I
253
Some Biological Characteristics of Bemisia tabaci
Activity
Strain
Emergence
A, B
Parthenogenetic
A, B
Mating
A, B
Preoviposition
A B
Longevity
A
B
Description Dorsal through a T-shaped integument fissure, 0800 to 1200 h, little movement up to 10 h while wax is transferred from an abdominal gland to wings. Unfertilized eggs produce males, sex ratios range 1:1 to 1:6.
Source [28, 73-75]
[28, 73, 76,79, 80,81] [28, 72]
On emergence, both sexes immature; males seldom accepted before 10-24 h of age. 1 to 22 days at 26.7°C. Field and insectary 2-3 days. [29, 73] Range 3.7 to 2.1 days at 16° to 28°C on tobacco, [76] range 4.3 to 2.2 days at 16° to 28°C on poinsettia. Females live longer than males, 13.2 to 62.0 days in [29, 73, the field, males 4.0 to 11.7 days at 14° to 32°C; 74, 92, females 10.0 to 35.0 days at 14° to 32°C in the 85, 86, laboratory. Variability apparent between cotton, 88, 157] poinsettia, eggplant, sweet potato, tomato, and lubia. Females live 9.9 to 50.8 days at 16.0° to 32.2°C. [76, 83, Variability occurs between cotton, poinsettia, 84, 86] tobacco, eggplant, tomato, sweet potato, cucumber, and garden bean.
and 28°C, respectively [76]. Age-specific mortalities for the A strain at temperatures between 20° and 32°C in the laboratory were, respectively, 17-28%, 2-16%, 3-14%, and 2-24% for first, second, third, and fourth instar nymphs on cucumber and cotton [82]. Results were similar for the B strain on eggplant {Solarium melongena L.) [83], tomato, sweet potato, cucumber, and green bean [84]. Under laboratory conditions males were observed to live 8-10 days at 16° to 32°C and females 10-35 days at 14° to 32°C [74, 76, 85, 86,]. Under field and insectary conditions, the range of longevity is considerably wider, depending on the time of year, with male longevity ranging from 6 to 34 days and female longevities from 15 to 55 days with temperatures ranging from 12.7° to 26.5°C [29]. 2.
LIFE HISTORY
a. Stages. Eggs are elongate with an attached pedicle, which is imbedded in plant tissue during oviposition. Certain leaf surface characteristics such as lamina trichomes may, at least for cotton, provide orientation cues for oviposition site selection [87], which may facilitate access of nymphs to minor vascular bundles for feeding. Under field conditions of varying temperature, oviposition by strain A has ranged from 28 eggs [28] to 160 eggs [88] per female on cotton, 37 to 44 on tobacco [34], 50 on eggplant [29], 108 on lubia [89], and 161 on sweet potato [73]
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TJ. HENNEBERRY AND S J . CASTLE
(Table II). In the laboratory, with temperatures ranging from 25.0° to 27.0°C, eggs per female on cotton ranged from 32 [86], 81 [74], and 93 [90] to 28 [85, 90] and 344 [9]. Fewer eggs per female of 72 [74] and 95 [91] were laid at temperatures of 30.0° to 32.6°C, whereas 56 and 76 eggs per female were laid on tomato at 14° and 25°C, respectively [92]. For strain B, females laid 51 eggs per female on cotton and 85 on poinsettia [86] at 16 to 28°C. Eggs per female were reported to range from 48 to 96 on poinsettia [76], and on eggplant, tomato, sweet potato, cucumber, and garden bean the values ranged from 21 to 324 [83, 84]. Egg hatch on cotton has been less variable than oviposition, ranging from 81 to 98% at 20° to 35°C, 59% at 40°C, and 62% at 15°C [74, 82, 88]. On cucumber, 73.7 to 96.5% egg hatch was observed [82] (Table II) and on cotton 81 to 99% at 16° to 28°C [76]. On eggplant, 90 to 96%o egg hatch was observed between 15° and 35°C [83], with a similarly high egg hatch of 95 to 96% occurring on eggplant, tomato, sweet potato, cucumber, and garden bean at 25°C [84]. b. Development Immature stages consist of the egg and four nymphal instars. Total developmental times in the field (egg to adult) are highly variable ranging, from 14 to 107 days (Table III). In the laboratory, egg development occurs in 5.2 to 10.7 days at 22° to 31°C [82, 93, 85]. Faster developmental rates occur at higher temperatures [6, 83, 84]. Development of nymphal instars of strains A and B on various hosts and at different temperatures is shown in Table 3. The results of several studies cited in this paper [94, 95] show the effect of different host plants on developmental rates. Differences in developmental rates of as much as 10 days have been observed on different hosts. The effects of the host plant on variable developmental rates must also be considered an important factor in population dynamics. c. Longevity. Age-specific mortalities for the A strain at temperatures between 20° and 32°C ranged from 17 to 28%, 2 to 16%, 3 to 14%, and 2 to 24% for first, second, third, and fourth instar nymphs on cucumber and cotton [82]. Results were similar for the B strain on eggplant [83] and tomato, sweet potato, cucumber, and green bean [84]. Under laboratory conditions males were observed to live 8 to 10 days at 16° to 32°C and females 10 to 35 days at 14° to 32°C [74,76, 85, 86] (Table I). Under field and insectary conditions the range of longevity is considerably wider, depending on the time of year, with male longevity ranging from 6 to 34 days and female longevities from 15 to 55 days with temperatures ranging from 12.7 to 26.5°C [29]. B.
Ecological Traits 1.
INTRAPLANT DISTRIBUTION
Bemisia tabaci is most frequently found on the undersurfaces of leaves, often frustrating conventional chemical control approaches. The higher rate of
Table II
Strain A
B
Effects of Temperature and Host on Bemisia tabaci Fecundity and Egg Hatch Temperature (°C) 9.4-42 9.4-34.4 12.7-26.5 29.0-33.0 31.9-38.0 15-40 25-26 26.7 32.6 30.0 22-30 27.0 30.0 25-26 25-26.5 14.0 25.0 26.7 26.7 20.0 25.5 29.0 32.0 20.0 25.5 29.0 32.0 26.7 26.7 16.0 19.0 22.0 25.0 28.0 16.0 22.0 28.0 20.0 25.0 27.0 30.0 35.0 25.0 25.0 25.0 25.0 25.0
Host«
Eggs per female (number)
Hatch
(%)
Source
28-43 44-77 50 108 48-394
_ -
[28] [34] [29] [89] [73] [88] [158] [74] [74] [91] [91] [85] [159] [9] [9] [92] [92] [86] [86] [82] [82] [82] [82] [82] [82] [82] [82] [86] [86] [76] [76] [76] [76] [76] [76] [76] [76] [83] [83] [83] [83] [83] [84] [84] [84] [84] [84]
Cotton* Tobacco* Eggplant* Lubia* Sweet potato* Cotton Cotton Cotton Cotton Cotton Cotton Cotton Tomato Cotton Cotton Tomato Tomato Cotton Poinsettia Cotton Cotton Cotton Cotton Cucumber Cucumber Cucumber Cucumber Cotton Poinsettia Tobacco Tobacco Tobacco Tobacco Tobacco Poinsettia Poinsettia Poinsettia Eggplant Eggplant Eggplant Eggplant Eggplant Eggplant Tomato Sweet potato Cucumber Garden bean
-
59-98
-
257 81 72 95 93 128 203 309-344 257-286 56 76 31.8 22.3
91.7
-
84.5 90.5 98.0 94.9 73.3 96.5 86.8 92.8
51.2 85.0 47.8 81.5 75.7 72.3 61.6 60.2 90.9 96.3 324.4 223.7 186.4 57.8 21.7 223.7 167.6 77.5 66.0 83.5
68 75
-
-
80.7 90.8 97.9 99.0 97.2
-
90.3 96.3 94.5 90.0 97.2 94.5 96.0 94.7 95.0 95.3
^ Experimental environment: asterisk = field or greenhouse conditions; no asterisk = controlled temperature and lighting conditions.
255
Table IZI Effects of Temperature and Host on Bemisia iabaci Egg and Nymph Development Nymph development (days) Temperature" ("C)
N vI
cn
9.4 to 42.0* 12.7 to 26.5* Summer* Dec* 29-33 25.4 31.0 25.4 28.0 14.9 16.7 20.0 22.5 25.0 27.5 30.0 32.5 36.0 30.0 30.0 29-33 15.0 20.0 25.0 30.0 35.0
Host Cotton Eggplant Tobacco Tobacco Lubia Sweet potato Sweet potato Potato Cauliflower Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Tomato Lubia Cotton Cotton Cotton Cotton Cotton
Egg development (days) 3-33 4-22 3 4 7-10 5.0 5.9 -
3.7
Instar I
Instar 2
-
-
-
-
3-5 8-10 2.4 4.5 3.0 4.5 3.5
2-6 6-9 1.8 2.7 2.0 3.0 1.9
2 4 &9 2.5 2.6 I .9 2.8 2.0
2-5 4-6 4.0 6.2 4.7 5.8 7.1
Instar 3
Instar 4 2-8
14107 11-75 -
17.5
65.1 48.7 34.7 27.8 23.6 17.8 16.6
22.5 11.5 9.9 7.6 6.1 5.4 5.0
**
*
-
5.3 6.2 5.0 20.5 11.8 8.3 7.5 5.8
Egg to adult (total days)
2.5 2.2 2.4
3.0 2.3 1.8
-
-
2.6 3.1 2.4
4.3 6.0 4.0
-
17.7 19.6 15.6
Source
h)
3
40.0 14.0 25.0 26.7 26.7 20.0 25.5 29.0 32.0 20.0 25.5 29.0 32.0 26.7 26.7 16.0 19.0 22.0 25.0 28.0 15.0 20.0 25.0 26.7 30.0 35.0 25.5 25.5 25.5 25.5 25.5
Cotton Tomato Tomato Cotton Poinsettia Cotton Cotton Cotton Cotton Cucumber Cucumber Cucumber Cucumber Cotton Poinsettia Poinsettia Poinsettia Poinsettia Poinsettia Poinsettia Eggplant Eggplant Eggplant Eggplant Eggplant Eggplant Eggplant Tomato Sweetpotato Cucumber Garden beans
5.2 16.5 6.4 -
8.8 4.8 4.2 3.9 12.4 5.8 4.1 4.1 -
45.5 23.3 16.9 14.0 10.4 25.8 10.3 5.9 4.9 4.2 4.9 6.0 6.2 6.1 6.4 6.0
-
-
-
-
9.0 2.8 2.9 5.2 4.9 3.4 5.2 4.6 6.6 4.1 3.5 3.1 4.1 4.9
7.0 2.4 2.1 3.2 3.3 1.7 2.3 2.1 4.4 2.1 2.3 2.2 2.3 2.5
9.5 3.0 3.0 3.4 3.4 2.4 2.3 2.2 4.0 2.4 2.3 2.0 3.2 3.3
27.5 4.7 5.3 6.4 8.3 5.3 5.2 5.4 10.8 5.8 5.1 5.5 6.6 4.4
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
-
15.0 4.1 2.0 1.7 1.7 3.0 2.0 2.2 2.4 2.3 3.1
13.1 3.6 2.3 2.0 1.7 2.3 2.4 2.2 2.1 2.4 3.1
15.0 3.2 2.3 2.1 1.6 2.8 1.9 2.2 2.4 2.3 2.9
36.0 8.5 5.1 4.8 4.1 5.8 5.1 5.1 5.1 5.9 5.9
-
19.3 23.3 25.6 28.6 17.7 19.1 18.3 38.2 20.2 17.4 17.4 23.6 23.2 168.1 86.1 49.9 41.2 29.9 104.9 29.8 17.6 16.0 13.6 18.8 17.3 18.0 18.1 19.3 21.0 Continued
Table I l l
Continued Nymph development (days)
Temperatureo("C)
28.5 21.3 (J)** 23.1 (F) 26.8 (M) h,
31.O(A)
$
33.3(MY) 33.5 (J) 28.2 (JLY) 26.0 (A) 28.0 (S) 28.2 (0) 26.6 (N) 22.3 (D)
Host Sweet potato Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton Cotton
Egg development (days)
4.7 10-12 1&11 8-10 7-9 7-8 &8 6-7 6-7 6 8 6.0 6-8 8-10
Instar I
Instar 2
lnstar 3
lnstar 4
Egg to adult (total days)
2.4 3.4 3.4 3.4 3.4 2.4 2.3 3.0 3.0 2.3 2.4 3.4 3.4
2.0 7-8 6 8 7-8 5-7 5-7 4-6 4 6 4 5 5 5 6 5.7
2.3 6-8 7-8 7-8 &7 5-6 4 6 4-6 4 5 4 5 4 6 4.6 5.6
4.3 7-8 7-8 7-8 6 7 5-7 5-7 4-6 4-6 4-6 4-7 5-7 6.8
37.0 36.0 35.0 31.0 29.0 27.0 28.0 28.0 29.0 28.0 32.0 35.0
Source
Experimental environment: one asterisk = field or greenhouse conditions; two asterisks = unspecified laboratory conditions; no asterisk = controlled temperature and lighting.
13.
BEMISIA: PEST STATUS, ECONOMICS, BIOLOGY, AND POPULATION DYNAMICS
259
oviposition that occurs on abaxial leaf surfaces appears to involve more cues than gravitational or leaf orientation alone. Experimental manipulation of leaves showed in one study that approximately 80% of eggs were laid on the adaxial surface when the leaves were oriented upward [78]. Using a similar approach, a second study observed from 32 ± 3% to 69 ± 5% of eggs laid on abaxial leaf surfaces turned upward, compared with a range of 86 ± 7% to 97 ± 1% on abaxial leaf surfaces oriented normally [96]. Higher rates of oviposition occur on younger leaves, with later developed nymphs found on progressively older leaves [97, 98]. The degree to which insecticide efficacies are reduced as a result of the underleaf habitat and its potential effect on population development have not been determined.
2.
DISPERSAL
Immigration and emigration are vital components of the population dynamics of B. tabaci. The spatial and temporal relationships of cultivated crops and weed hosts are important factors in colonization by dispersing whiteflies. Unfortunately, as is the case with most insect pest species, it has been difficult to quantify and integrate dispersal parameters, such as the relative numbers of dispersing whiteflies, their reproductive status, their sex ratio, and other biological factors that influence population dynamics. Migrant forms of the A but not the B strain have been identified on the basis of morphological differences as compared with trivial flyers [99]. An unusually high lipid content (40% of total dry weight) suggests that lipids function as an important energy source in longdistance flight [100]. Laboratory studies show takeoff activity between 0600 and 1900 hours, with males flying longer than females. Maximum flight activity occurred between 0600 and 1000 hours and both sexes were capable of sustaining flight for over 2 hours [101]. Greater ascension rates for females than for males may be important in attaining higher altitudes, where wind becomes more of a factor in dispersal. Plant senescence, whitefly age, host plant quality, and population crowding may also affect flight behavior. In the field, distribution patterns have been found to be heavily dependent on wind direction. Numbers of dispersing whiteflies were observed to be concentrated near source fields and distributed bimodally between short-range and long-range fliers, with the greatest distance measured from a source field of 2.7 km [102]. Longrange dispersal studies ofB. tabaci adults based on aircraft net collections have suggested a concentration effect of vertical convection currents and horizontal air movement on whitefly densities [103]. Dispersal from heavily infested cotton and melon fields in Arizona to lettuce fields 1.4 to 4.8 km away resulted in average catches of 384 adults per yellow sticky trap in 30-minute sampling periods during peak dispersal activity [104]. These large-scale movements contributed to population doubling in lettuce in 7 to 10 days.
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Environmental Factors 1.
PLANT CHARACTERISTICS AND HOST PLANT DIVERSITY
Although the number of plant hosts ofB. tabaci is currently listed in excess of 500 [70], there is great variation in preference and infestations on different hosts. For example, in cotton the hirsute leaf character has been reported by many investigators to result in higher B. tabaci populations than the glabrous leaf types. Some authors [87] have hypothesized that lamina trichomes of cotton leaves that originate from elongated epidermal cells overlying leaf veins provide cues to first instar crawlers to locate leaf vascular tissue. Access to phloem tissue is limited by the geometric relationships of the length of the stylet, the abaxial leaf plan, and the distance of vascular tissue from the point of stylet insertion [105]. Stylets of first instar B. tabaci nymphs were reported to be about 80 |im long [106]. In one report, nymphs settled within 60 to 80 jiim of vascular bundle-associated epidermal cells [87], which supports the suggestion that stylet length is a limiting factor in feeding site selection. Okra-leaf cottons have also been found to support lower B. tabaci populations as compared with normal-leaf ones. The difference between the two types has been attributed to smaller leaf area and more open canopy, which provides a less suitable habitat. Differences in utilization of various plant species has long been recognized in B. tabaci. In India during the 1920s and 1930s, B. tabaci was intensively studied as a serious pest of cotton, but it was also observed to colonize other crops and wild hosts throughout the year [28]. In other regions, B. tabaci has been a pest principally on crops other than cotton, such as vegetables in Israel [29], soybeans {Glycine max (L.) Merr. in Brazil [107], and cassava (Manihot esculenta Crantz) in central Africa [108]. In many cases, the relative pest pressure within any particular crop has no doubt involved cultural factors such as the types of crops grown in a particular area and their relative acreages, as well as the cropping sequences and phenologies. However, there are well-documented cases of apparent host specialization among certain populations of B. tabaci. In Puerto Rico, a strain of B. tabaci was identified that colonized only Jatropha spp., whereas the "Sida" strain demonstrated a more typical polyphagous nature but did not include Jatropha spp. [109]. A similar situation was observed in the Ivory Coast, with the "cassava" strain specializing on cassava, but the "okra" strain demonstrating polyphagy exclusive of cassava [108]. A subsequent study conducted in Uganda found a similar difference in host utilization between the specialist cassava strain and the generalist strain, which colonized a wide host range with the exception of cassava [110]. An important difference between the two cassava strains was that the Ugandan population was observed to colonize cotton and sweet potato to a limited extent, whereas the Ivory Coast population was not capable of colonizing any hosts other than cassava [110]. Besides having implications for the epidemiology of the disease caused by African cassava mosaic virus [110], the incom-
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plete specialization of the Ugandan population on cassava points out differences in the capacity of two specialist populations ofB. tabaci, the Ivory Coast cassava strain and the Uganda cassava strain, to utilize a narrow host range. Similar differences in capacity to utilize various hosts have also been routinely observed among polyphagous populations of 5. tabaci. One of the best examples of differences among polyphagous forms was already spelled out in the description of the events that marked the invasion of the B strain into North America beginning in the mid-1980s. Economic infestations of poinsettia in greenhouses and melons in the field were uncommon or unknown for the A strain prior to 1986 but subsequently served to indicate the probable presence of the B strain. Although both A and B strains are widely polyphagous and probably have a similar experimental host range, the natural host range of the B strain appeared to be much greater, based on the larger number of crops and ornamental hosts on which the B strain regularly developed economic infestations. 2.
CROP IRRIGATION AND PLANT NUTRITION
Higher populations of B. tabaci have been associated with increased fertilization of cotton [111, 112] and tomatoes [92]. Although the mechanisms involved in cultural-plant nutrition-^, tabaci interactions are unknown, model simulations for plant insect damage and water stress resulting in reduced photosynthate suggested increases in vegetative growth to be more favorable for B. tabaci population development [90]. Higher populations ofB. tabaci have often been reported on water-stressed cotton [113-115]. Leaf carbohydrate concentrations and honeydew from whiteflies have been shown to be higher in water-stressed melons than in non-water-stressed melons [116]. The largest differences in honeydew carbohydrate concentrations were for glucose and sucrose, suggesting isomerization of simpler sugars as an osmoregulation mechanism. 3.
WEATHER
Extremes in weather conditions appear to play an important role in whitefly population dynamics in some areas [117]. Upper temperature thresholds for growth and development are probably higher than 35°C [74, 83]. Although temperature is a well-documented factor affecting life functions under laboratory conditions, there have been few reports under field conditions except for population reductions following ambient temperatures of 43° to 45°C and low humidity (8-17%) in cotton fields [118]. On eggplant grown under field conditions, adverse effects of low humidity and a lower oviposition threshold of 14°C were reported [29]. Rainfall also has an adverse effect on adult populations [74, 89, 118-120] and oviposition behavior [120]. Wind is an important factor in dispersal, and crops grown downwind in close proximity to infested crops are more likely to be infested than crops at more distant locations [121].
262 4.
TJ. HENNEBERRY AND S J . CASTLE OVERWINTERING
Bemisia tabaci is particularly adapted to tropical and subtropical areas but also is capable of surviving in mild, temperate climates where protected niches are available, under greenhouse conditions, or both [122]. Little emphasis has been placed on overwintering survival and its effects on subsequent summer population development on cultivated crops. Adult and immature populations decrease dramatically, and oviposition is greatly reduced on cultivated crops and weed hosts during the fall and winter months [94, 121-125]. Numerous overwintering hosts have been reported in areas of the world where B. tabaci is an important economic pest (Table IV). In southern California, overwintering B. tabaci and their parasites have been found on Malva parviflora L., a frost-hardy winter weed, on Helianthus annuus L., on Convolvulus arvensis L., and on Lactuca serriola [126-128]. Bemisia tabaci completes development in winter months on carrot (Daucus carota L.), broccoli, squash, eggplant, guar (Cyamopsis tetragonoloba (L.) Taub.), guayule (Parthenium argentatum A. Gray), alfalfa (Medicago sativa L.), and lettuce [94]. At least one generation and a partial second has been observed on lettuce in southern California in winter months. An estimate of the number of empty pupal cases in late autumn indicated that one adult was produced for every 25 mature lettuce leaves, or about 5,000 adults per hectare of lettuce. Empty pupal cases in December on lettuce and reproducing populations on alfalfa, London rocket, Sisymbrian irio L., and alkali mallow {Sida hederocea [Doug.] Terr.) have also been reported in the Yuma Valley of Arizona [121]. Lowlevel egg and nymph populations exist on collards, mustard (Brassica juncea [L.]), canola {Brassica oleraca var. acydela DC), and turnip {Brassica rapa L.) during the winter in South Carolina (temperature range 9.7° to 16.9°C) [122]. One of the major effects of low winter temperatures is the asynchrony of whitefly development and leaf aging [124]. Eggs and larval development slow down or cease at low temperatures while leaf aging continues. Plants vary greatly in their ability to retain leaves under low-temperature conditions. When older leaves die before the completion of 5. tabaci development, high mortality occurs. Heavy rains under low temperatures also increase the mortality of the immature stages [124]. Parasites {Eretmocerus mundus Merc, and Encarsia lutea Masi) have been found throughout the winter and spring in association with overwintering B. tabaci in Israel [125]. Percentages of parasitism were variable, but in some cases more than 50% of the fourth instar B. tabaci nymphs were parasitized. Similarly, Eretmocerus spp. and Encarsia spp. parasites were found throughout the year in the Imperial Valley, California on wild lettuce, Lactuca serriola L., and wild sunflower, Helianthus annuus L. [95]. 5.
NATURAL ENEMIES
Outbreaks in many cases have been associated with the overuse of synthetic organic insecticides and their adverse effects on natural enemy populations [46].
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Table IV
Reported Overwintering Hosts of Bemisia tabaci
Location India
Jordan Sudan Israel
United States Southern California
Southern California Arizona
South Carolina
Source
Overwintering host Rape (Brassica napus L.), cauliflower {Brassica oleracea L.), turnip {Brassica rapa L.), potato (Solanum tuberosum L.), sowthistle {Sonchus spp.), Euphorbia spp., Convolvulus arvensis L. Tomato (Lycopersicon esculentum L.), squash (Cucurbita spp.), potato, cauliflower Ipomoea cardofona Abutilon grandiflorum, Lantana camora L., Chrysanthemum indicum L., Malva parviflora L., Gerbera spp., Solanum vilosum, Withania somnifera (L,), Dunal, Celtis australis, Lonicera estrusca, Verbena spp., Cercis siliquastrum L., Convolvulvus arvensis L., Plumbago europaea, Alcea setosa L., Tropaeolum majus L., Calendula spp., Sonchus oleraceus, Ipomoea batatas L.
[28]
Carrot (Daucus carota L.), broccoli (Brassica oleracea L.), squash (Cucurbita spp.), eggplant (Solanum melongena L.), guar (Cyamopsis tetragonoloba L. Taubert), gua5aile (Parthenium argentatumA. Gray), alfalfa (Medicago sativa L.), lettuce (Lactuca sativa L.) Malva parviflora L., Helianthis annuus L., Convolvulus arvensis L., Lactuca serriola L. Alfalfa, London rocket Sisymbrian irio L., alkaU mallow (Sida hederocea Doug. Torr.); okra, Abelmoschus esculentus L., squash (Cucurbita spp.) watermelon, Citrullus lanatus (Thunb.), cantaloupe (Cucumis melo L.), broccoli, cauliflower, spiny sowthistle (Sonchus asper (L.)) Hill Collard (Brassica oleraca var acydela DC), mustard (Brassica juncea (L.)), canola (Brassica rapa L.)
[94]
[124] [88] [125]
[126, 127, 128] [121, 127, 128]
[122]
Parasitism of fourth instar nymphs as high as 70-80% in southern Cahfornia cotton has been reported under full-season production [126, 129, 130]. Under shortseason cotton production in the Imperial Valley of California parasitism seldom reaches 40% [131], although parasitism rates higher than 80%o have been observed [132]. Early-season parasites in southern California have not sufficiently controlled B. tabaci population growth, but Eretmocerus parasitism always exceeded that of Encarsia [95]. Similarly, in Israel, peak populations of Encarsia lutea (IVlasi) and Eretmocerus mundus (Mercet) occur in early and midSeptember, respectively, following B. tabaci peak populations [133]. In most cases in the United States and Israel, indigenous species have rarely provided con-
264
T.J. HENNEBERRY AND S.J. CASTLE
trol, and additional inputs have been necessary to suppress populations [131, 134]. One exception, in the United States, was the reported high level of parasitism in peanuts [135]. In the Sudan, the highest parasitism of B. tabaci was reported to be due to E. lutea {66%) followed by E. mundus (34%) [136]. A subsequent study from the Sudan reported that B. tabaci was effectively controlled by natural enemies in untreated cotton fields [137]. In Syria, natural enemies appear to be effective at controlling B. tabaci in untreated cotton, but it becomes a major cotton pest where insecticides are used [138]. In Egypt, average percentages oiB. tabaci parasitism by E. mundus in insecticide-treated cotton and cabbage ranged from 44.4 to 73.0% and from 34.0 to 55.4%), respectively, and in untreated Lantana camara L from 78.6 to 80.8%, suggesting that E. mundi had a major role in regulating populations [139]. Similar results in Egypt were reported on cotton, soya, cauliflower, and tomato [140] and a number of other vegetable crops [141]. Numerous predaceous arthropod species have been reported to attack B. tabaci. A recent review of predaceous arthropods of eggs, nymphs, and adults lists 13 identified and 1 unidentified predaceous mite species, 12 species of spiders (5 undetermined species), 16 Coleoptera, 3 Diptera, 13 Hemiptera, 1 Hymenoptera, 8 Neuroptera (1 unidentified species), and 1 unidentified species of Thysanoptera [142]. Quantification of the impact on whitefly population development has been difficult because predators rarely leave evidence of attack. A major advance in our ability to evaluate predation on whitefly populations was the development of a pest-specific monoclonal antibody for detection of B. tabaci types A and B [143] in predator gut contents. Of nine heteropteran predator species collected from field populations in cotton, 4.0 to 39.4%) tested positive for B. tabaci egg antigens [144, 145]. Of two coleopterous predators, 58%) of the Collops vittatus (Say) and 38%) of the Hippodamia convergens Guerin-Meneville tested positive for 5. tabaci [145]. The most commonly observed fungal pathogens of B. tabaci and/or other whiteflies are Paecilomyces fumosoroseus (Wize), Verticillium leucanii (Zimmerman), and Aschersonia aleyrodides Webber [146]. Under natural conditions, P. fumosoroseus epizootics have occasionally been observed to decimate B. tabaci populations. Overall, little is known regarding the role of fungi in regulating populations. 6.
INSECTICIDE RESISTANCE
In some areas of the world, resistance to all major chemical groups (chlorinated hydrocarbons, pyrethroids, organophosphates, carbamates, cyclodienes) and the insect growth regulators buprofezin and pyriproxyfen has been reported [8]. Resistance to all types of chemicals may not exist in any one area. No predictable relationship has been found between the frequency or patterns of resistance and biotypes. Quantification of the impact of resistance on population dynamics and pest status is notably lacking [8]. However, the interactions between insecticide
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resistance and B. tabaci fertility stimulation by DDT have been suggested as the most probable causes of epidemic populations in the Sudan in the 1970s [9, 10]. Generalizations regarding the role of resistance as a factor in the increasing problems, however, are somewhat risky. For example, in the Imperial Valley of California, insecticides have been used intensively against B. tabaci populations for many years and particularly since the 1991 outbreak [3]. Insecticide resistance monitoring between 1992 and 1995 failed to detect progression to higher levels of resistance to many pyrethroid and organophosphate compounds [147, 148]. Although good protection of crops was attainable in closely managed fields, the Imperial Valley consistently produced high populations each year on a regional basis. This appeared to be more related to a highly favorable growing environment than to inability to control infestations with insecticides. It also reflected the capacity ofB. tabaci to capitalize on the wide assortment of host crops, including 200,000 acres of alfalfa, which probably played an important role in increasing populations as well as in acting as a refuge from insecticides and helping to sustain susceptibility levels to insecticides. D.
Life Tables and Population Models
Under field conditions on cotton, immature-stage B. tabaci mortality was found to be highest during the crawler and second-instar nymphal stages [149]. About 17% of the eggs survived to adults in the field as compared with 75% in the laboratory. First-instar mortality was attributed primarily to climatic factors (e.g., low humidity and high temperatures) and predation by lacewings, spiders, and Orius spp., but also to host plant effects. The "moderate" mortality that incurred in other nymphal instars and pupae also was attributed to weather and parasitism. In the laboratory, temperature and cotton leaf age have been reported as the most significant factors affecting reproduction [85]. Under greenhouse conditions, dramatic differences in egg and nymphal mortalities associated solely with plant effects were found on peppers, cotton, and melons. Except for small nymphs on cotton, mortalities were greater in all cases on pepper, followed by cotton and then melons [150]. In the laboratory, increasing temperatures within the range of 16° to 28°C shortened developmental times, adult longevity, and preoviposition time, reduced immature mortality, and increased oviposition on poinsettia [76]. This resulted in a faster generation time (169 days at 16°C versus 32 days at 28°C) and population doubling time (550 days at 16°C versus 6.0 days at 28°C), and increased both the numbers per generation and the intrinsic rate of population increase (0.001 at 16°C to 0.126 at 28°C) (Table V). On eggplant, at temperatures ranging from 20° to 35°C, decreases in intrinsic rates of increase occurred along with increases in generation doubling times at 30° and 35°C compared with 27°C [83]. At 25°C, population doubling time was approximately 8 days on poinsettia as compared with approximately 4 on eggplant. Reported intrinsic rates of increase of 0.153, 0.138, 0.131, and 0.120 occurred on tomato, sweet potato, cucumber, and garden
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TJ. HENNEBERRY AND S.J. CASTLE
bean at 25°C [84], compared with 0.09 on poinsettia [76], and generation doubling times at 23° to 27°C ranged from 3.7 to 5.7 days, respectively. Model development for simulating B. tabaci populations has been slow, and progress is limited by a lack of information from multicropping systems. However, a model has been developed that includes temperature-dependent development and survival of all stages and age groups, temperature- and nutrition-dependent oviposition effects (e.g., changes in leaf age), parasitism, and dispersal [85, 90]. The population growth model simulates exponential population growth, and the cotton model interface describes increasing leaf mass production; leaf, stem, and fruit dry-matter production; and cumulative density of fruiting forms. Simulations showed that late-season 5. tabaci population declines were attributable primarily to a decline in host plant quality, emigration, and cooler temperatures. Parasitism appeared to contribute to some of the late-season mortality and population decline. A degree-day (lower [10.0°C] and upper [32.2°C] thresholds) exponential population growth-rate equation was used in a B. tabaci model in California and assumed an average accumulation of 316 degree-days between field generations [151]. The model is very sensitive to the accuracy of initial population density estimates but has been useful in describing population growth in California.
v. Concluding Remarks The biological characteristics of B. tabaci, including high reproductive rates, polyphagy, and dispersal, interact with climatic inputs within multicrop environments to drive its sometimes explosive population dynamics. These factors have been studied extensively throughout the world in the field and under laboratory conditions. From this collective body of information, some threads of continuity have been revealed, which suggest a commonality of factors that influence the population dynamics of the B, tabaci complex throughout its geographical range. Laboratory life table results show potential population doubling times of 4 to 10 days at temperatures ranging from 25° to 35°C on six different hosts used in each study [76, 83, 84]. This is well within the temperature conditions of most cultivated crop and greenhouse growing seasons. Thus, inherent B. tabaci biological and reproductive abilities expressed under favorable environmental conditions can result in devastating population levels on most cultivated crops. Although universally accepted as having an important role in B. tabaci bionomics, quantification of the impact of natural enemies has been difficult to achieve. Even though high percentages of parasitism have been recorded in certain crops such as cotton, and predators such as Orius spp. and Geocoris spp. have been positively identified for predation [144,145], most reviews indicate that natural enemies independently have not achieved acceptable control in the field in
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the United States, Israel, or Argentina [120, 131, 134]. In contrast, indigenous natural enemy populations have been reported to provide effective control in the Sudan [137], Egypt [139-141], and Syria [138]. The differences have been reported to occur because of insecticide effects on natural enemies under intense crop systems, as opposed to less intense crop systems, in which fewer insecticides are used and natural enemies readily contain infestations. However, this contention has rarely, if ever, been supported by rigorous field data. Natural enemy impact on population development in most crop systems warrants integrated pest management practices with reduced insecticide inputs to optimize natural enemy control ofB. tabaci. The spatial and temporal relationships of cultivated and weed hosts, large area monocultures, and crop production imputs such as fertilizer and irrigation have all been shown to have a significant impact on B. tabaci population dynamics. The importance of these factors in limiting or encouraging natural enemy populations, however, is virtually unknown. Higher populations of B. tabaci occur in waterstressed cotton and in cotton with increased fertilizer (N, P, K) inputs. The inter- and intracrop dispersal ofB. tabaci in conjunction with its wide host range encourages population expansion on a continuum of food sources and crop habitats. Dispersal from weeds and winter crops to spring cucurbits and summer cotton, followed by a return in autumn to short-season vegetable crops and weeds, has been reported from many parts of the world [28, 152-154]. In most areas of epidemic outbreaks, the largest acreages and most suitable microhabitats for B. tabaci, occur within the cotton crop. Additionally, high insecticide use in cotton discourages natural enemy populations and often leads to insecticide resistance. Although insecticide resistance has been well documented in B. tabaci populations around the world, there has been little quantification of its impact on B. tabaci population dynamics [9]. In Sudan, heavy insecticide use reportedly led to resistant B. tabaci populations but also acted to stimulate fertility levels of B. tabaci, these effects together acting to increase populations to outbreak levels. Although high levels of resistance to insecticide were documented in the Imperial Valley in the 1980s, insecticide resistance has not been an overwhelming factor in the 1990 outbreaks, although it may have contributed to problems in Arizona. Concern over the increasing pest status of B. tabaci began to surface with the increasing frequency of severe B. tabaci outbreaks in cotton in India in 1919, Sudan in 1950, Iran in 1955, Nicaragua in 1967, Turkey in 1974, Israel in 1976, and the United States in the late 1970s. Thus cotton as a crop and the evolving production methodology and technology may have favored conditions for B. tabaci exploitations. In conjunction with logarithmic population development on cotton, an increase in ability to exploit commercially produced cole crops, alfalfa, and melons may be of particular importance in some ecosystems [155]. The early spring melon crop in California and Arizona becomes infested from B. tabaci that has overwintered on weeds, ornamentals, or cole crops. Recently, it has been suggested that alfalfa may also play a role in contributing reproductive biomass throughout the year [156]. Thus, the
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T.J. HENNEBERRY AND S.J. CASTLE
potential for higher overwintering populations has been increased dramatically, at least in California and Arizona, with crops that bridge the gap between annual plantings of large, monocultured cotton crops. Overwintering survival is a major population regulating factor, with annual decreases in B. tabaci populations coinciding with declining temperatures and reduced cotton acreages. It is unlikely that any one factor can be found to explain the occurrence oiB. tabaci epidemics in all instances for any given area, and much less so on a global basis. It is likely that a combination of factors are involved in most instances in which unmanageable B. tabaci populations occur. The population dynamics of 5. tabaci become more complex with overlapping generations and in multicrop environments. A more complete understanding at the agroecosystem level of B. tabaci population dynamics is a goal for the future.
References 1. Gonzalez, R.A., Goldman, G.E., Natwick, E.T., Rosenberg, H.R., Grieshop, J. I., Sutter, S.R., Fanakoshi, T., and Davila-Garcia, S. (1992). Whitefly invasion in Imperial Valley cost growers, workers millions in losses. Calif. Agr. 46, 7-8. 2. Norman, J.W., Jr., Sparks, A.N., Jr., and Riley, D.G. (1995). Impact of cotton leaf-hairs and whitefly populations on yields in the Lower Rio Grande Valley. In "Proceedings of the Cotton Production and Research Conference, National Cotton Council of America" (D. A. Richter and J. Armor, eds.), pp. 102-104. Memphis, TN. 3. Perring, T.M., Cooper, A., Kazmer, D.J., Shields, C , and Shields, J. (1991). New strain of sweetpotato whitefly invades California vegetables. Calif. Agr. 45, 10-12. 4. Gill, R.J. (1992). A review of the sweetpotato whitefly in Southern Cahfomia. Pan-Pac. Entomol.6H, 144-152. 5. Polston, J.E., and Anderson, P.K. (1997). The emergence of whitefly-transmitted geminiviruses in tomato in the Western Hemisphere. Plant Dis. 81, 1358-1369. 6. Polston, J.E., Bois, D., Carmona-Serra, A., and Concepcion, S. (1994). First report of tomato yellow leaf curl-like geminivirus in the Western Hemisphere. Plant Dis. 78, 831. 7. Dittrich, V, Ernst, G.H., Ruesch, O., and Uk, S. (1990). Resistance mechanisms in sweetpotato whitefly (Homoptera: Aleyrodidae) populations from Sudan, Turkey, Guatemala and Nicaragua. J. Econ. Entomol. 83, 1665-1670. 8. Denholm, I., Cahill, M., Byrne, F.J., and Devonshire, A.L. (1996). Progress with documenting and combating insecticide resistance in Bemisia. In "Bemisia 1995: Taxonomy, Biology, Damage, Control and Management" (D. Gerling and R.T. Mayer, eds.), pp. 577-603. Intercept Ltd., Andover, Hampshire, U.K. 9. Dittrich, V, Hassan, S.O., and Ernst, G.H. (1985). Sudanese cotton and the whitefly: a case study of the emergence of a new primary pest. Crop Protect. 4, 161-176. 10. Byrne, F.J., Cahill, M., Denholm, I, and Devonshire, A.L. (1995). Biochemical identification of interbreeding between B-type and non-B-type strains of the tobacco whitefly Bemisia tabaci. Biochem. Genet. 3, 13-23. 11. Schuster, D.J., Stansly, P. A., and Polston, J.E. (1996). Expressions of plant damage by Bemisia. In "Bemisia 1995: Taxonomy, Biology, Damage, Control and Management" (D. Gerling and R.T. Mayer, eds.), pp. 153-177. Intercept Ltd., Andover, Hampshire, U.K.
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133. Gerling, D., Motro, U, and Horowitz, R. (1980). Dynamics of Bemisia tabaci (Gennadius) (Homoptera: Aleyrodidae) attacking cotton in the coastal plain of Israel. Bull. Entomol. Res. 70, 213-219. 134. Gerling, D. (1986). Natural enemies of Bemisia tabaci: Biological characteristics and potential as biological control agents—a review. Agr. Ecosys. Environ. 17, 99-110. 135. McAuslane, H.J., Johnson, F.A., Knauft, D.A., and Colvin, D.L. (1993). Seasonal abundance and within-plant distribution of parasitoids of Bemisia tabaci (Homoptera: Aleyrodidae) in peanuts. Environ. Entomol. 22, 1043-1050. 136. Gameel, O.I. (1969). Studies on whitefly parasites Encarsia lutea and Eretmocerus mundus Mercet (Hymenoptera: Aphelinidae). Rev. Zool. Bot. Afr. 79, 65-77. 137. Abdelrahman, A.A., and Munir, B. (1989). Sudanese experience in integrated pest management of cotton. Insect Sci. Appl. 10, 787-794. 138. Stam, P. A., and Elmosa, H. (1990). The role of predators and parasites in contolling populations of Earias insulana, Heliothis arrnigera and Bemisia tabaci on cotton in the Syrian Republic. Entomophaga 35, 315—327. 139. Hafez, M., Awadallah, K.T., Tawfik, M.F.S., and Sarhan, A.A. (1979). Impact of the parasite Eretmocerus mundus Mercat on population of the cotton whitefly, Bemisia tabaci (Genn.) in Egypt. Bull. Entomol. Soc. Egypt 62,23-32. 140. Abdel-Fattah, M.I, Hendi, A., and El-Said, A. (1986). Ecological studies on parasites of the cotton whitefly 5emwzfl tabaci (Genn.) in Egypt. Bull. Entomol. Soc. Egypt Econ. Sen 14, 95-105. 141. Abdel-Gawaad, A.A., El Sayed, A.M., Shalaby, F.E, and Abo-El-Ghar, M.R. (1990). Natural enemies of Bemisia tabaci Genn. and their role in suppressing the population density of the pest. Agr Res. Rev 6H,n5-l95. 142. Nordlund, D.A., and Legaspi, J.C. (1996). Whitefly predators and their potential for use in biological control. In ''Bemisia: 1995 Taxonomy, Biology, Damage, Control and Management" (D. Gerling and R.T. Mayer, eds.), pp. 499-513. Intercept Ltd., Andover, Hampshire, U.K. 143. Hagler, J.R., Brower, A.G., Tu, Z., Byrne, D.N., Bradley-Dunlop, D., and Enriquez, FJ. (1993). Development of a monoclonal antibody to detect predation of the sweetpotato whitefly, Bemisia tabaci. Entomol. Exp. Appl. 68, 231-236. 144. Hagler, J.R., and Naranjo, S.E. (1994). Qualitative survey of two coleopteran predators of Bemisia tabaci (Homoptera: Aleyrodidae) and Pectinophora gossypiella (Lepidoptera: Gelechiidae) using a multiple prey gut content ELISA. Environ. Entomol. 23, 193-197. 145. Hagler, J.R., and Naranjo, S.E. (1994). Determining the frequency of heteropteran predation on sweetpotato whitefly and pink bollworm using multiple ELISA's. Entomol. Exp. Appl. 72,59-66. 146. Lacey, L.A., Fransen, J.J., and Carruthers, R. (1996). Global distribution of naturally occurring fungi of Bemisia, their biologies and use as biological control agents. In "Bemisia 1995. Taxonomy, Biology, Damage, Control and Management" (D. Gerling and R.T. Mayer, eds.), pp. 401^33. Intercept Ltd., Andover, Hampshire, U.K. 147. Castle, S., Henneberry, T.J., Toscano, N., Prabhaker, N., Birdsall, S., and Weddle, D. (1996). Silverleaf whiteflies show no increase in insecticide resistance. Calif. Agr 50, 18-23. 148. Castle, S.J., Henneberry, T.J., Prabhaker, N., and Toscano, N.C. (1996). Trends in relative susceptibilities of whiteflies to insecticides through the cotton season in the Imperial Valley, California. In "Proceedings of the Beltwide Cotton Production and Research Conference" (P. Dugger and D.A. Richter, eds.), pp. 1032-1035. National Cotton Council of America, Memphis, TN. 149. Horowitz, A.R., Podoler, H., and Gerling, D. (1984). Life table analysis of the tobacco whitefly Bemisia tabaci (Gennadius) in cotton fields in Israel. Acta Oecol. Oecol. Appl. 5, 221-233. 150. Riley, D., Nava-Camberos, U, and Allen, J. (1996). Population dynamics of Bemisia in agricultural systems. In ''Bemisia: 1995 Taxonomy, Biology, Damage, Control and Management" (D. Gerling and R.T. Mayer, eds.), pp. 93-109. Intercept Ltd., Andover, Hampshire, U.K.
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151. Zalom, EG., Natwick, E.T., and Toscano, N.C. (1985). Temperature regulation of Bemisia tabaci (Homoptera: Aleyrodidae) populations in Imperial Valley cotton. J. Econ. Entomol 73, 61-64. 152. Johnson, M.W., Toscano, N.C, Reynolds, H.T., Sylvester, E.S., Kido, K., and Natwick, E.T. (1982). Whiteflies cause problems for southern CaUfomia growers. Calif. Agr. 36, 24—26. 153. Melamed-Madjar, V, Cohen, S., Chen, M., Tam, S., and Rosilio, D. (1979). Observations on populations of Bemisia tabaci Gennadius (Homoptera: Aleyrodidae) on cotton adjacent to sunflower and potato in Israel. Israeli Entomol 18, 71-78. 154. Nachapong, M., and Mabbett, T. (1979). A survey of some wild hosts of Bemisia tabaci Genn. around cotton fields in Thailand. Thai J. Agr. Sci. 12, 217-222. 155. Toscano, N.C, Henneberry, T.J., and Castle, S. (1994). Population dynamics and pest status of silverleaf whitefly in the USA. Arab J. Plant Protect. 12, 137-142. 156. Yee, W.L., and Toscano, N.C. (1996). Ovipositional preference and development of Bemisia argentifolii (Homoptera: Aleyrodidae) in relation to alfalfa. J. Econ. Entomol. 89, 870-876. 157. El-Kihidir, E. (1965). Bionomics of the cotton whitefly {Bemisia tabaci Genn.) in the Sudan and the effects of irrigation in population density of whiteflies. Sudan Agr J. 1, 8-22. 158. Hassan, O. (1982). Investigations of DDT residues on cotton leaves on the fertility of the cotton whitefly Bemisia tabaci (Gennadius) (Homoptera: Aleyrodidae). M.S. Thesis, University of Reading, U.K. 37 pp. 159. Hendi, A., Abdol-Fahah, M.I., and El-Sayed, E. (1987). Biological study on the whitefly, Bemisia tabaci (Genn.) (Homoptera: Aleyrodidae). Bull. Entomol. Soc. Egypt 5, 101-108.
CHAPTER 14
Whitefly-Borne Viruses in Continental Europe PlERO C.
CACIAGLI
/- Introduction In a review on whitefly transmission of plant viruses in 1987, Duffus [1] pointed out that whiteflies, as vectors of disease agents, "although not considered as important as aphids on a worldwide basis," caused significant losses throughout the world, and that there had been an increasing awareness of losses caused by whitefly-transmitted viruses in temperate areas. In 1999, a search on a bibliographic database such as CABPEST CD shows that the number of articles dealing with whiteflies and viruses almost doubled in the last 10 years, whereas the number of articles on aphids and viruses remained almost constant. About one-tenth of the papers on whitefly-bome viruses discuss problems within Europe. The increasing menace of whitefly-borne viruses caused the European Union (EU) to set up in 1999 the "European network on European whiteflies, their associated plant pathogens and disorders," or more briefly, the European Whitefly Studies Network (EWSN). Geographical Europe includes many more countries than those in the EU, and whitefly-borne viruses and their vectors tend to ignore borders, being more affected by geographical than political barriers. These are the reasons why, in this review, I have included within continental Europe countries extending over the Caucasus mountains (traditionally, one of the geographical limits of Europe) and Turkey, which geographically is mostly in Asia.
Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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//. A.
Virus and Virus-Like Diseases
Closteroviridae
Members of the family Closteroviridae [2] have long, highly flexuous particles with distinct cross-banding. Virions contain linear, positive-sense, ssRNA [3]. Closteroviruses may have either monopartite or bipartite genomes, but both types encode for a heat-shock protein (HSP-70) homolog [4]. Table I summarizes the closteroviruses known to be present in Europe. 1.
BEET PSEUDO-YELLOWS CLOSTEROVIRUS
Beet pseudo-yellow virus (BPYV) was the first whitefly-borne closterovirus described [5, 6]. The particles measure 900-950 nm in length [4] and 12 nm in diameter. This virus is transmitted by the greenhouse whitefly Trialeurodes vaporariorum, has a broad host range of crops, weeds, and ornamentals, including lettuce (Lactuca sativa), endive (Lactuca serriola), melon (Cucumis melo), and cucumber (Cucumis sativus), but excluding tomato (Lycopersicon esculentum) [4, 5]. Plants are generally stunted; leaf symptoms range from chlorotic spotting to interveinal yellowing. Leaves of diseased plants are thickened and brittle [5], a characteristic common to other whitefly-bome closteroviruses. Beet pseudo-yellows virus, currently placed in the genus Closterovirus, has a number of features of the new genus Crinivirus [2], and its status may change as soon as information on its genome organization is available.
Table I
Whitefly-Bome Closteroviruses in Europe
Virus Beet pseudo-yellows closterovirus (BPYV)
As melon yellows virus As muskmelon yellows virus As muskmelon yellowing virus As cucumber chlorotic spot virus As cucumber infectious chlorosis virus Tomato infectious chlorosis crinivirus (TICV) Cucurbit yellow stunting disorder crinivirus (CYSDV)
Country The Netherlands France Belgium Italy United Kingdom Turkey Greece and Crete Spain France Spain France Bulgaria Italy Spain Turkey
Reference(s) [7] [8] [17] [18] [19] [20] [21,22] [12, 13, 15] [4,9] [14] [10,11] [4, 16] [25, 26] [11,22,28] [22]
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281
In Europe, BPYV was found almost contemporaneously in the Netherlands [7] and France [8] in 1980. It was again reported in France in 1982, when it was described as muskmelon yellows [9], now recognized as BPYV [4]. Cucumber chlorotic spot virus, obtained from cucumber in France in 1992 [10], is also now considered an isolate of BPYV [11]. In 1982 BPYV was found in Spain and reported as melon yellows [12], then in 1993 as muskmelon yellowing [13], and again in 1999 as melon yellows [14]. In 1991, BPYV was identified as causing the latter diseases[15]. In Bulgaria, BPYV was reported in 1986 [16] as cucumber infectious chlorosis virus, now recognized as BPYV [4]. In Belgium, Italy, and the United Kingdom, BPYV was found and correctly identified and reported in 1987 [17], 1989 [18], and 1990 [19]. The relationships between different isolates of yellowing viruses from cucumber and the original North American BPYV have been studied and clarified by Coffin and Coutts [20] using reverse-transcriptase polymerase chain reaction (RT-PCR) techniques. An isolate from Turkey was included in this research because Turkey is among the countries where BPYV is present. This virus has also been detected in continental Greece [21] and in Crete [22]. 2.
TOMATO INFECTIOUS CHLOROSIS CRINIVIRUS
Tomato infectious chlorosis virus (TICV) in the Closteroviridae has a bipartite genome [23] and particles with a modal length of 850 to 900 nm. Transmitted by T. vaporariorum, TICV was originally isolated in California in 1993 from tomato plants with interveinal yellowing, necrosis, and leaf brittleness [24]. It has a relatively narrow host range, which includes potato (Solarium tuberosum), lettuce, and artichoke (Cynara scolymus) but not cucurbits [24]. A very similar virus, almost contemporarily isolated in northwest Italy (Riviera) from tomato plants, was reported in 1995 [25] and identified as TICV in 1996 [26]. Since then, the virus has also been detected in naturally infected artichokes in the same area (Caciagli, unpubHshed). 3.
CUCURBIT YELLOW STUNTING DISORDER CRINIVIRUS
Cucurbit yellow stunting disorder virus (CYSDV) was first found in the Arab Emirates in 1982 and described in 1990 [27]. A Spanish isolate of CYSDV has been characterized by Celix et al [28] and its presence in southern Spain was again reported in 1999 [11, 22]. The virus has a bipartite genome, is transmitted 'in a semipersistent manner by Bemisia tabaci biotypes B and Q [11], and is restricted to Cucurbitaceae [28], in which its symptoms are very similar to those of BPYV [4]. It has also been detected in cucurbits from Turkey [22]. Symptoms of whitefly-borne closteroviruses on hosts they have in common are not very distinctive; nevertheless, biological differentiation of the viruses found in Europe is easy (Table II). There are additional ways to identify these and other
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Table II
Biological Differentiation of Whitefly-Bome Closterovimses Found in Europe Virus
Host or vector Curcurbit Tomato Lettuce Vector
Beet pseudoyellows virus (BPYV)
Tomato infectious chlorosis virus (TICV)
Yes No Yes T. vaporariorum
No Yes No T. vaporariorum
Cucurbit yellow stunting disorder virus (CYSDV) Yes No No B. tabaci (biotype B)
related viruses not yet known in Europe: lettuce infectious yellows virus (LIYV), lettuce chlorosis virus (LCV), tomato chlorosis virus (ToCV), and the whiteflyborne viruses infecting sweet potato [4]. Antisera are available against BPYV [6], TICV [24, 25], and LIYV [29] for enzyme-linked immunosorbent assay (ELISA) or Western blot tests. Degenerate primers have been designed from deduced amino acid sequences for the conserved phosphate 1 and 2 motifs of the virusencoded HSP-70 homologue [30] and used in reverse transcription-polymerase chain reactions (RT-PCRs) to produce species-specific cDNAs for several whitefly- and aphid-borne viruses in Closteroviridae. These primers provide a good general tool for detecting whitefly-bome closteroviruses, although there is at least one such virus not amplified with these primers [31]. B.
Geminiviridae
Member viruses of the family Geminiviridae have geminate particles measuring 30 nm long and 18-20 nm wide, with icosahedral symmetry. They possess a closed, circular, single-stranded, monopartite or bipartite DNA genome [3]. All geminiviruses found in Europe belong to the genus Begomovirus [2], type species bean golden mosaic virus. Some begomoviruses are mechanically transmissible; all have the whitefly B. tabaci as vector, if any, and a narrow host range in the dicotyledonous plants [3]. Begomoviruses known to be present in Europe are Usted in Table III. 1.
ABUTILON MOSAIC BEGOMOVIRUS
Abutilon mosaic virus (AbMV), known since 1906 (Bauer, cited in [32]), was classified as a geminivirus in 1986 [32]. Isolates of AbMV are known from different parts of Europe [33, 34], but the decoratively infected plant of Abutilon pictum 'Thompsonii' is so widely grown as ornamentals that it is useless to try to map its presence in different countries. The threat of AbMV to European crops is probably low; attempts to transmit European isolates by B. tabaci have thus far been unsuccessful [35, and Caciagli, unpublished].
14.
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WHITEFLY-BORNE VIRUSES
Table III
Whitefly-Borne Geminiviruses in Europe
Virus
Country
Abutilon mosaic virus (AbMV)
Reference(s)
X15983, X15984«
Germany Scotland England
Honeysuckle yellow vein mosaic virus (HYVMV) Ipomoea yellow vein virus (lYW) Tomato yellow leaf curl virus-Sardinia (TYLCV-Sar) Tomato yellow leaf curl virus-Israel (TYLCV-Is)
Present in many countries Great Britain Present in many countries Spain Italy Spain Cyprus Turkey Portugal Spain
Sequence accession EMBL/GenBank
AF058013^ AF058014^ [35, 36] [42]
AJ132548
[49-51] [52, 53] [46] [47] [55] [56]
X61153,Z28390 Z25751,L27708 XI5656^
AF022219 (VI), AF022220(C2)
" Sequence of the West Indian isolate present in Germany. * Partial sequences of the coat protein (AVI) genes from two unpublished Rothamsted field strains. ^ Sequence of the original isolate from Israel, not a European isolate.
2.
HONEYSUCKLE YELLOW VEIN IMosAIC BEGOMOVIRUS
The presence and distribution of honeysuckle with yellow vein mosaic are similar to that of abutilon mosaic. Plants of Lonicera Japonica var. aureo-reticulata with bright yellow mosaic are widely grown as ornamentals. In the plants tested in Europe, symptoms are associated with honeysuckle yellow vein mosaic virus (HYVMV) [36]. Yellow vein symptoms in L. japonica can also be caused by tobacco leaf curl begomovirus (TLCV) [37-39]. Japanese isolates of TLCV from honeysuckle are transmitted by B. tabaci [37, 38], whereas the tested isolate of HYVMV is not [35]. For a while HYVMV was considered synonymous with TLCV [37, 40]; however, they are now listed as different viruses in the same genus [41]. 3.
IPOMOEA YELLOW VEIN VIRUS
One more geminivirus, provisionally named Ipomoea yellow vein virus (lYVV), has been recently detected in vegetatively propagated Ipomoea indica in Spain [42]. The virus is associated with yellow vein symptoms and is not transmissible by B. tabaci biotypes B, Q, and S. It has 70.8% identity to the ACl gene
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Table IV
Unidentified Whitefly-Bome Disease Agents Reported in Europe.
Unidentified agent
Country
Maple mosaic Citrus chlorotic dwarf
Hungary Turkey
Vector Trialeurodes vaporariorum Parahemisia myricae
Reference(s) [61] [62,63]
of Agemtum yellow vein virus and the highest homology to the coat protein gene of tomato leaf curl virus from southern India [42]. 4.
TOMATO YELLOW LEAF CURL BEGOMOVIRUSES
Tomato yellow leaf curl as an infectious disease was described in Israel in the 1960s [43, 44]. Its etiology was linked to tomato yellow leaf curl begomovirus in 1988 [45]. In the meantime, TYLCV had been detected in Cyprus in 1985 [46] and in Turkey [47]. Since then, a number of viruses with similar characteristics have been characterized, so that there are now two different tomato yellow leaf curl viruses recognized in Europe, TYLCV-Is from Israel and TYLCV-Sar from Sardinia [41]; TYLCV-Is [48] and TYLCV-Sar [49] both have monopartite genomes, cause typical curling and yellowing of tomato leaves, have overlapping host ranges, and are transmitted by B, tabaci. a. TYLCV-Sar. TVLCV-Sar was identified in Italy in 1988 [50, 51], and in Spain in 1994 [52, 53]. A typical isolate does not infect French bean (Phaseolus vulgaris) [54]. b. TYLCV-Is. TYLCV-Is was identified as such in Portugal in 1996 [55] and then in Spain [56], where it also causes a severe disease of French bean [57, 58]. The viruses detected in Cyprus [46] and Turkey [47] are suppose4 only for geographical reasons, to be TYLCV-Is. The geminiviruses already identified in Europe can be distinguished by their biological properties, but this may not be the easiest way. Degenerate primers from highly conserved regions of the Begomovirus genome are usefiil in PCR detection of geminiviruses, even when their genomes are not well characterized [59, 60]. Although restriction fragment length polymorphism analysis seems to be more useful in identifying geminiviruses from the Americas [59], it can also distinguish among some isolates from Africa and Europe [60]. For final identification of known viruses, a panel of monoclonal antibodies, used in the correct combination, can identify a number of viruses in the genus Begomovirus [34, 36]. C.
Unidentified Whitefly-Borne Agents
Two other whitefly-associated diseases have been reported from Europe (Table IV). Their causal agents are unknown, but suspected of being viruses.
14.
WHITEFLY-BORNE VIRUSES
1.
285
MAPLE MOSAIC DISEASE
A disease of maple (Acer negundo mid Acer pseudoplatanus) with symptoms of ochre-yellow mosaic, leaf deformation, and irregular incisures of the leaf blade was described in 1972 in Hungary. The disease develops in about 10 years through pronounced shortening of intemodes of adventitious shoots to a broomlike appearance of the crown of the tree [61]. The disease agent has been transmitted to A. negundo by T. vaporariorum collected from diseased plants or reared on healthy tomato plants before exposure to diseased plants and transfer to test plants [61]. 2.
CITRUS CHLOROTIC DWARF DISEASE
An infectious chlorosis of citrus found in the eastern Mediterranean region of Turkey in the 1980s reached epidemic levels in the 1990s and was described in 1994 by Cinar et al [62]. Symptoms consist of V-shaped notches in leaves, crinkling and upward cupping of leaves, and variegation, particularly on lemon. The disease agent is transmitted, although inefficiently, by stem slash. The Japanese bayberry whitefly, Parabemisia myricae (Kuwana), a newcomer among virus vectors, transmits the disease agent in a semipersistent or persistent manner [63]. This completes the inventory of whitefly-bome viruses or suspected viruses identified in Europe. There is no doubt that others will be found or reported in new areas because, as we shall see, the whitefly vectors are present in regions of European where no whitefly-bome virus or disease has yet been reported.
///. Vectors Since the spread of viruses and virus-like diseases often depends on the presence of their vectors, we shall look at locations in which vectors of whitefly-borne disease have been found in Europe (Table V). A.
Trialeurodes vaporariorum
The most common and most diffuse of the whitefly vectors of viruses in Europe is certainly the greenhouse whitefly, T. vaporariorum Westwood. At present, this whitefly is so common all over the world that there is no off'icial distribution map for it. In Europe, T. vaporariorum has been reported as an "important pest" as far north as Scotland [64] and Norway [65]. The distribution limit of Z vaporariorum seems to be greenhouses more than anywhere else. In southern Europe its presence is not limited to protected crops. B.
Parabemisia myricae
The Japanese bayberry whitefly P. myricae wa,s first described in Japan in 1927 (Kuwana, cited in [66]) but now occurs almost worldwide. This whitefly is largely
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PlERO C. CACIAGLI
Table V Whitefly Vectors of Unidentified Viruses and Disease Agents in Europe Whitefly Bemisia tabaci
Parabemisia myricae
Trialeurodes vaporariorum
Country
Reference(s)
Austria Azerbaijan Cyprus Denmark France Georgia Greece Greece (Crete) Hungary Italy Norway Poland Portugal Spain
[72] [70] [70] [70] [70] [70] [70] [74] [71] [70] [72] [72] [70] [42] [70] [73]
Turkey Cyprus Greece and Crete Italy Portugal Spain Turkey All over Europe, in general In particular: Belgium Bulgaria France Greece and Crete Hungary Italy The Netherlands Norway Scotland Spain United Kingdom Turkey
[70] [73] [68] [68] [68] [69] [68] [68]
Virus or disease agent
TYLCV-Is«
TYLCV-Sar«
TYLCV-Sar« CYSDV,^TYCV-Sar,« TYCV-Is« CYSDV,^ TYLCV-Is«
Citrus chlorotic dwarf*^
BPYV* BPYV* BPYV* BPYV* Maple mosaic^ BPYy^TICV* BPYV^ [65] [64] BPYV* BPYV^ BPYV*
" See Table III for references. * See Table I for references. *^ See Table IV for references.
polyphagous on fruit trees, on which it can complete five generations per year in southern Italy [66] and up to eight on the southern shores of the Mediterranean [67]. Adult P. myricae generally appear more similar to the greenhouse whitefly than to Bemisia tabaci, but with brown areas on the thorax. Puparia have a pronounced waxy
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WHITEFLY-BORNE VIRUSES
287
band all around. In Europe, B myricae was first reported in Cyprus and Turkey, then in Greece and Crete, Italy, Spain [68], and Portugal [69]. It is a menace to southern Europe, considering that citrus is an important crop in all these countries. C. Bemisia tabaci Bemisia tabaci was first described in Greece in 1889 by Gennadius, who separated this species from T. vaporariorum. It is present, at least in some areas, in Cyprus, France, Greece, Italy, Portugal, Spain, Turkey, Azerbaijan, and Georgia in the open [70]. It also has been reported from Hungary [71], Austria, Germany, Denmark, Norway, and Poland [72] in greenhouses, mainly in connection with ornamentals, particularly poinsettia. As for which biotypes are present, the B biotype has been detected in Austria, Cyprus, Denmark, France, Germany, Italy, the Netherlands, Norway, Poland, Portugal, and Spain [72], and one or more non-B biotypes have been reported in Spain and Turkey [73] and Crete [74], and Q and S biotypes in Spain [42]. The A biotype, as defined by Costa and Brown [75] has not yet been detected in Europe. Differences in vectoring efficiencies among B and non-B silvering and nonsilvering biotypes, if any, are not as dramatic as between A and B biotypes [76], as so far tested [10, 35, 77]. D.
Maps
Distribution maps of some diseases and vectors mentioned in this chapter are available on the web site of the Istituto di Fitovirologia Applicata: http://www.ifa.to.cnr.it/.
IV. Concluding Remarks The problems presented by whitefly-borne viruses in Europe, although not as bad as in other parts of the world [74], do cause concern, even prompting attempts to eradicate one of the vectors, B. tabaci, from countries where it can only become established on protected crops [78, 79]. Considering the increasing movement of living plant materials, the spreading of both B. tabaci and B myricae, and the appearance of B. tabaci biotypes more adapted to relatively low temperatures [80], one can easily forecast that the problems due to these pests and vectors will progress from bad to worse, especially in view of global warming. The latter will undoubtedly dramatically increase the areas fit for these typically subtropical insects [80].
References 1. Duffus, J.E. (1987). Whitefly transmission of plant viruses. In "Current Topics in Vector Research" (K.F. Harris, ed.), Vol. 4, pp. 73-91. Springer-Verlag, New York.
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2. Mayo, M.A. (1999). Developments in plant virus taxonomy since the publication of the 6th ICTV Report. Arch. Virol. 144, 1659-1666. 3. Murphy, F.A., Fauquet, CM., Bishop, D.H.L., Ghabrial, S.A., Jarvis, A.W., MartelH, G.P., Mayo, M.A., and Summers, M.D. (1995). Virus Taxonomy. Classification and Nomenclature of Viruses. Sixth Report of the International Committee on Taxonomy of Viruses. Springer-Verlag, Vienna. Arch. Virol., Suppl. 10. 4. Wisler, G.C., Duffiis, J.E., Liu, H.-Y., and Li, R.H. (1998). Ecology and epidemiology of whiteflytransmitted closteroviruses. Plant Dis. 82, 270-280. 5. Duffus, J.E. (1965). Beet pseudo-yellows virus, transmitted by the greenhouse whitefly, Trialeurodes vaporariorum. Phytopathology 55, 450^53. 6. Liu, H.-Y., and Duffus, J.E. (1990). Beet pseudo-yellows virus: Purification and serology. Phytopathology 80, 866-869. 7. Van Dorst, H.J.M., Huijberts, N, and Bos, L. (1980). A whitefly-transmitted disease of glasshouse vegetables, a novelty for Europe. Neth. J. Plant Pathol. 86, 311-313. 8. Lot, H., Onillon, J.-C, and Lecoq, H. (1980). Une nouvelle maladie a virus de la laitue en serre: La jaunisse transmise par la mouche blanche. P.H.M. Revue Horticole 209, 31-34. 9. Lot, H., DelecoUe, B., and Lecoq, H. (1982). A whitefly transmitted virus causing muskmelon yellows in France. Acta Hort. Ill, 175-182. 10. Woudt, L.P., de Rover, A.P, de Haan, P.T. and van Grinsven, M.Q.J.M. (1993). Sequence analysis of the RNA genome of cucumber chlorotic spot virus (CCSV), a whitefly transmitted closterovirus. In "Abstracts of the 9th International Congress of Virology, Glasgow, Scotland, p. 326. 11. Berdiales, B., Bemal, J.J., Saez, E., Woudt, B., Beitia, E, and Rodriguez-Cerezo, E. (1999). Occurrence of cucurbit yellow stunting disorder virus (CYSDV) and beet pseudo-yellows virus in cucurbit crops in Spain and transmission of CYSDV by two biotypes of Bemisia tabaci. Eur J. Plant Pathol. 105,211-215. 12. Cuartero, J., Esteva, J., and Nuez, F. (1985). Sintomatologia y desarrollo del amarilleamiento del melon en cultivo bajo invemadero. In "Proceedings of the IV Congreso Nacional de Fitopatologia, Pamplona (Spain)", as cited by Soriat et al. [15]. 13. Jorda-Gutierrez, C , Gomez-Guillamon, M.L., Juarez, M., and Alfaro-Garcia, A. (1993). Clostero-like particles associated with a yellows disease of melons in South-eastern Spain. Plant Pathol. 42, 722-727. 14. Nuez, F, Pico, B., Iglesias, J., Esteva, J., and Juarez., M. (1999). Genetics of melon yellows virus resistance derived from Cucumis melo spp. agrestis. Eur J. Plant Pathol. 105, 453^64. 15. Soria, C, Gomez-Guillamon, M.L., and Duffiis, J.E. (1991). Transmission of the agent causing a melon yellowing disease by the greenhouse whitefly Trialeurodes vaporariorum in southeast Spain. Neth. J. Plant Pathol. 97, 289-296. 16. Hristova, D.P., and Natskova, VT. (1986). Interrelation between Trialeurodes vaporariorum West, and the virus causing infectious chlorosis in cucumbers. Biol.-Virol. Compt. Rend. Acad. Bulgare Sci.3% 105-118. 17. Meunier, S., and Verhoyen, M. (1987). Identification de la cause de jaunissements observes sur les laitues en plein air et en serre en Belgique. Medelingen van der Faculteit Landbouwwetenschappen, Rijksuniversiteit Gent 52, 1033-1039. 18. Ragozzino, A., Alioto, D., and lengo, C. (1989). The yellowing virus and mycoplasma diseases of lettuce in Campania and Latium Regions. Riv. Patol. Veg. S. IV IS, 15-19. 19. Coffin, R.S., and Coutts, R.H.A. (1990). The occurrance of beet pseudo-yellows virus in England. Plant Pathol. :39,6n-6^5. 20. Coffin, R.S., and Coutts, R.H.A. (1993). Relationships among Trialeurodes vaporariorum-traiasmitted yellowing viruses from Europe and America. J. Phytopathol. 143, 375-378.
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21. Livieratos, I.C., Katis, N., and Courts, R.H.A. (1998). Differentiation between cucurbit yellow stunting disorder virus and beet pseudo-yellows virus by a reverse transcription-polymerase chain reaction assay. Plant Pathol 47, 362-369. 22. Rubio, L., Soong, J., Kao, J., and Falk, B.W. (1999). Geographic distribution and molecular variation of isolates of three whitefly-bome closteroviruses of cucurbits: Lettuce infectious yellows virus, cucurbit yellow stunting disorder virus, and beet pseudo-yellows virus. Phytopathology 89,707-711. 23. Wisler, G.C., Liu, H.-Y., Klaassen, VA., Duffiis, J.E., and Falk, B.W. (1996). Tomato infectious chlorosis virus has a bipartite genome and induces phloem-limited inclusions characteristic of the closteroviruses. Phytopathology 86, 622-626. 24. DuflFus, J.E., Liu, H.-Y., and Wisler, G.C. (1994). Tomato infectious chlorosis virus-a new clostero-like virus transmitted by Trialeurodes vaporariorum. Eur. J. Plant Pathol. 102, 219-226. 25. Dellavalle, G., Caciagh, P., Bosco, D., Lisa, V, d'Aquiho, M., Milne, R.G., and Masenga, V (1995). A whitefly-transmitted clostero-like virus isolated from diseased tomato. In "Proceedings of the 8th Conference on Virus Diseases of Vegetables, Prague, Czech Republic, July 9-15,1995, pp. 35-38. 26. Duflfus, J.E., Caciagh, R, Liu, H.-Y, Wisler, G.C, and Li R.H. (1996). Occurrence of tomato infectious chlorosis virus in Europe. In "Silverleaf Whitefly 1996 Supplement to the Five-Year National Research and Action Plan." USDA-ARS Publ 1996-01:36. 27. Hassan, A.A., and Duffus, J.E. (1991). A review of a yellowing and stunting disorder of cucurbits in the United Arab Emirates. Emir. J. Agr. Sci. 2, 1-16. 28. Celix, A., Lopez-Sese, A., Almarza, N., Gomez-Guillamon, M.L., and Rodriguez-Cerezo, E. (1996). Characterization of cucurbit yellow stunting disorder virus, a Bemisia ^a6ac/-transmitted closterovirus. Phytopathology 86, 1370-1376. 29. Duffus, J.E., Larsen, R.C., and Liu, H.-Y. (1986). Lettuce infectious yellows virus—a new type of whitefly-transmirted virus. Phytopathology 76, 97-100. 30. Tian, T, Klaassen, VA., Soong, J., Wisler, G., Dufais, J.E., and Falk, B.W (1996). Generation of cDNA specific to lettuce infectious yellows closterovirus and other whitefly-transmitted viruses by RT-PCR and degenerate oligonucleotide primers corresponding to the closterovirus gene encoding the heat shock protein 70 homolog. Phytopathology 86, 1167-1173. 31. Wisler, G.C, Li, R.H., Liu, H.-Y, Lowry, D.S., and Duffus, J.E. (1998). Tomato chlorosis virus: A new whitefly-transmitted, phloem-limited, bipartite closterovirus of tomato. Phytopathology 88, 402^09. 32. Abouzid, A., and Jeske, H. (1986). The purification and characterization of gemini particles from Abutilon mosaic virus infected Malvaceae. J. Phytopathol. 115, 344-353. 33. Frischmut, T, Zimmat, G., and Jeske, H. (1990). The nucleotide sequence of abutilon mosaic virus reveals prokaryotic as well as eukaryotic features. Virology 178,461-468. 34. Macintosh, S., Robinson, D.J., and Harrison, B.D. (1992). Detection of three whitefly-transmitted geminiviruses occurring in Europe by tests with heterologous monoclonal antibodies. Ann. Appl. Biol. 121,279-303. 35. Bedford, I.D., Briddon, R.W., Brown, J.K., Rosell, R.C, and Markham, RG. (1994). Geminivirus transmission and biological characterisation oiBemisia tabaci (Gennadius) biotypes from different geographic regions. Ann. Appl. Biol. 125, 311-325. 36. Harrison, B.D, (1994). Detection, identification and assessment of variation of whitefly-transmitted geminiviruses. Arab J. Plant Prot. 12, 120-116. 37. Osaki, T, and Inouye, T. (1981). Tobacco leaf curl virus. CMI/AAB Descriptions of Plant Viruses No. 232. 38. Osaki, T, Kobatake, H., and Inouye, T. (1979). Yellow vein mosaic of honeysuckle (Lonicera japonica Thunb), a disease caused by tobacco leaf curl virus in Japan. Ann. Phytopathol. Soc. Jpn. 45, 62-69.
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CHAPTER 15
Transmission Properties of Whitefly-Borne Criniviruses and Their Impact on Virus Epidemiology GAIL C. WISLER JAMES E. DUFFUS
/. Introduction The genus Crinivirus is in the family Closteroviridae [1]. The type member of this new genus is Lettuce infectious yellows virus (LIYV). The genus was separated from the genus Closterovirus because its members have a bipartite genome as opposed to the monopartite genome of closteroviruses [2,3]. As is consistent with a divided genome, criniviruses have particle lengths that are roughly half the lengths reported for closteroviruses [4]. All viruses that have been assigned to the genus Crinivirus are transmitted by whiteflies, as opposed to aphids for members of the genus Closterovirus. Those viruses that have not been characterized on a molecular level to fully document their genome organization but are whitefly-transmitted, for example, Lettuce chlorosis virus (LCV) dindi Beet pseudo-yellows virus (BPYV), are currently assigned to the genus Crinivirus. The incidence and impact of viruses belonging to the genus Crinivirus are dependent on interactions among several factors, including the virus itself, the host range of that virus, the specific whitefly vector(s), surrounding crops and weeds, and climatic conditions. Unlike viruses belonging to the genus Begomovirus in the family Geminiviridae, which are transmitted by Bemisia tabaci (Gennadius) biotypes A and B (also known as the silverleaf whitefly, Bemisia argentifolii) [5, 6], criniviruses are unique in that they are transmitted by one or more of the following whitefly species: B. tabaci biotype A (sweet potato whitefly, SPWF), B. tabaci biotype B (silverleaf whitefly, SLWF), Trialeurodes vaporariorum Westwood (greenhouse whitefly, GHWF), and Trialeurodes abutilonea Haldeman (banded wing whitefly. Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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BWWF). A unique characteristic of criniviruses is that some members can be transmitted by more than one whitefly vector species or biotype with different levels of efficiency. Another important difference in transmission between these two genera is that begomoviruses are transmitted persistently while criniviruses are transmitted semipersistently. In the persistent relationship, virus is acquired by insects feeding on an infected plant, usually from the phloem, and passes through the gut wall to the hemolymph and salivary glands, and from there to the insect's saliva [7, 8]. The virus is retained in the insect for a relatively long time, from weeks to the lifetime of the insect, and it retains transmissibility through a molt. In the semipersistent relationship, there is a positive correlation between the length of the acquisition feeding period and the probability of virus acquisition. There does not appear to be an accumulation of inoculativity after the acquisition period has ended. Those criniviruses that have been studied have retention periods from 1 to 9 days (Table I). Preliminary evidence suggests that the Tomato infectious chlorosis crinivirus (TICV) is limited to the cibarium of the GHWF (Wisler, unpublished data) and does not appear to circulate in the insect, which is consistent with semipersistent transmission. This chapter addresses criniviruses and their whitefly vectors, particularly the effect of transmission characteristics on their disease epidemiology. Little research has been reported on whitefly transmission characteristics of criniviruses; however, knowledge of the vectors is crucial to management of the diseases caused by these viruses. Although molecular probes and antisera have been developed for several of the criniviruses, the virus titer in agronomic hosts, in particular cucurbits, tomatoes, and lettuce, is characteristically low. Transmission experiments are, therefore, often more sensitive than laboratory assays and provide important backups to the latter.
//. A.
Vector Transmission and Virus-Vector Relationstiips Transmission Efficiency
Tests for efficiency of transmission involve allowing aviruliferous whiteflies to feed on virus-infected plants singly or in groups of 5,10,20, or 40 insects per leaf cage for a 24-hour acquisition access feeding period (AAFP) and then transferring them to appropriate inoculation indicator plants to test for virus transmission. Except for cucurbit yellow stunting disorder virus (CYSDV), which is restricted to members of the Cucurbitaceae [9], Physalis wrightii Gray is an excellent inoculation (transmission) test plant for most criniviruses evaluated in our laboratory. Ten replications are routinely used for each test group, and inoculation test plants must be young, generally at the three- to four-leaf stage. Virus source plants must also be in good condition. Ideally, virus source plants should be just starting to
Table Z Transmission Properties of Whitefly-Borne Criniviruses
Vector transmission characteristics of criniviruses
Specific vector(s)b,C
Virusa
GHWF *SLWF~SPWF *SPWF>SLWF SLWF=**SPWF
BPYV CYSDV LIYV LCV
2 wl
SPCSV
GHWF SLWF>BWWF> *SPWF>GHWF SLWF
AYV
BWWF
TICV ToCV
~
Transmission efficiency 1-40 insects
Acquisition threshold 1 4 3 hr
Inoculation threshold 1 4 8 hr
Persistence in vector
10-83.3% 3-85% 22.2-97.4% *o-57.5% * *2 .9-74.4% 8-83% 18-80%
4.2-70.8% 3-85% 7.5-96.7% +96%
8.3-95.7% 8-97% 6 hr minimum 3476%
6 days 7-9 days 3 days 4 days
wide, tomato (-) Cucurbits only wide, cucurbits (+) wide, cucurbits (-)
6-94% ndd
1680% 672%
3 days <1 day
wide, lettuce (+) wide, lettuce (-)
9.3-79.1%
8.3-66.7%
4.2-87.5%
2 days
4.1-80.8% (50 insects)
18.8-76.6%
42.2-100%
3 days
Impomeu spp., It clevelundii N. edwurdsonii moderate, Anodu ubutiloides
Diagnostic host range
~
AYV (abutilon yellows virus), BPYV (beet pseudo-yellows virus), CYSDV (cucurbit yellow stunting disorder virus), LCV (lettuce chorosis virus, L I W (lettuce infectious yellows virus), SPCSV (sweet potato chlorotic stunt virus), TICV (tomato infectious chlorosis virus), and ToCV (tomato chlorosis virus). BWWF (banded wing whitefly, Trialeurodes ubutiloneu), GHWF (greenhouse whitefly, T vuporuriorum), SLWF (silverleaf whitefly, Bemisiu argentifolii = B. tubaci biotype “B’), and SPWF (sweet potato whitefly, B. tubuci biotype “A”). Transmission efficiency (third column) is for the specific vector indicated (* or **) in the second column. Data not determined.
(I
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show symptoms, usually 2 to 4 weeks post-inoculation depending on the time of year and quality of light. Infected P. wrightii plants show characteristic symptoms of interveinal yellowing and bronzing, and leaves become stiff and brittle within 2 to 4 weeks post-inoculation. B.
Minimum Acquisition Access Feeding Period
The minimum AAFP necessary for aviruliferous whiteflies to acquire a virus (acquisition threshold) is studied over AAFPs ranging from 1 to 48 hours. Actual AAFPs tested vary among experimenters. After feeding aviruliferous whiteflies on virus-infected plants in groups of 20 per leaf cage for each AAFP chosen, the insects are removed and transferred to healthy test seedlings. In general, the transmission efficiency of whiteflies increases up to a saturation point, with corresponding increases in the AAFP C.
Inoculation Feeding Period
Minimum inoculation feeding period (IFP) (inoculation threshold) needed by viruliferous whiteflies to inoculate healthy test seedlings with virus is determined by allowing whiteflies an appropriate AAFP, usually from 24 to 48 hours, before transferring them to indicator plants in large enough groups, usually 20 to 40 insects per leaf cage, to ensure some transmission for incremental IFPs of 1 to 48 hours. D.
Persistence
The length of time a viruliferous whitefly retains its ability to transmit virus is referred to as persistence. After a 24- to 48-hour AAFP on virus source plants, viruliferous whiteflies are collected in leaf cages, usually 20 to 40 per cage, which are used to make daily serial transfers to healthy F wrightii seedlings for up to 10 days. All of the above experiments (section IIA through D) are replicated at least three times, five replications being standard in our laboratory.
///.
Criniviruses Infecting Cucurbits
A. Beet Pseudo- Yellows Virus Beet pseudo-yellows virus (BPYV) was the first whitefly-transmitted member of the family Closteroviridae described [10]. Full details about the genome organization of BPYV are still lacking, but molecular probes are available for diagnosis of BPYV based on the hallmark heat shock protein (HSP70), which is present in all closteroviruses studied to date [11, 12]. Owing to the extremely low titer of BPYV in infected plants, virion purification is difficult.
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Beet pseudo-yellows virus was also the first virus reported to be transmitted by the GHWF. It has a wide host range, including agronomic crops, in particular cucurbits, ornamentals, and weeds. Since its discovery in California greenhouses, it has been reported worldwide in sugarbeet, spinach, cucumber, melons, flax, lettuce, carrot, dandelion, Gomphrena sp., Callistephus sp., Aguilegia sp., Tagetes sp. (marigold), zinnia, and Godetia sp. Hot-house cucumbers and melons are frequently found to be infected with BPYV The greenhouse vegetable industry has expanded in North America, Europe, and the Middle East. Greenhouse-grown cucumbers account for about 10% of the greenhouse vegetable production in the United States [13]. The greenhouse vegetable and ornamental industry has expanded the natural range of the GHWF to areas where it would not normally survive the winter. Symptoms of BPYV infection, regardless of the host, are typical of many criniviruses described, with interveinal yellowing and necrosis, and stiffness or brittleness of the lower leaves. Symptoms are typically produced on the middle to lower parts of the plant; young growth appears normal. Symptoms resemble early senescence and nutritional deficiencies in plants. This is due to the plugging of the phloem with large viral inclusion bodies, which are likely to restrict movement of nutrients in the plant. For cucurbits in particular, chlorotic flecks are first produced in intermediate and older leaves. These enlarge and coalesce to produce large areas of yellowing. Necrosis forms in these yellowing areas, and leaves die prematurely. The latter effect leads to reduced yield and vigor of infected plants. Because symptoms of criniviruses can mimic those of nutritional disorders and natural senescence, they are difficult to recognize as viral and losses caused by BPYV infections are not easily or fully documented. The GHWF is a relatively efficient vector of BPYV Virus can be acquired and inoculated by whiteflies during AAFPs as brief as 1 hour. Viruliferous whiteflies given a 6-hour IFP transmitted BPYV at a 70% level of efficiency. The virus persists in the vector for 6 days during daily serial transfers. Single viruliferous whiteflies can transmit BPYV at 10% efficiency, whereas groups of 40 can do so at 83% efficiency [10]. Although it is difficult to assess losses due to BPYV, it is presumed to be fairly widespread in the greenhouse cucurbit and ornamental industries. Reduction of whitefly populations, through the use of either insecticides or natural predators, is only somewhat effective in limiting the spread of BPYV in greenhouse and field culture. B.
Cucurbit Yellow Stunting Disorder Virus
Symptoms of CYSDV are very similar to those produced by BPYV However, CYSDV is easily distinguished from BPYV on the bases of whitefly vectors, host ranges, and specific molecular probes [12]. Cucurbit yellow stunting disorder virus is transmitted more efficiently by the SLWF than by the SPWF, and the host range is limited to the Cucurbitaceae [9]. Until recently, CYSDV was restricted to
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the Middle East and the Mediterranean. The most efficient vector of CYSDV, B. tabaci biotype B, originated in the Middle East, which corresponds to the original range of CYSDV Between 1970 and 1990 the populations of the SLWF increased throughout the world in tropical and subtropical areas [14, 15] as a result of international export of plants infested with the SLWF. Even with large acreages planted to cucurbits throughout the sunbelt states of the United States, it was not expected that CYSDV could move from the Middle East and Mediterranean regions to the United States because cucurbit germplasm is generally not moved internationally. Recent reports indicate that CYSDV has now been found in North America, in greenhouse-grown melon in Texas and Mexico [16]. This was confirmed by nucleic acid hybridization and reverse transcriptase-polymerase chain reaction (RT-PCR) analyses. Bemisia tabaci is more likely to be a pest of fieldgrown than of greenhouse-grown cucurbits, and this corresponds to the higher incidence of CYSDV in field production. The SLWF is a relatively inefficient vector of CYSDV compared with the GHWF's efficiency in transmitting BPYV Single whiteflies are able to transmit CYSDV only 3% of the time, and groups of 60 are needed to reach 85% transmission efficiency. Relatively long AAFPs are needed to achieve transmission (18 hours for 80% transmission efficiency), and IFPs are long as well (24 hours for 83% transmission efficiency). In daily serial transfers, CYSDV persisted in the SLWF for at least 7 days when a Spanish isolate was tested and for 9 days in the case of an Israeli isolate (Wisler, unpublished data), the longest persistence time of all criniviruses tested to date [9]. The A biotype of ^. tabaci also transmits CYSDy but with a much lower efficiency than the B biotype. Although the SLWF is not a highly efficient vector for CYSDV as compared with other criniviruses, the long persistence of the virus in the vector, the preference for cucurbits by the SLWF, and the high populations of the vector are responsible for significant economic losses due to this virus. The heavy use of the insecticide imidacloprid has helped to maintain the whitefly populations at low levels on treated crops, although resistance to the insecticide has been reported in some areas [17, 18]. In addition, progress is being made toward tolerance to CYSDV in melons [19].
IV. Criniviruses Infecting Lettuce A. Lettuce Infectious Yellows Virus The development of Lettuce infectious yellows virus (LIYV) in the southern United States to epidemic proportions and subsequently its apparent disappearance in current cropping systems is an excellent example of the impact that insect populations and transmission characteristics have on virus ecology. Although LIYV was first described in lettuce, it caused significant losses in sugarbeets,
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cucurbits, and other crops in the southwestern United States between 1980 and the early 1990s [20]. This virus has a wide host range and infects 45 species in 15 plant families. The most efficient vector of LIYV is the SPWF, or B. tabaci biotype A, which was the predominant biotype in the southern United States until the late 1980s. At that time, a new biotype emerged throughout the southern United States, which caused silvering on leaves of cucurbit plants. Because of the symptoms induced on cucurbits, it was called the silverleaf whitefly (SLWF). Although it has been designated as a separate species, B. argentifolii [5,6], the designation as B. tabaci biotype B is still the more accepted terminology worldwide [14, 21]. The SLWF displaced the SPWF quickly and by the mid-1990s increased its populations to levels 1,600 times those of previous years [22]. Prior to the introduction of the SLWF, the epidemics of LIYV in lettuce were strongly influenced by fall-planted melons. Since cucurbits are also a host of LIYV, they provided a link from one lettuce crop to another. As a new lettuce crop was planted, viruliferous whiteflies from the melons moved to lettuce and caused virtually 100% infection of the new crop. The SLWF increased to such high numbers and had such a preference for cucurbits that it essentially destroyed fallplanted melons and eliminated the crop. Thus, in the absence of a fall melon crop, LIYV was no longer transferred to the young lettuce, and lettuce crops were essentially virus-free (Fig. 1). Another factor that influenced this is that the B biotype ofB. tabaci is a very inefficient vector of LIYV, being less efficient than biotype A by a factor of 100. In spite of the return of fall melons in the Imperial Valley of California, LIYV has not reemerged. In fact, recent surveys indicate that LIYV no longer exists in the desert southwest United States, which was the original range of the virus [23]. Lettuce infectious yellow virus has never been reported outside the southwestern United States. In addition, the efficient vector of LIYV, the A biotype of 5. tabaci, appears to have been completely displaced by the SLWF and is no longer found in these areas. It is not known if the A biotype exists in nature today. However, it is still a valuable research tool, and colonies are maintained in our laboratory for transmission experiments. The A biotype retains LIYV for 3 days or less. Single viruliferous whiteflies can transmit LIYV at 22% efficiency and groups of 40 whiteflies at almost 100% efficiency. Although the A biotype can acquire LIYV after a 10-minute AAFP, greater efficiency (68 to 87%) is achieved with longer AAFPs ranging from 1 to 48 hours [20]. The time required for viruliferous whiteflies to inoculate LIYV to plants is less than 6 hours. Complete transmission experiments for the B biotype have not been performed, but the efficiency of transmission of LIYV is extremely low compared with the A biotype [24, 25]. Recent studies by Tian et al [26] have shown that partially purified preparations of LIYV can be used for in vitro acquisition by B. tabaci. This is the first evidence that the determinants for whitefly transmission of criniviruses may be located on the purified virions. Resistance or tolerance to LIYV was developed in lettuce, sugarbeet, and cucurbit varieties in response to disease epidemics. Preliminary studies are under-
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No melons planted Fig. 1 Lettuce infectious yellows virus (LIYV) infection and Bemisia populations in lettuce in the Imperial Valley, California from 1990 to 1994. Shown are the virus incidence (visual counts) and whitefly populations made in 2-minute collections with a hand-held collection device from 20 randomly sampled mature lettuce fields at each date. Because of the destructive effect of the B biotype of Bemisia, virtually no melons were planted in the fall growing seasons from 1991 through 1993. The percentages of LIYV infected plants are indicated by black bars and Bemisia populations by lines. Thus, in spite of record whitefly populations during the years since the B biotype became predominant, the absence of the major source of LIYV (fall melons) resulted in a very low incidence of LIYV
way to determine if the resistance to LIYV in lettuce and sugarbeet confers resistance to Lettuce chlorosis virus (LCV) as well. There has been some speculation that perhaps LCV was present during the LIYV epidemics and that LIYV resistance may be useful for controlling more than one crinivirus infecting these crops. For example, the U.S. Department of Agriculture, Agricultural Research Service sugarbeet breeding program for the "virus yellows" complex of luteoviruses and closteroviruses was ongoing in the Imperial Valley when LIYV was prevalent. Because selection was based on absence of yellowing, varieties resistant to LIYV were developed before the causal agent was even characterized [27, 28]. B. Lettuce Chlorosis Virus A second, potentially destructive whitefly-transmitted crinivirus, LCV, was found in the Imperial Valley of California after LIYV was no longer present. Lettuce chlorosis virus has several characteristics in common with LIYV, including the typical symptomatology of interveinal yellowing and stiffness of lower leaves and
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leaf necrosis. In contrast to LIYy both A and B biotypes of B. tabaci are relatively efficient vectors. Lettuce chlorosis virus is similar to LIYV with respect to its host range, except that LCV does not infect members of the Cucurbitaceae. The whitefly population is currently being controlled in lettuce by use of imidacloprid; thus LCV has not been an economically significant problem, although it is consistently identified in symptomatic lettuce and sugarbeet when yellowing symptoms are found in the southwestern United States. Studies by McLain et al [29] indicate that LCV can cause significant yield losses in lettuce under field inoculum pressure, compared with plots treated with insecticide. These diagnoses are made by dot blot hybridization with riboprobes, western blot analyses using antisera to the purified virion, reverse transcriptase-polymerase chain reaction, and transmission to indicator plants followed by retesting from these source plants. The low titer of LCV in lettuce and sugarbeet can preclude positive results directly from infected plants. In those cases, a negative result in a serological or molecular assay can be misleading, and transmission to and retesting from hightitered indicator plants such as P. wrightii, Nicotiana clevelandii, or Nicotiana benthamiana can confirm questionable results. Immunospecific electron microscopy (ISEM) using a gold-labeled goat-antirabbit conjugated serum has also been useful in identifying LCV in both single and mixed infections with LIYV (Wisler, unpublished data). Transmission efficiency experiments using B. tabaci biotype A indicate that single whiteflies allowed 24-hour AAFPs on LCV-infected plants can subsequently transmit the virus to healthy test plants. Transmission rates established for whiteflies tested singly or in groups of up to 40 per leaf cage ranged from 2.9% (singly) to 74.3% (groups of 40). The B biotype was not as efficient, ranging from 0 to 57.5% transmission rates in tests using five replications of 10 plants each with 30 insects per leaf cage [30]. The acquisition threshold is 1 hour, but LCV is transmitted with greater efficiency (4 to 96%) by the A biotype after 3- to 48-hour AAFPs. Lettuce chlorosis virus remains inoculative in viruliferous whiteflies for 4 days, with transmission being fairly efficient for the first 3 days. No transmission was observed after 5 or more days post-acquisition.
V. Criniviruses Infecting Tomatoes A.
Tomato Infectious Chlorosis Virus
A new whitefly-transmitted virus infecting tomato was detected in 1993 in Orange County, California [31]. In one season the growers in this county incurred $2 million in losses due to Tomato infectious chlorosis virus (TICV). The high virus incidence coincided with extremely high populations of the GHWF, and TICV was established in surrounding weeds, including bristly oxtongue (Picris
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echioides L.), tree tobacco (Nicotiana glauca Graham), and wild artichoke {Cynara cardunculus L.). This new virus, TICV, is solely transmitted by the GHWF, and is the first crinivirus since BPYV reported to be transmitted by this vector. Since tomato is not a host of BPYV, this is clearly a new and distinct virus. Subsequent studies have shown that TICV has a bipartite genome, making it the second crinivirus to be characterized [32]. Tomato production in the United States was valued at about $1.64 billion in 1998, with a fresh-market production at $1.1 billion and production for processing at $540 million. More than half of the fresh-market tomato production is concentrated in Florida and California. Of 800 acres of greenhouse vegetables in the United States, 700 acres are planted in tomatoes [13]. Tomatoes are the most popular vegetable in the Israeli market, where they are grown in greenhouses yearround. In Europe, greenhouse tomato production is common. Although the GHWF is found in field conditions, it is well known as a common greenhouse pest. Since its initial detection, TICV has been found infecting tomatoes and other vegetables, ornamental crops, and weeds in field and greenhouse situations. Serological and nucleic acid probes have been developed that work well for diagnosis of TICV in tomato and other hosts [12, 33]. Tomato infectious chlorosis virus has been found by hybridization or serological assays, or both, in California, North Carolina, Italy, and Taiwan. The international movement of the GHWF and the associated movement of viruses in ornamentals and tomato breeding material is likely responsible for these widespread findings. The GHWF is a relatively efficient vector of TICV Single GHWFs allowed a 24-hour AAFP on TICV-infected plants can transmit TICV As expected, transmission rates increase with longer AAFPs. Transmission rates for GHWFs tested singly or in groups of up to 40 per leaf cage range from 8% (singly) to 83% (groups of 40). The TICV is acquired by the GHWF during 1-hour AAFPs, but is transmitted more efficiently (6 to 94%) after AAFPs ranging from 3 to 48 hours. IFPs of 1 to 48 hours result in plant inoculation rates of 16 to 80% [31]. The persistence of TICV in viruliferous GHWFs, as determined by daily serial transfers, is 3 to 4 days, although most of them ceased to be inoculative after 1 day. The virus has a wide host range, which includes many epidemiologically significant plants, from weeds to ornamentals and bedding plants to agronomic hosts, including lettuce and potato. Although all potato varieties tested were susceptible, no natural infection has yet been found in this host. B.
Tomato Chlorosis Virus
As a result of interest generated in whitefly-transmitted yellowing viruses of tomato, a second distinct crinivirus called Tomato chlorosis virus (ToCV) was recognized and later described from Florida [34]. Symptoms of ToCV in tomato are very similar to those of TICV, with characteristic interveinal yel-
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lowing, necrotic flecks, and brittleness of leaves, accompanied by yield reduction. This virus is also bipartite, with a genome size and organization similar to those of TICV. Tomato chlorosis virus is unique among known whitefly-transmitted viruses because it is transmitted by four whitefly vectors: the greenhouse (GHWF), banded wing (BWWF), sweet potato (SPWF), and silverleaf (SLWF) whiteflies. It was first detected in Florida, where the SLWF is prevalent. Since its initial detection, it has been identified in Louisiana, Taiwan, Spain, and South Africa (Wisler, unpublished data). With the availability of four whitefly vectors, the geographic distribution of ToCV should increase relative to that of TICV. Comparative transmission studies have shown wide differences in efficiency among the four vectors. The SLWF has the highest efficiency, ranging increasingly from 12.5 to 98% for 1 up to 40 whiteflies per leaf cage, respectively. This is expected because of the occurrence of ToCV in regions highly populated by the SLWF. Surprisingly, the BWWF is almost the equal of the SLWF in transmission efficiency. Tested singly, BWWFs had an efficiency of 7.5%, whereas groups of 40 per cage were 100% efficient. The SPWF had increasing transmission efficiencies, ranging from 0 to 68% for insects tested singly or in groups of up to 40 per leaf cage. The GHWF had the lowest efficiency, ranging from 0 to 28% for test insects numbering from 1 up to 40 per leaf cage. The higher efficiencies of the SLWF and BWWF would suggest that ToCV may be more prevalent in fieldgrown tomatoes than in those grown under greenhouse conditions, as is common for TICV When the viruliferous SPWFs were tested for persistence by daily serial transfers, they lost their ability to transmit the virus during the first 24-hour feeding period. Like TICV, ToCV also has a fairly wide host range (24 plant species in seven families), which includes many ornamental, agronomic, and weed hosts, including Beta sp. and potato. In contrast to TICV, ToCV does not infect lettuce. Thus, in addition to specific probes designed to distinguish between these two viruses, the use of different whitefly vectors and selective hosts provides additional diagnostic information, particularly since the titers of these viruses are characteristically low and laboratory analyses directly from tomato can yield negative results. Because many greenhouse tomato production facilities rely on bumblebees for pollination, insecticides cannot be used for whitefly control. Natural predators are used, but biocontrol is not highly effective, and low-level vector populations are enough to maintain moderate levels of infection. Since the tomato crops are grown for long periods of time, up to 8 or 10 months, infections can become chronic. A common practice in greenhouse tomatoes is to interplant with small tomatoes after virus-infected, nonproductive plants are removed. Without completely removing all plant material and starting a new crop, TICV or ToCV or both can continue to cause yield losses in these hydroponic operations. Since each plant can produce up to 30 pounds of fruit, yield reductions caused by these viruses are of concern to the growers.
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W. Criniviruses Infecting Sweet Potato A. Sweet Potato Chlorotic Stunt Virus Together with coinfection by sweet potato feathery mottle potyvirus (SPFMV), Sweet potato chlorotic stunt virus (SPCSV) produces a serious disease of sweet potato, which was first reported in Nigeria [35]. Sweet potato, Ipomoea batatas (L.) Lam., germ plasm is vegetatively propagated and shipped throughout the world as an important food source for many countries. The whitefly component of the sweet potato virus disease complex was characterized according to the transmission test setups of Cohen et al [36] and Larsen et al [37]. Bemisia tabaci was used in both transmission experiments, and although it is not stated which biotype was used, the time and source of whiteflies indicate that it was the B biotype. The transmission efficiencies reported by Cohen et al [36] ranged from 12.5% to 93.8% for test insect groups ranging from 10 to 50 whiteflies per leaf cage. Acquisition (transmission) rates increased from 20.8% to as high as 87.5% for AAFPs ranging from 1 to 24 hours. Inoculation rates increased from 15.6% to 84.3% following IFPs ranging from 1 to 24 hours. Again, these results indicate a semipersistent mode of transmission. Transmission of SPCSV can occur by grafting but not by mechanical transmission. The host range of SPCSV is limited primarily to Ipomoea spp., A'^ clevelandii Gray, A^. benthamiana Domin, and Amaranthus palmeri L. Wats. Although SPCSV is whitefly-transmitted, the most important means of distribution of virus-infected propagative material is likely to be by vegetative propagation. All closterovirus-like isolates of SPCSV have bipartite genomes with indistinguishable mobilities in gel analysis. Although the clustering of geographically diverse isolates is different depending on serological or molecular results, all bipartite closterovirus isolates studied to date are closely related (S. Winter and H.J. Vetten, personal communication). Management of sweet potato virus disease is best accomplished through large-scale production of certified, virus-tested, micropropagative stock. For example, the Micropropagation Unit at North Carolina State University in Raleigh has an ongoing virus-indexing program for sweet potato which includes evaluation for true-to-variety and routine pathogen testing. These explants are distributed to other states, countries, and breeders for further increase and distribution for commercial use and research [38].
VII. Criniviruses Infecting Weed Hosts A. Abutilon Yellows Virus Abutilon yellows virus (AYV) was first found in collections of the BWWF on the weed Abutilon theophrasti Medic, collected in Illinois. This virus has been characterized with a bipartite genome, as is similar to other described criniviruses
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[39]. Abutilon yellows virus is unique in that it is one of only two crinivirus known to be transmitted only by the (BWWF). The significance of AYV in regard to agronomic crops and ornamentals, if any, is not known. Molecular probes are available for diagnosing AYV and distinguishing it from other described criniviruses. It shows symptoms in infected Anoda abutiliodes A. Gray that are distinct from those of most criniviruses. Instead of characteristic interveinal yellowing, AYV produces distinct vein yellowing on leaves of infected plants 2 to 3 weeks postinoculation. The host range of AYV appears limited, and no agronomically important hosts have yet been identified. Transmission studies indicate that the BWWF transmits AYV with efficiencies ranging from 4.1 to 80.8% for whiteflies tested singly or in groups of up to 50 per leaf cage. The acquisition efficiency ranges from 18.8 to 76.6% for 1 to 40 insects per leaf cage, and the inoculation efficiency ranges from 42 to 100%). The AYV persists in the BWWF for 3 days. B. Diodia Vein Chlorosis Virus Diodia vein chlorosis virus (DVCV) was first described infecting Virginia buttonweed, Diodia virginiana L. which occurs commonly throughout the southeastern United States. This virus is similar to AYV with respect to its vector, the BWWF, and the vein clearing symptoms that it produces on infected plants rather than the typical interveinal chlorosis. This virus has not been fully characterized on either a biological or molecular basis, but cytological studies show cytoplasmic vesiculation and highly flexuous rods typical of the Family Closteroviridae. Diodia vein chlorosis virus also has a restricted host range, which at present includes only D. virginiana [40].
VIIL Concluding Remarks Since the advent of molecular biology, emphasis on biological characterization of plant viruses has diminished with respect to research priorities. However, where disease management and control are concerned, knowledge of the biological characteristics of plant viruses is crucial. These characteristics include host range, cropping patterns, weed associations, climatic factors, and virus-vector-plant interactions. The nonpersistently transmitted potyviruses and persistently transmitted geminiviruses have received a great deal of attention at the molecular level, especially as that approach relates to virus disease control. The semipersistent, whitefly-transmitted criniviruses belong to a relatively new genus of plant viruses, and less is known, as compared with poty- and geminiviruses, of their biology, specific transmission determinants, and epidemiology. However, descriptive data on viruses were critical in the elimination of LIYV, for example, in the Imperial Valley of California between the 1980s and early 1990s.
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Molecular probes and antisera developed for several criniviruses serve as useful diagnostic tools. Degenerate primers that are specific for both aphid- and whitefly-transmitted closteroviruses and criniviruses, respectively, have been developed to detect most known viruses in these two viral groups using PCR technology [12, 41]. Nonetheless, some probes considered to be universal have failed to detect all members of target groups. For example, ToCV does not amplify with degenerate primers that target a conserved region of the hallmark heat shock protein (HSP70) gene of whitefly-transmitted criniviruses because of differences in nucleotide sequences. Likewise, the "universal" potyvirus monoclonal antibody does not react with Tulip break virus, Lily mottle virus [42], or Peanut mottle virus [43]. Therefore, negative results in nucleic acid-based assays or serological tests should be confirmed by transmission studies. Our experience with criniviruses indicates typically low titers in infected crop plants. The insect vector can be the most sensitive means of detecting criniviruses, particularly previously undescribed criniviruses. Since awareness of criniviruses is growing in both agricultural and academic communities, presumably numerous criniviruses await discovery and description. For example, a possible new crinivirus infecting tomato in the Canary Islands [22] is similar to ToCV, but it infects lettuce and ToCV does not. Accurate diagnoses without molecular or serological tools can still be made for several criniviruses by using specific hosts and vectors for transmission studies (Table I). Still, it is best to combine results from biological and laboratory assays to obtain and confirm the maximum amount of information regarding these viruses. This dual approach will best identify their diversity and the intra- and interrelationships that exist among criniviruses and between the genus Geminivirus and other viral taxa, respectively.
References Martelli, G.P. et al. (2000). Family Closteroviridae. In "Virus Taxonomy: Seventh Report of the International Committee on Taxonomy of Viruses" (M.H.V Van Regenmortel, CM. Fauquet, D.H.L. Bishop, E.B. Carsten, M.K. Estes, S.M. Lemon, J. Maniloff, M.A. Mayo, D.J. McGeoch, C.R. Pringle, and R.B. Wickner, eds.), pp. 943-952, Academic Press, San Diego. Klaassen, VA., Boeshore, M., Dolja, VV, and Falk, B.W. (1995). Partial characterization of the lettuce infectious yellows virus genomic RNAs, identification of the coat protein gene and comparison of its amino acid sequence with those of other filamentous RNA plant viruses. J. Gen. Virol. 75, 1525-1533. Klaassen, VA., Boeshore, M.L., Koonin, E.V, Tian, T, and Falk, B.W. (1996). Genome structure and phylogenetic analysis of lettuce infectious yellows virus, a whitefly-transmitted, bipartite closterovirus. Virology 208, 99-110. Liu, H.-Y, Wisler, G.C., and Duffus, J.E. (2000). Particle length of whitefly-transmitted criniviruses. Plant Dis., 84, 803-805. Bellows, T.S., Perring, T.M., Gill, R.J., and Headrick, D.H. (1994). Description of a species of Bemisia [Homoptera, Aleyrodidae].^««. Entomol. Soc.Am. 87, 195-206.
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6. Perring, T.M., Cooper, A.D., Rodriguez, RJ., Farrar, C.A., and Bellows, T.S. (1993). Identification of a whitefly species by genomic and behavioral studies. Science 259, 74-79. 7. Cicero, J.M., Hiebert, I., and Webb, S.E. (1995). The alimentary canal of Bemisia tabaci and Trialeurodes abutilonea (Homoptera, Stemorrhynchi [sic]: Histology, ultrastructure and correlations to function. Zoomorphology 115, 31-39. 8. Harris, K.F., Pesic-Van Esbroeck, Z., and Duffus, J.E. (1996). Morphology of the sweet potato whitefly, Bemisia tabaci (Homoptera, Aleyrodidae) relative to virus transmission. Zoomorphology 116, 143-156. 9. Celix, A., Lopez-Sese, A., Almarza, N., Gomez-Guillamon, L. and Rodriguez-Cerezo, E. (1996). Characterization of cucurbit yellow stunting disorder virus, a Bemisia tabaci-\xQnsmi\X.Qd closterovirus. Phytopathology 86, 1370-1376. 10. Duffus, J.E. (1965). Beet pseudo-yellows virus, transmitted by the greenhouse whitefly (Trialeurodes vaporariorum). Phytopathology 55, 450^53. 11. Coutts, R.H.A., and Coffin, R.S. (1996). Beet pseudo-yellows virus is an authentic closterovirus. Virus Genes n,\19-\U. 12. Tian, T, Soong, J., Wisler, G.C., Duflus, J.E., and Falk, B.W. (1996). Generation and cloning of specific cDNAs corresponding to four whitefly-transmitted viruses using RT-PCR and degenerate oligonucleotide primers corresponding to the closterovirus gene encoding the heat shock protein 70 homolog. Phytopathology 86, 1167-1173. 13. Snyder, R.G. (1995). Greenhouse tomatoes - the basics of successful production. In "Proceedings of the Greenhouse Tomato Seminar, Montreal, Canada," pp. 3-6. 14. Brown, J.K. (1994). Current status of Bemisia tabaci as a plant pest and virus vector in agroecosystems worldwide. FAO Plant Protection Bull. 42, 3-33. 15. Cohen, S., Duffus, J.E., and Liu, H.-Y. (1992). A new Bemisia tabaci (Gennadius) biotype in southwestern United States and its role in silverleaf of squash and transmission of lettuce infectious yellows virus. Phytopathology 82, 86-90. 16. Kao, J., Jia, L, Tian, T, Rubio, L. and Falk, B.W. (2000). First report of cucurbit yellow stunting disorder virus (CYSDV, genus Crinivirus) in North America. Plant Dis. 84,101. 17. Prabhaker, N., Toscano, N., Castle, S., and Henneberry, T. (1996). Evaluation of insecticide rotations and mixtures as a resistance management strategy for whiteflies. In "Silverleaf Whitefly: 1996 Supplement to the Five-year National Research and Action Plan," USDA-ARS Publ. 1996-01,95. 18. Prabhaker, N., Toscano, N., Castle, S., and Henneberry, T (1995). Hydroponic bioassay technique to monitor responses of whiteflies to imidacloprid. In "Silverleaf Whitefly: 1995 Supplement to the Five-year National Research and Action Plan," USDA-ARS Publ. 1995-01, 89. 19. Lopez-Sese, A.I., Gomez-Guillamon, M.L., (2000). Resistance to cucurbit yellowing stunting disorder virus (CYSDV) in Cucumis melo L. Hort.Sci. 35, 110-113. 20. Duffus, J.E., Larsen, R.C., and Liu, H.-Y. (1986). Lettuce infectious yellows virus—a new type of whitefly-transmitted virus. Phytopathology 76, 97-100. 21. Markham, RG., Bedford, I.D., Liu, S., Frolich, D., Rosell, R., and Brown, J.K. (1996). The transmission of geminiviruses by biotypes of Bemisia tabaci (Gennadius). In "Bemisia 1995: Taxonomy, Biology, Damage Control and Management" (D. Gerling and R.T. Mayers, eds.), pp. 69-75. Intercept Ltd., Andover, U.K. 22. Wisler, G.C., Duffus, J.E., Liu, H.-Y, and Li, R.H. (1998). Ecology and epidemiology of whiteflytransmitted closteroviruses. Plant Dis. 82, 270-280. 23. Rubio, L., Soong, J., Kao, J., and Falk, B.W. (1999). Geographic distribution and molecular variation of isolates of three whitefly-bome closteroviruses: Lettuce infectious yellows virus (LIYV), cucurbit yellow stunting disorder virus (CYSDV) and beet pseudo-yellows virus (BPYV). Phytopathology, 89, 707-711.
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24. Duffiis, J.E. (1995). Whitefly transmitted yellowing viruses of the Cucurbitaceae. In "Cucurbitaceae 94" (G.E. Lester, and J.R. Dunlap, eds.), pp. 12-16, Gateway Printing, Edinburgh. 25. Duffus, J.E. (1996). Whitefly-bome viruses. In "Bemisia 1995: Taxonomy, Biology, Damage Control and Management" (D. Gerling and R.T. Mayers, eds.), pp. 255-263. Intercept Ltd., Andover, U.K. 26. Tian, T., Rubio, L., Yeh, H.H., Crawford, B., and Falk, B.W. (1999). Lettuce infectious yellows virus: In vitro acquisition analysis using partially purified virions and the whitefly Bemisia tabaci. J. Gen. F/ra/. 80, 1111-1117. 27. Lewellen, R.T., and Skoyen, I.O. (1987). Registration of 17 monogerm, self-fertile germplasm lines of sugarbeet derived from three random-mating populations. Crop Sci. 27, 371-371. 28. Lewellen, R.T., and Skoyen, I.O. (1988). Registration of four monogerm, self-fertile randommated sugarbeet germplasms. Crop Sci. 28, 873-874. 29. McLain, J., Castle, S,, Holmes, G., and Creamer, R. (1998). Physiochemical characterization and field assessment of lettuce chlorosis virus. Plant Dis. 82, 1248-1252. 30. Duffus, J.E., Liu, H.-Y., Wisler, G.C., and Li, R.H. (1996). Lettuce chlorosis virus—a new whitefly-transmitted closterovirus. Eur J. Plant Pathol. 102, 591-596. 31. Duffus, J.E., Liu, H.-Y, and Wisler, G.C. (1996). Tomato infectious chlorosis virus—a new clostero-like virus transmitted by Trialeurodes vaporariorum. Eur J. Plant Pathol. 102, 219-226. 32. Wisler, G.C, Liu, H.-Y, Klaassen, VA., Duffus, J.E., and Falk, B.W. (1996). Tomato infectious chlorosis virus has a bi-partite genome and induces phloem-limited inclusions characteristic of the closteroviruses. Phytopathology 86, 622-626. 33. Li, R.H., Wisler, G.C, Liu, H.-Y, and Duffus, J.E. (1998). Comparison of diagnostic techniques for detecting tomato infectious chlorosis virus. Plant Dis. 82, 84-88. 34. Wisler, G.C, Li, R.H., Liu, H.-Y, Lowry, D.S., and Duffus, J.E. (1998). Tomato chlorosis virus: A new whitefly-transmitted, phloem-limited, bicomponent closterovirus of tomato. Phytopathology 88,402^09. 35. Schaefers, G.A., and Terry, E.R. (1976). Insect transmission of sweet potato disease agents in Nigeria. Phytopathology 66, 642-645. 36. Cohen, J., Franck, A., Vetten, H.J., Lesemann, D.E., and Loebenstein, G. (1992). Purification and properties of closterovirus-like particles associated with a whitefly-transmitted disease of sweet potato. Ann. Appl. Biol. 121, 257-268. 37. Larsen, R.C, Laakso, M., and Moyer, J.W (1991). Isolation and vector relations of a whiteflytransmitted component of the sweet potato virus disease (SPVD) complex from Nigeria. Phytopathology %\, 1157. 38. Pesic-Van Esbroeck, Z, Averre, CW, Schultheis, J.R., Daykin, M.E., and Milholland, R.D. (1997). North Carolina improved sweet potato foundation seed program. (Abstr.) Phytopathology 87, S77. 39. Liu, H.-Y, Li. R.H., Wisler, G.C, and Duffus, J.E. (1997). Characterization of Abutilon yellows virus—a new clostero-like virus transmitted by banded-wing whitefly (Trialeurodes abutilonea). (Abstr.) Phytopathology 87, S58-S59. 40. Larsen, R.C, Kim, K.S., and Scott, H.A. (1991). Properties and cytopathology of diodia vein chlorosis virus-a new whitefly-transmitted virus. Phytopathology 81, 227-232. 41. Karasev, A.V, Nikolaeva, O.V, Koonin, E.V, Gumpf, D.J., and Gamsey, S.M., (1994). Screening of the closterovirus genome by degenerate primer-mediated polymerase chain reaction. J. Gen. Virol 75,1415-1422. 42. Deng, T.-C, and Zettler, FW (1995). Incidence of three aphid-transmitted Z///ww viruses. (Abstr.) Phytopathology %5, 1137. 43. Li, R.H., Zettler, FW, Elliott, M.S., Petersen, M.A., Still, PE., Baker, C.A., and Mink, G.I. (1991). A strain of peanut mottle virus seedbome in bambarra groundnut. Plant Dis. 75,130-133.
CHAPTER 16
Classical Biological Control of Bemisia and Successful Integration of Management Strategies in the United States A. A. KJRK L. A. LACEY J. A. GOOLSBY
/. Introduction Since its description in 1889 by Gennadius, Bemisia tabaci Gennadius has been widely recorded on many plant hosts throughout the zone between latitudes 40° North and 40° South around the world [1,2]. Taxonomically it is treated as part of a species complex containing three species, and also as a new species, Bemisia argentifolii (Bellows and Perring), [3]. Bemisia argentifolii has been recorded from over 900 plant species in 74 families [4, 5]. Outbreaks of 5. argentifolii in Arizona, California, Florida, and Texas caused estimated crop losses in excess of $500 million in 1992 [6]. Bemisia tabaci s. 1 is resistant to most insecticides and, despite massive pesticide spraying, its numbers continue to increase. Besides causing rapid development of resistance to insecticides, pesticide treatments have an impact on nontarget organisms, leading to a severe reduction or elimination of natural enemies and further compounding the problem. The history of the occurrence, distribution, and severity of 5. argentifolii, the silverleaf whitefly (SLWF), in the United States is a curious but salutary one. First recorded in Florida in 1894 and in Texas and Georgia in the late 1940s, it was not known as a serious pest until 1986-1988 in Florida [7] and late 1989 in Texas and California [8]. The circumstances leading to the outbreaks are unknown but are probably related to the use of extensive pesticide applications, which reduced natural controls and enhanced resistance to chemical products. A run of mild winters might have allowed SLWF to overwinter on an increased weed species population 309
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[6]. Cole crops are grown as a winter crop in the southern United States. Silverleaf whitefly was unknown on these before the late 1980s. After its accidental introduction, SLWF became adapted to cole crops in the agricultural region of southern California and was recognized as a problem when several visible disorders of cruciferous crops appeared at the same time [9]. At this time Perring et al [9] indicated that cole crop producers were using multiple pesticide applications against whiteflies for the first time and that there was particularly severe damage to cauliflower and broccoli, Brassica oleracea L. [10]. In southern Texas severe damage to cabbage was also noted at the same time [11]. In addition to sustaining considerable direct damage, cole crops act as reservoir plants for overwintering whitefly populations, which move onto melons in spring when the crucifers are harvested. A number of physiological disorders leading to weight loss of cole crops have been associated with the presence of immature whitefly stages: in Hawaii, yellowing and stem blanch in kai choy [12] and in 1990 in Arizona, white streaking disorder of broccoli and cauliflower [8]. The appearance of an apparently new gemini virus, cabbage leaf curl virus, followed the arrival of B. argentifolii in southern Florida [13]; leaf yellowing was reported by Bedford et al (14,15) on B. argentifolii-infQstQd cabbage and cauliflower. Increased transportation of infested ornamental plants fi"om greenhouses around the country may have resulted in widespread distribution of resistant SLWF. Recent genetic evidence points to the expansion in range of the B biotype or B. argentifolii from an ancestral Mediterranean home throughout the world. Without a doubt, increased transportation of ornamental plants as seedlings and full-grown plants has led to this global expansion. As a rule, natural enemies do not travel with their host and in the case of SLWF, extraordinary attempts at obtaining clean plants for export would have eliminated them. Silverleaf whitefly, however, because of its very comprehensive resistance to pesticides, would have traveled with its host plant. Crop losses mainly of cotton and vegetables in the Rio Grande Valley of Texas reached $100 million by 1991. The situation was grave and a series of planning meetings led to a 5-year national research and action plan for development of management and control methodology for silverleaf whitefly. The plan identified six areas of priority research, including biocontrol. Biological control based integrated pest management of SLWF in the United States using imported natural enemies is presented in this chapter, the main sections of which include Foreign Exploration, Pathogenic Fungi for Biological Control of Silverleaf Whitefly, and Evaluation and Release of Silverleaf Whitefly Parasitoids. The diverse landscapes and agricultural systems present worldwide suggested a potential for many suitable habitats for whiteflies and natural enemies. From 1991 to 1998, exploration for natural enemies for biological control of B. argentifolii was conducted throughout the world by the U.S. Department of Agriculture-Agricultural Research Services (USDA-ARS)'s European Biological Control Laboratory (EBCL) based at Montpellier, France [16-18]. As a component of the SLWF natural enemy complex, the potential of the Aphelinidae (Hymenoptera) insect
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parasitoids as biocontrol agents was considered to be very high. They are widespread, and their ability to find and attack whitefly nymphs is well documented [4, 5]. Owing to the feeding strategy oi Bemisia (e.g., a piercing and sucking feeding apparatus), the only pathogens with microbial control potential for Bemisia spp. are the entomopathogenic fungi. A wide variety of fungi have been isolated from B. argentifolii and B. tabaci, but only a few have been developed as microbial control agents [17]. Species in the order Hyphomycetes offer the most potential for commercial production and ease of application. They can be grown on artificial media, have relatively good shelf lives, especially when refrigerated, and can be applied using conventional pesticide application equipment [19, 20].
//. Foreign Exploration A.
Collections
Collection sites were selected initially based on climate matching with whiteflyinfested areas in Arizona, California, and Texas [21]. First collections were made in June of 1991 in Greece, in November and December of 1991 in southeastern Spain, and then in the Indian subcontinent, which was believed to be the center of origin of B. tabaci based on the apparent abundance of its natural enemies. In both cases, very large amounts of leaves infested by SLWF nymphs were collected. It was immediately apparent that a simple technique was required to prevent rotting of the leaves and subsequent loss of parasitized nymphs. The objective was to deliver parasitized whitefly nymphs on leaves to quarantine facilities in the United States by express mail and air freight, sometimes resulting in transit periods as long as 12 days. Shipment of very large numbers reduced the chance of complete loss of material. Absorbent rather than nonabsorbent material was required for wrapping. Preshipment drying of leaves and cooling also improved survival during shipping. Paper bags that absorbed condensation and allowed for ventilation were used. Leaves were individually wrapped in tissue paper before being placed in the paper bags. In the case of material from very humid areas such as southern India, an inner layer of children's diapers was spread throughout the shipment box to absorb ambient humidity. Paper bags containing the infested leaves were placed in large cool boxes or containers with "blue ice" for shipment. Refinements of this basic system led to a very high emergence rate at the receiving quarantine facilities. Field collections were made by pulling infested leaves from plants, placing them directly into paper bags, and storing them in cool boxes containing blue ice. Walk-in refrigerators and room refrigerators in hotels were used to hold material until shipment. Studies were carried out to elucidate optimal temperatures for storage of collected material. Lacey et al. [22] showed that the eggs and pupae of SLWF and the
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pupae of the parasitoid Encarsia formosa survived after prolonged storage at 10°C. Shipments were made during and at the end of long trips. Where no suitable means of shipment was found, material was hand carried to Montpellier and shipped from there. Traceable means of shipment were used whenever possible so that shipments could be tracked during transit. The use of local express mail was not reliable; certain national and international express mail services were adequate, and air freight, where feasible, was always the best. All samples were collected and stored separately, on the basis of host plant and locality. Exploration for fungi with entomopathogenic activity against Bemisia was also conducted; nymphs suspected of being infected were stored over silica gel for transportation. Collections of infected nymphs were also made in the United States and Mexico. Permits for importing infested, parasitized, and infected material into France and the United States were obtained. It was sometimes necessary to obtain export permits for this material from the countries of collection. In these cases, we depended on and received help from in-country cooperators. Collections were shipped in insulated boxes by air freight to the Mission Biological Control Center (MBCC) in Mission, TX and to the USDA-ARS quarantine facility in Montpellier, France, where parasitoids emerged. From 1991 to 1998, the processing of 235 accessions resulted in 41 cultures of parasitic Hymenoptera, 17 of which were determined to be distinct species (Table I). Twenty-five countries were visited at least once, and collections were made in hot dry to hot humid climates. Hundreds of isolates of the fringal pathogen Paecilomyces fumosoroseus came from Bemisia populations in five countries and were stored at the USDAARS laboratory in Ithaca, NY. The collections of pathogenic fungi in North America resulted in the isolation of several dozen strains with good insecticidal activity against Bemisia [23-26]. B.
Taxonomic Identification by Morphological Characters
Microscope slides of adult parasitoid specimens and whitefly nymphal case remains were prepared by the techniques described by Noyes [27] and Martin [28], respectively. Whitefly specimens were sent to S. Nakahara at the Systematic Entomology Laboratory in Beltsville, MD and to R. Gill at the California Department of Food and Agriculture in Sacramento, CA. All Encarsia species were sent to M. Schauff at the Systematic Entomology Laboratory in Beltsville and to J. B. WooUey at Texas A&M University, College Station, TX. Eretmocerus species were sent to M. Rose and G. Zolnerowich at Texas A&M University. Table 1 gives the details of the insect parasitoids collected, as well as their identifications and DNA patterns (see section IV on evaluation and release of silverleaf whitefly parasitoids). C. Whitefly Populations and Apparent Field Parasitism Examples of apparent field parasitism shown by parasitoids in Spain and Thailand are given here.
16.
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Table I
Parasitoids of Silverleaf Whitefly Evaluated for Biological Control
Parasitoid
Accession DNA no. pattern
Encarsia sp. nr. strenua Encarsiaformosa
M92018 M92017
EN-1 EN-2
Encarsia formosa
M92030
EN-2
Encarsiaformosa Encarsiaformosa
M94051 M94089
EN-2 EN-2
Encarsia sophia (^ transvena) Encarsia sophia
M94017
EN-3
M94019
EN-4
Encarsia sophia
M94041
EN-5
Encarsia sophia
M94047
EN-5
Encarsia sophia Encarsia adrianae
M95107 M94024
EN-5 EN-6
Encarsia sophia Encarsia lutea Encarsia lutea Encarsia lutea Encarsia lutea Encarsia lutea
M93003 M93064 M94096 M94107 M94115 M94129
EN-7 EN-10 EN-10 EN-10 EN-10 EN-10
Encarsia lutea
M96044
EN-10
Encarsia sophia
M94014
EN-11
Encarsia sophia Encarsia nr. pergandiella Encarsia nr. hispida
M94016 M94055
EN-11 EN-15
M94056
EN-16
Encarsia sp. (parvella group) Eretmocerus spp Eretmocerus mundus Eretmocerus mundus Eretmocerus mundus Eretmocerus mundus Eretmocerus mundus
M95001
EN-18
Eretmocerus mundus
Collection locality India, Parbhani Greece, Angelohori Egypt, Nile Delta Thailand, Saen Italy, Borgo Corso Taiwan, Shan-Hua Taiwan, Shan-Hua Thailand, Chiang Mai Malaysia, Kuala Lumpur Pakistan, Multan Thailand, Kampang Saen Spain, Murcia Cyprus, Mazotos Italy, Testa Di Lespe Israel, Givat Haim Israel, Dead Sea Spain, Mazarron Casas Nuevas Sicily, Ragusa Philippines, Benguet Taiwan, Shan-Hua Brazil, Sete Lagoas
Host plant(s)
Bean
Autoparasitoid Uniparental
Lantana
Uniparental
Snakeweed Tomato
Uniparental Uniparental
Soybean, tomato Soybean, tomato Poinsettia
Autoparasitoid
Mussaenda sp. Cotton Snakeweed
Autoparasitoid Autoparasitoid Autoparasitoid
Lantana Lantana Eggplant Cotton Lantana Ipomea sp.
Autoparasitoid Autoparasitoid Autoparasitoid Autoparasitoid Autoparasitoid Autoparasitoid
Solanaceous weed White potato
Autoparasitoid
Poinsettia Poinsettia/ soybean Poinsettia/ Brazil, Sete Lagoas soybean Dominican Republic, Tomato Azua
M92014 ERET-1 Spain, Murcia M92019 ERET-1 India, Padappai M92027 ERET-1 Egypt, Cairo M93058 ERET-1 Taiwan, Tainan M94092 ERET-1 Italy, Castel Gondolfo M94097 ERET-1 Italy, Testa Di Lespe
Biology
Cotton Eggplant Lantana Tomato Ipomea sp. Hibiscus Eggplant
Autoparasitoid Autoparasitoid
Autoparasitoid Autoparasitoid Uniparental Uniparental Autoparasitoid
Biparental Biparental Biparental Biparental Biparental Biparental continued
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A. A. KIRK, L. A. LACEY, AND J. A. GOOLSBY
Continued
Table I
Parasitoid
Accession DNA no. pattern
Eretmocerus mundus
M94103
ERET-1
Israel, Gat
Eretmocerus mundus
M94125
ERET-1
Israel, Golan Heights
Eretmocerus mundus Eretmocerus mundus Eretmocerus mundus Eretmocerus hayati Eretmocerus melanoscutus Eretmocerus melanoscutus Eretmocerus melanoscutus Eretmocerus eremicus Eretmocerus staufferi Eretmocerus tejanus Eretmocerus hayati
M94124 M96028 M97046 M93005 M94036
ERET-1 ERET-1 ERET-1 ERET-2 ERET-3
M94040
ERET-3
M95097
ERET-3
Israel, Negev Desert Sicily, Santa Groce Cyprus, Nicosia India, Thirumala Thailand, Chiang Mai Thailand, Kampang Saen Taiwan, Tainan
M94001 M94002 M94003 M95012
ERET-4 ERET-5 ERET-6 ERET-10
Brawley, CA College Station, TX Mission, TX Pakistan, Multan
Eretmocerus hayati Eretmocerus sp. nr. furuhashii Eretmocerus emiratus Eretmocerus sp.
M95105 M95098
ERET-10 ERET-11
Pakistan, Multan Taiwan, Tainan
M95104 M96076
ERET-12 ERET-13
United Arab Emirates Ethiopia, Melka Werer
1.
SPAIN 1991 AND
Collection locality
Host plant(s)
Biology
Kohhabi, Sonchus sp. Euphorbia spp.. Melons Cucumber Eggplant Lantana
Biparental
Chromolaena odorata Cotton
Biparental Biparental Biparental Biparental Biparental Biparental Biparental
Tomato
Biparental
Okra Tomato Cabbage Eggplant, cotton Eggplant Tomato, cabbage Okra Cotton
Biparental Uniparental Biparental Biparental
Biparental Biparental Biparental
1992
Collections were made in Murcia, southeastern Spain, an intensively farmed area in a dry Mediterranean climate resembling that of southern California. Ten leaves were taken from ten cotton plants in Murcia, which were the source of parasites in December of 1991 and ten leaves from ten Sonchus oleraceae plants under the cotton in November of 1992. Five counts were made across each leaf Each counting area was 1 cm^, and all whitefly pupae within the area were counted. Parasitized nymphs were recognized by the presence of an aphelinid larva, the presence of meconia, and the typical circular hole in the SLWF nymph made by emerging aphelinid adults. Nymphs parasitized by Eretmocerus were citron in color, versus black or gray for those parasitized by Encarsia spp. Leaves were senescent on the cotton, and pesticide spraying had stopped 2 weeks before the collections were made from cotton leaves in 1991. The Sonchus was underneath cotton plants that were still being sprayed at the time of collection in 1992. Populations of whitefly nymphs on cotton were (n = 50 counts) 12 nymphs/cm^.
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standard error (SE) 1.61, and {n = 50) 13 nymphs/cm^ on Sonchus, SE 1.21. Percentage parasitism by Eretmocerus mundus of whitefly nymphs on cotton was {n = 50) 39%, SE 4.24 and on Sonchus (n = 50) 44%, SE 4.9, despite the spraying. Eretmocerus mundus from this collection were found to be tolerant of several insecticides when evaluated in Texas [29]. 2.
THAILAND 1994
In 1994 collections were made from 11 host plant species in 19 localities throughout Thailand. Six Bemisia parasitoid emerged. The localities were in tropical to subtropical areas that have heavy seasonal rains and a distinct dry season resembling the climate of southern Texas. Parasitized nymphs were identified as described previously. All whitefly eggs and nymphs within the sampling area were counted. Whitefly nymphal populations were generally sparse on weeds and field crops (one or fewer, nymphs/per cm^ of leaf. Numbers of whitefly eggs, an indicator of current whitefly activity, ranged from 0 to 12 per cm^ of leaf in the field. Percentage parasitism under field conditions ranged from 0 to 65%. An Eretmocerus sp. parasitized 65% of Bemisia on Physalis minima L., a solanaceous weed.
///. Pathogenic Fungi for Bioiogicai Controi of Silverleaf Whitefly A.
Applied and Basic Research on Factors Influencing Entomopathogenic Activity
The majority of basic studies on fungal pathogens of Bemisia have been conducted on P. fumosoroseus. The focus of laboratory and greenhouse research has been the elucidation of factors that enhance or reduce insecticidal activity. These include a wide range of biotic and abiotic factors. 1.
FUNGAL E\CTORS
a. Isolate, The P. fumosoroseus isolates collected from whiteflies comprise a genetically diverse group [30]. Those that have been bioassayed demonstrated a broad range of insecticidal activity against 5. argentifolii [26, 31-33]. Two of the most active isolates in terms of rapid growth and insecticidal activity are the Pfr 97 isolate used in the Thermo Trilogy product [23] and the EBCL Pfr 42 isolate [33]. Although Beauveria bassiana is rarely reported from Bemisia [34], laboratory assays and field evaluation of several isolates demonstrated good insecticidal potential against SLWF [26, 35].
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A. A. KIRK, L. A. LACEY, AND J. A. GOOLSBY
b. Propagule, Although recently produced blastospores demonstrate greater insecticidal activity than conidia in some cases [33, 36], their persistence can be significantly curtailed by drying [36]. The age of aerial conidia can also have a marked effect on their viability and infectivity Hall et al [37] observed that aerial conidia of R fumosoroseus from young cultures germinate faster when compared with those from older ones. c. Media. Jackson et al. [36] and Vidal et al [38] investigated the effect of media on blastospore production and infectivity. The MS medium [34] was one of the most effective in producing the maximum number of propagules (5 x 10^ blastospores per milliliter) in the relatively short time of 72 hours [39]. The MS medium contains a complex mineral and vitamin mixture and a carbon to nitrogen ratio of approximately 36:1 (casamino acids and glucose). 2.
PHYSICAL FACTORS INFLUENCING GROWTH AND ACTIVITY
a. Humidity. Laboratory assays conducted by Wraight et al [35] demonstrated the capacity of R fumosoroseus and B. bassiana to infect SLWF nymphs on excised hibiscus leaves incubated at relative humidities as low as 25% (23°C). b. Temperature. The effect of temperature on growth of/?^mo5'or(0^^w^ has been quantified by Vidal et al [40] for several strains isolated from Bemisia and other host insects. There was a correlation between the geographical origin of the various isolates and the optimum temperature for growth and the range of temperatures that could be tolerated. Those from the Indian subcontinent grew optimally at 25° to 28°C and were the most tolerant to high temperatures (up to 35°C), whereas the optimal temperature for those from temperate climates was 20° to 25°C. c. Ultraviolet Light. Research conducted by Fargues et al [41, 42] and Smits et al [43] on the susceptibiHty of R fumosoroseus to ultraviolet (UV) radiation indicates that the ftingus is sensitive to UVB (280-320 and 295-320 nm) and UVA irradiance (320-400 nm). The authors concluded that UVB was the most detrimental waveband of sunlight in terms of persistence of the fungus. 3.
BioTic FACTORS
INFLUENCING ACTIVITY
a. SLWF Stage. Osborne et al [23] demonstrated that all stages of SLWF were susceptible to R fumosoroseus. However, Hall et al [44] observed that first instar nymphs of Bemisia were refractory to infection. Lacey et al [33] reported significant but low ovicidal activity of blastospores of the EBCL Rfr 42 isolate. b. Host Plant Effect Comparisons of Rfr 97 against SLWF on cucumber, tomato, and cabbage showed no significant difference in efficacy as a fiinction of
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317
host plant [39]. Secondary plant compounds (allelochemicals) can have detrimental effects on germination and growth of P. fumosoroseus [45]. Because many allelochemicals act as fungicides, they may also affect the activity of entomopathogens directly or indirectly by altering host susceptibility. Whether they can actually affect the efficacy of R fumosoroseus or other fungi against SLWF can only be definitely determined with bioassays on host plant varieties with varying levels of allelochemicals in identical microclimatic conditions. B.
Field Studies 1.
NATURAL
OccuioiENCE
Large epizootics caused by P. fumosoroseus in Bemisia spp. have been observed in several locations in the Indian subcontinent, the Imperial Valley of California, and the lower Rio Grande valley of Texas [17, 34, 46]. Although they may drastically reduce some populations, epizootic predictability is uncertain, and they may occur after economic thresholds have been surpassed or transmission of plant pathogens has taken place. 2.
APPLICATION OF FUNGI FOR MICROBIAL CONTROL OF
SLWF
The inundative application of entomopathogenic fungi provides a degree of predictability but relies on the appropriate microclimatic conditions in order to be effective. Certain environmental conditions that provide high humidity and optimal temperatures for germination and infection or for protection from UV inactivation of spores will facilitate better and more predictable control. The effectiveness of environmental manipulation to enhance infection will depend heavily on the agroecosystem and climate. For example, manipulation will be most feasible in greenhouses, where high humidity and optimal temperatures can be maintained. The majority of field studies on entomopathogenic fixngi for control of SLWF have been conducted on P fumosoroseus and B. bassiana, Both species of fungi have provided effective control of SLWF in cotton [47, 48], cucurbits [35], and greenhouse crops [39, 49]. Numerous other examples are informally published in the yearly supplements to the 5-year National Research and Action Plan for Silverleaf Whitefly. The most detailed field tests for control of SLWF to date have been conducted by Wraight et al [35] on various cucurbit crops in the lower Rio Grande valley in Texas and by Vidal et al. [39] on greenhouse crops. Multiple (five to seven) applications of 5 x 10^^ conidia per hectare of P fumosoroseus and 5. bassiana at 4to 5-day intervals provided better than 90% control of large nymphs [35]. A single appHcation of 5 x lO'* conidia per cm^ of P fumosoroseus (Pfr 97) to greenhouse tomatoes and cucumbers resulted in 82 to 88% control of SLWF at 14 days posttreatment. Nearly 100% of fungus-killed nymphs sporulated within 14 days posttreatment, providing an important source of secondary inoculum.
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A. A. KIRK, L. A. LACEY, AND J. A, GOOLSBY
Because Bemisia nymphs are attached to the underside of leaves, application strategies must be used that deliver the majority of conidia to this surface. Wraight and Carruthers [20] and Wraight et al [35] address application systems that enable treatment of the undersurface of leaves. Wraight et al. [35] also present a method for measurement of coverage using plastic coverslips attached to the undersurface of host plant leaves. Two commercial products based on B. bassiana that have been used for control of Bemisia are available in the United States from Troy Biosciences, Inc. (Phoenix, AZ) and from Mycotech Corp. (Butte, MT). Paecilomyces fumosoroseus is available from Thermo Trilogy Corp. (Columbia, MD) and Agrobiologicos del Noroeste, S.A. (Culiacan, Sinaloa, Mexico). Garza Gonzalez and Arredondo Bernal [50] reported insecticidal activity of Verticillium lecanii for SLWF, but no large-scale field trials have been reported against Bemisia. E. T. Meeks and J. Fransen (cited in Lacey et al. [34]) evaluated several Aschersonia species with activity for Bemisia. Steinberg and Prag [51] reported on the combined activity of Aschersonia aleyrodis and the coccinellid predator Delphastus pusillus in controlling B. tabaci in a greenhouse. The efficacy of these fungi and their specificity for whiteflies and scale insects warrant further attention. No commercial products based on Aschersonia spp. are currently available. C.
Compatibility of Entomopathogenic Fungi with Other Natural Enemies
Brooks [52] has reviewed the interaction of entomopathogenic fungi and a number of beneficial organisms, including predators and parasitoids. The compatibility of fungi, particularly /? fumosoroseus, V lecanii, and A. aleyrodis, and insect natural enemies of whiteflies and aphids has been reviewed by Lacey et al. [34, 53]. Several mechanisms minimize the antagonistic interaction between entomopathogenic fiingi and parasitoids, including the production of antimycotic substances by internal parasitoids and the avoidance of infected hosts by ovipositing females. Poprawski et al. [54] and Steinberg and Prag [51] have reported the compatibility ofB. bassiana and A. aleyrodis with coccinellid predators of Bemisia, respectively.
IV. Evaluation and Release of Sllverleaf Whitefly Parasitoids A.
Prerelease Surveys and Evaluations
Prior to release of exotic natural enemies for SLWF, prerelease evaluations were conducted in California [55], Florida [56, 57], South Carolina [58], and Texas [59]. A complex of parasitoids in the genera Encarsia and Eretmocerus was found attacking B. tabaci on a wide variety of hosts. Hunter et al. [60], using mating and
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allozyme studies, found that the indigenous Eretmocerus attacking B. tabaci was actually a complex of species. Several of the new species were described, including Eretmocerus eremicus Rose & Zolnerowich from Arizona and California and Eretmocerus tejanus Rose & Zolnerowich from Texas [61]. Several Encarsia species were recovered from B. tabaci, including Encarsia pergandiella Howard, Encarsia formosa Gahan, Encarsia luteola Howard, Encarsia meretoria Gahan, and Encarsia nigricephala Dozier. The autoparasitoid E. pergandiella was the most common indigenous parasitoid species parasitizing SLWF in the southeastern United States. Encarsia pergandiella was not collected in surveys of the Imperial Valley of California despite the presence of its primary host, Trialeurodes abutilonea. B.
Quarantine Methods
Natural enemies ofB. tabaci imported into the MBCC were placed in cages containing B. tabaci infested plants. Rearing conditions were 24° to 29°C, 50 to 70% relative humidity, and a light regime of 14:10. Unique species or biotypes of natural enemies were isolated into separate emergence containers by date, geographic location, and host plant. Individuals from each candidate population were collected from the emergence containers and used to start separate cultures. Foreign material arriving into quarantine showed extensive diversity, especially in the genus Eretmocerus [62, 63]. The methods used at the USDA-Animal and Plant Health Inspection Service (APHIS) quarantine facility in Mission, TX are described here. To best handle the issues of cryptic species, species complexes, and the need to initiate pure cultures representing the maximum available diversity of natural enemies, a unique quarantine protocol was developed, which integrated biosystematics and molecular techniques. Foreign collections were categorized in quarantine, by plant type, site location, and the macrocharacters of the parasitic Hymenoptera and Aleyrodidae. Only parasitoids reared from individuals of the B. tabaci complex met the requirements for obtaining release permits, as stated in the environmental assessments of the genera Eretmocerus and Encarsia [64]. Furthermore, the imported species must have had biology described as uniparental, biparental, or autoparasitoid. Species that displayed obligate hyperparasitism of other taxa were not considered suitable for release [65]. The requirements of the environmental assessments were intended to identify the parasitoid species with the most specificity to the B. tabaci complex. Species that met these criteria were acceptable for processing using our quarantine protocol. Eretmocerus and Encarsia were separated into distinct groups according to the morphology of the pupae and adult females. Individuals from each unique accession were immediately characterized at the MBCC Genetics Laboratory by using a rapidly amplified polymorphic DNA (RAPD) procedure that employed primers C04 and AlO [66, 67]. Detailed methodology and representative electrophoretic gel patterns for Eretmocerus and Encarsia parasitoids
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are presented in Legaspi et al [62]. Cohorts of the original parental material were sent to cooperating systematists. Information from the collaborating systematists and geneticists allowed for characterization of quarantine material while the original parental cohort was still alive. Typically, material was characterized by both methods within 2 to 3 days after acceptance into quarantine. Unique parasitoid accessions were set up in pure cultures reared on the local B. tabaci with hibiscus var. Kona pink, Hibiscus rosasinensis L., as the plant host. Duplicate accessions were combined or, in the later stages of the program, processed only for reference purposes. Representatives of all the accessions were cryogenically stored at the MBCC Genetics Laboratory, and vouchered at Texas A&M University, Department of Entomology Collection in College Station, TX and the USDA-ARS, Systematic Entomology Laboratory in Washington, D.C. Some species, such as Eretmocerus mundus Mercet, were widely distributed in the Old World and displayed the same RAPD banding pattern from all 18 locations. By comparison, Eretmocerus emiratus Rose & Zolnerowich was collected from only one locality and displayed a unique banding pattern. Apart from differences in RAPD patterns, new species were only distinguishable from each other by minute differences in the first ftmicular antennal segment of the females. Screening by RAPD techniques was the only method available for distinguishing such cryptic species. Without the use of this DNA-based technology, multiple cultures of widely distributed species such as E. mundus would have used most of the quarantine resources. Resources were conserved, which allowed for new species to be imported later in the program. With both RAPD patterns and morphological identifications available, it became apparent that unique RAPD patterns directly corresponded to distinct species for both Eretmocerus and Encarsia accessions. The one exception appeared to be Encarsia transvena Timberlake (now Encarsia sophia Girault). This species was collected in many different geographic regions and was characterized by five separate banding patterns (Table 1). However, Viggiani (personal communication) has found some crossing incompatibility between the Spanish and Pakistani populations of E. sophia, which is correlated with morphometric differences. It is possible this could reflect species level differences in the two E. sophia populations, just as suggested by the results of the RAPD analyses. C.
Quarantine Screening
A total of 38 exotic and two native parasitoids were evaluated at the quarantine facility in Mission, TX [63]. The purpose of the screening was to assess the fecundity of the parasitoid species on selected crop plants. Promising species were then prioritized for subsequent field evaluations. The plant varieties chosen, owing to the considerable economic losses that occurred annually to these crops, were cantaloupe melons (Cucumis melo L. cv 'Perlita'), cotton {Gossypium hirsutum L. cv
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'Delta Pine 51'), and broccoli (Brassica oleracea L. cv 'Patriot'). Highest attack rates were found for Encarsia nr. pergandiella (Brazil) and Eretmocerus mundus Mercet (Spain) on melons, for Eretmocerus hayati Rose & Zolnerowich (Pakistan) on cotton, and for E. mundus (Spain) on broccoli. In the laboratory, these three exotic parasitoids attacked significantly greater numbers of hosts than the native species ofE, pergandiella and E. tejanus. D.
Field Evaluation
Following several years of foreign exploration in the early 1990s, a wealth of parasitoid diversity was in culture at Mission, TX, Gainesville, FL, and Riverside, CA [63, 68, 69]. Various teams of researchers became involved in screening the diversity of natural enemies under localized field conditions. In the desert southwest of the United States, Hoelmer [70] evaluated several of the imported Eretmocerus spp. in field cage trials with melons, cotton, broccoli, and alfalfa as whitefly hosts. Four imported species, E. emiratus Rose & Zolnerowich (M95104 United Arab Emirates), E. mundus (M92014 Spain), and Eretmocerus sp. (M96076 Ethiopia) performed better than the indigenous Eretmocerus eremicus. In the lower Rio Grande valley of Texas, field evaluation of several introduced Eretmocerus spp. on cotton, melons, broccoli and cucumbers produced significantly different results from those of similar studies in California [63, 71]. Eretmocerus hayati Rose Si Zolnerowich (M95012 Pakistan) and E. mundus (M92014 Spain) were found to be more effective under field conditions than the indigenous E. tejanus. Eretmocerus hayati performed well on all crop types except cole crops, where E. mundus dominated. Four populations of Encarsia Sophia from Spain (M93003), Thailand (M94041), Malaysia (M94047) and Pakistan (M95107), as well as Encarsia nr. strenua from India (M92018), were evaluated in the Imperial Valley of California with use of field cages [72]. Encarsia Sophia (M93003) from Spain performed best in broccoli and cantaloupe, with E. Sophia (M95107) from Pakistan parasitizing significantly more whitefly nymphs on cotton. Recoveries of the E. sophia populations were confirmed by using RAPD profiles. The imported Eretmocerus spp. consistently performed the best in field trials. Selection of the most promising imported Eretmocerus spp. from field evaluation studies led to implementation of large-scale field trials. It was proposed that seasonal augmentative releases of natural enemies could overcome the seasonal lack of parasitoids in the ephemeral agroecosystems where SLWF was most damaging [73]. Large-scale augmentation programs were developed in California and Texas to evaluate the efficacy of early season releases oi Eretmocerus spp. in cantaloupe melons. Simmons et al [74] demonstrated that augmentative release ofE. emiratus could successfully control SLWF in cantaloupe melons and that the parasitoids could be integrated with subsurface applications of Admire TM (imidacloprid). Parasitoid-inoculated seedling transplants, "banker plants," were developed to
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make augmentation of parasitoids in short-season vegetable crops more efficient [75]. Banker plants could be preinoculated in the greenhouse for subsequent transplanting in the field. Application of Admire 1 week postinoculation with parasitoids was shown to have no significant effect on emergence. Encouraging results from the seasonal augmentation programs led to the development of areawide biological control-based integrated pest management (BCIPM) programs directed against SLWF. Inoculative releases into key crops such as spring melons could sufficiently suppress SLWF populations and in turn produce parasitoids for dispersal into surrounding summer and fall crops. Integration of selective insecticides and augmented parasitoids led to season-long control, thus avoiding late season SLWF population outbreaks. Using this BC-IPM approach, melon fields could be transformed at season's end from net SLWF producers to parasitoid refuges. Large-scale implementation of this approach was tested in Arizona, California, and Texas during 1998 and 1999. Further field testing is needed to determine the efficacy and economics of this approach. In the San Joaquin Valley of California, a similar areawide augmentation program was developed to suppress SLWF in cotton. Field surveys determined that citrus groves were the most important overwintering sites for SLWF. In the San Joaquin Valley, blocks of citrus are often isolated and surrounded by cotton fields. Following defoliation of cotton, citrus, which is normally a marginal host, is one of the few hosts available for SLWF. Releases of Eretmocerus spp. during the fall to attack the overwintering population in citrus were tested [76]. Several million parasitoids (E. emiratus, E. mundus, and E. hayati) were released, first into adjacent cotton, then into citrus. Parasitoids were found attacking silverleaf whitefly on cotton, citrus, and adjacent weeds through the summer of 1998. E.
Establishment of Parasitoids—Classical Biological Control
Establishment of imported parasitoid species for SLWF in the United States has varied widely by geographic region. Four species of Eretmocerus are reported to be established. In California, E. emiratus and Eretmocerus sp. nov. (M96076 Ethiopia), both from hot dry climates similar to that of southern California, have been recovered consistently [77]. Gould et al [78] recorded in-season reproduction of the same species in Arizona. Releases in Arizona were made in urban areas on ornamental plantings to mitigate the effects of host plant discontinuity and broad-spectrum pesticide use. In Texas, a survey program for establishment of exotic parasitoids was implemented by placing "sentinel" plants, which were preinfested with immature whiteflies, in field locations to sample parasitoid species composition. Locations across the lower Rio Grande valley of Texas were selected that represented a varied mix of agricultural and urban sites. Muskmelon wdiV.perlita, followed by cotton in the summer months, was used for sentinels. Males were mounted on slides to determine if they were exotic or native. The antennal pedicel of males of introduced species is uniformly fuscous, compared with an amber col-
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oration in indigenous E. tejanus males. This character was used to determine the overall percentage of indigenous versus introduced Eretmocerus spp. The percentage assumes a standard female to male sex ratio of 60:40 across Eretmocerus spp. Female Eretmocerus were used for species determinations. Species level determinations were made by using slide mounts and keys to the imported species oiEretmocerus, RAPD analyses, or satellite DNA probes [67, 79, 80]. Results of the sentinel plant recovery survey showed a dramatic increase in the numbers of introduced Eretmocerus spp. At the beginning of the survey in June of 1997, native E. tejanus represented more than 95% oiEretmocerus species. Three months later, during the fall of 1997 (August-October), exotic populations began to increase, representing 85% {n = 700) of the Eretmocerus spp. recovered. Analysis of the female Eretmocerus using both morphological techniques and molecular genetics identified three species: E. tejanus, E. mundus, and E. hayati. The two exotic species, E. mundus and E. hayati, from areas with hot seasonally humid climates similar to that of southern Texas, are widely established across the Rio Grande flood plain [63].
V
Concluding Remarks
There are several reasons for the successful implementation of the biological control program against silverleaf whitefly in the United States. The USDA-APHIS-ARS "conception to success" model for a biological control program that consists of program planning, progress meetings, and identification of key people to implement the plan is one. In this case scientists from the ARS EBCL in Montpellier, France carried out the foreign exploration. The quarantine and field evaluations and RAPD profiles of the insect parasitoids in the United States were made at the APHIS Mission Biological Control Center in Texas, which received and reared parasitoids. Identifications of the parasitoids were carried out at the USDA Systematic Entomology Laboratory in Washington, D.C. The California Department of Food and Agriculture is involved in the field evaluations at the producer level. Cooperators in France, at the ARS Weslaco Texas Research Station, and at many universities in the United States carried out experimental work, particularly on the fungal pathogens. This team effort enabled implementation of exotic parasitoids into the field within 5 to 7 years. It will be sometime before the potential distribution of these parasitoids has been attained. The parasitoids, which showed high field parasitism in their areas of origin, have been shown to be more efficient at killing Bemisia nymphs than the native parasitoids. A large amount of ecological work that is not reported here has identified the habitats in which Bemisia overwinters. These overwintering populations constitute the reservoir for infestations later in the season. Large-scale releases directed against this reservoir population resulted in a decrease
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of Bemisia populations and the establishment of the efficient exotic parasitoids in California and Texas. (Although isolates of pathogenic fungi are not in widespread use, concerted progress was made in evaluating and testing such isolates for use in biocontrol of SLWF.) Using RAPD profiles in quarantine as a way of measuring and identifying diversity appears to be an excellent means of making preliminary separations of geographic populations or strains, and possibly new species. It proved possible in California and Texas to track the presence and establishment of exotic parasitoids in the field by using RAPD analyses and morphological techniques. Biological control programs may be well served if RAPD profiles could be included as part of the protocol in the quarantine phase of the program. Additional field studies are needed to quantify the impact of the biological control agents. Quantifying the benefits of the program to agriculture and the environment should be compared with potential nontarget impacts of the agents. The silverleaf whitefly program was unique in that prerelease studies were conducted prior to release of the introduced species [59, 81, 82]. A thorough postrelease evaluation program of SLWF agents could provide the scientific evidence to facilitate a productive discussion between the biological control community and public policy makers. Bemisia numbers have decreased year by year since 1997 in California and Texas. Classical biological control has an important role to play when used to suppress reservoir populations of SLWF. There is clear evidence, from the information presented here, that augmentative releases of exotic parasitoids, combined with changes in farm cropping since the appearance of SLWF and with the use of insect growth regulators, has resulted in a successful integrated approach. By incorporating several compatible technologies, advances in Bemisia management have been made.
Acknowledgments Administrators, scientific colleagues, farmers, their workers, and often village youngsters contributed in the friendliest way to the foreign explorations. Colleagues in the United States took up the natural enemies collected, worked with them, added to our knowledge of them, and supported the objectives of areawide biological control-based integrated management of SLWF. We thank them all.
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53. Lacey, L.A., Mesquita, A.L.M., Mercadier, G., Debire, R., Kazmer, DJ., and Leclant, R (1997). Acute and sublethal activity of the entomopathogenic fungus, Paecilomyces fumosoroseus (Deuteromycotina: Hyphomycetes) on adult Aphelinus asychis (Hymenoptera: Aphelinidae). Environ. Entomol 26, 1452-1460. 54. Poprawski, T.J., Legaspi, J.C, and Parker RE. (1998). Influence of entomopathogenic fungi on Serangium parcesetosum (Coleoptera: Coccinellidae), an important predator of whiteflies (Homoptera: Aleyrodidae). Environ. Entomol. 11^ 785-795. 55. Hoelmer, K.A., and Culver, G. (1997). Survey of desert host plants for whiteflies and parasitoids. In "Silverleaf Whitefly: 1997 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, T.M. Perring, and R.M. Faust, eds.). USDA-ARS Publ. No. 1997-01. 56. McAuslane, H.J., Johnson, F.R., and Knauft D.A. (1994). Population levels and parasitism of Bemisia tabaci (Gennadius) (Homoptera: Aleyrodidae) on peanut cultivars. Environ. Entomol. 23,1203-1210. 57. Schuster, D.J., Evans, G.A., Bennet, F.D., Stansly, PA., Jansson, R.K., Leibee, G.L., and Webb, S.E. (1998). A Survey of parasitoids of Bemisia spp. whiteflies in Florida, the Caribbean, and Central and South America. Int. J. Pest Management 44, 255-260. 58. Simmons, A.M. (1998). Survey of the parasitoids of Bemisia argentifolii (Homoptera: Aleyrodidae) in coastal South Carolina using yellow sticky traps. J. Entomol. Sci. 33, 7-14. 59. Riley, D.G., and Ciomperlik, M.A. (1997). Regional population dynamics of whitefly (Homoptera: Aleyrodidae) and associated parasitoids (Hymenoptera: Aphelinidae). Environ. Entomol. 26, 1049-1055. 60. Hunter, M.S., Antolin, M.F., and Rose, M. (1996). Courtship behavior, reproductive relationships, and allozyme patterns of three North American populations of Eretmocerus nr. californicus (Hymenoptera: Aphelinidae) parasitizing the whitefly Bemisia spp., tabaci complex (Homoptera: Aleyrodidae). Proc. Entomol. Soc. Wash. 98, 126-137. 61. Rose, M., and Zolnerowich, G. (1997). Eretmocerus Haldeman (Hymenoptera: Aphelinidae) in the United States, with descriptions of new species attacking Bemisia {tabaci complex) (Homoptera: Aleyrodidae). Proc. Entomol. Soc. Wash. 99, 1-27. 62. Legaspi, J.C, Legaspi, B.C., Jr., Carruthers, R.I., Goolsby, J.A., Jones, W.A., Kirk, A.A., Moomaw, C , Poprawski, T.J., Ruiz, R.A., Talekar, N.S., and Vacek, D. (1996). Foreign exploration for natural enemies of Bemisia tabaci from Southeast Asia. Subtrop. Plant Sci. 48,48-53. 63. Goolsby, J.A., Ciomperlik, M.A., Legaspi, B.C., Jr., Legaspi, J.C, and Wendel, L.E. (1998). Laboratory and field evaluation of exotic parasitoids of Bemisia tabaci (Biotype "B") in the Lower Rio Grande valley of Texas. Biol. Control 12, 127-135. 64. U.S. Department of Agriculture, Animal and Plant Health Inspection Service. (1995). "Field Releases of Nonindigenous Parasitic Wasps in the Genus Eretmocerus and Encarsia (Hymenoptera: Aphelinidae) for Biological Control of Whitefly Pests (Homoptera: Aleyrodidae). Environmental Assessment." USDA-APHIS, Riverdale, MD. 65. Hunter, M.S., Rose, M., and Polazek, A. (1996). Divergent host relationships of males and females in the parasitoid Encarsia porteri (Hymenoptera: Aphelinidae). Ann. Entomol. Soc. Am. 89,667-675. 66. Black, W C IV, DuTeau, N.M., Puterka, G.J., Nechols, J.R., and Pettorini, J.N. (1992). Use of the random amplified polymorphic DNA polymerase chain reaction (RAPD-PCR) to detect DNA polymorphisms in aphids (Homoptera: Aphididae). Bull. Entomol. Res. 82, 151-159. 67. Vacek, D.C, Ruiz, R.A., and Wendel, L.E. (1996). RAPD-PCR identification of natural enemies of SLWF. In "Silverieaf Whitefly: 1996 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C Toscano, R.M. Faust, and J.R. Coppedge, eds.). USDA-ARS Publ. 1996-02. 68. Nguyen, R., and Bennett, F.D. (1995). Importation and field release of parasites against silverleaf whitefly, Bemisia argentifolii (Bellows & Perring) in Florida from 1990-1994. Proc. Fla. State Hort. Soc. 108, 4 3 ^ 7 .
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69. Headrick D.H., Bellows, T.S., Perring, T.M., and Orr, B. (1997). Natural enemy releases against the silverleaf whitefly in the Imperial Valley. In "Silverleaf Whitefly: 1997 Supplement to the 5Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, R.M. Faust, and J.R. Coppedge, eds.). USDA-ARS Publ. 1997-02. 70. Hoelmer, K.A. (1998). Comparative field cage evaluations of top-performing introduced parasitoids in desert cantaloupes. In "Silverleaf Whitefly: 1998 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, T.M. Perring, and R.M. Faust, eds.). USDA-ARS Publ. 1998-01. 71. Ciomperlik, M.A., Goolsby, J.A., Poprawski, T.J., Wendel, L.E., and Wraight, S. (1997). Demonstration of biological control-based IPM of Bemisia in the lower Rio Grande valley of Texas. In "Silverleaf Whitefly: 1997 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, T.M. Perring, and R.M. Faust, eds.). USDA-ARS Publ. 1997-01. 72. Roltsch, W.J., and Goolsby, J.A. (1998). Field cage evaluation of nonindigenous parasitoiods in desert crops. In "Silverleaf Whitefly: 1998 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, T.M. Perring, and R.M. Faust, eds.). USDA-ARS Publ. 1998-01. 73. Hoelmer, K.A. (1995). Whitefly parasitoids: Can they control field populations of Bemisia? In "Bemisia 1995: Taxonomy, Biology, Damage Control and Management" (D. Gerling and R. Mayer, eds.), pp. 451-476. Intercept Ltd., Andover, Hampshire, U.K. 74. Simmons, G.S., Hoelmer, K.A., Staten, R.S., Boratynski, T., and Natwick, E. (1998). Biological control of silverleaf whitefly infesting cantaloupe with large-scale releases of exotic parasitoids in the Imperial Valley of California. In "Silverleaf Whitefly: 1998 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, T.M. Perring, and R.M. Faust, eds.). USDA-ARS Publ. 1998-01. 75. Goolsby, J.A., and Ciomperlik, M.A. (1999). Development of Parasitoid Inoculated Seedling Transplants for Augmentative Biological Control of Silverleaf Whitefly (Homoptera: Aleyrodidae). Florida Entomol. 82, in press. 76. Pickett, C.H., Simmons, G.S., Goolsby J.A., and Overholt, D. (1999). Fall releases of parasites into citrus by. In "Silverleaf Whitefly: 1999 Supplement to the 5-year National Research and Action Plan" (TJ. Henneberry, N.C. Toscano, TM. Perring, and R.M. Faust, eds.). USDA-ARS Publ. 1999-01. 77. Hoelmer, K.A., and Kirk, A.A. (1999). An overview of natural enemy explorations and evaluations for Bemisia in the U.S. lOBCAVPRS Bull. 22, 109-112. 78. Gould, J., Waldner, D., Colleto, N., Antilla, L., and Santangelo, R. (1998). Release of exotic parasitoids for establishment in Arizona. In "Silverleaf Whitefly: 1998 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, T.M. Perring, and R.M. Faust, eds.). USDA-ARS Publ. 1998-01. 79. Heilmann, L. (1997). Development of species specific DNA probes for Eretmocerus species. In "Silverleaf Whitefly: National Research, Action and Technology Transfer Plan: 1997 Supplement to the 5-Year National Research and Action Plan" (T.J. Henneberry, N.C. Toscano, R.M. Faust, and J.R. Coppedge, eds.). USDA-ARS Publ. 1997-02. 80. Rose, M., and Zolnerowich, G. (1998). Eretmocerus Haldeman (Hymenoptera: Aphelinidae) imported and released in the United States for control of Bemisia (tabaci complex) (Homoptera: Aleyrodidae). Proc. Entomol. Soc. Wash. 100, 31-323. 81. Moomaw, C. (1995). Survey of the indigenous parasitoids of Bemisia tabaci in the lower Rio Grande valley of Texas. M.S. Thesis, Dept. of Entomology. Texas A&M University, College Station, TX. 82. Legaspi, B.C., Jr., Legaspi, J.C, Carruthers, R.I., Goolsby, J.A., Hadman, J., Jones, W.A.D., Murden, D., and Wendel, L.E. (1997). Areawide population dynamics of silverleaf whitefly (Homoptera: Aleyrodidae) and its parasitoids in the lower Rio Grande valley of Texas. J. Entomol. Sci. 32, 445^59.
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CHAPTER 17
Interference with Ultraviolet Vision of Insects to Impede Insect Pests and Insect-Borne Plant Viruses YEHEZKEL ANTIGNUS MOSHE L A P I D O T SHLOMO COHEN
/, Introduction Chemical control of pests in agriculture was the method of choice for a long time [1]. However, this approach is ephemeral, and its negative effects on the environment, such as toxicity and destruction of the natural balance, are well known and documented. During the last decade, people's awareness of the drawbacks of chemical control has increased. Integrated pest management (IPM) is a better alternative for controlling various pests, including insect-borne plant viruses. It is considered to be a system that, in the socioeconomic context of farming systems and the associated environmental and populational dynamics of pest species, utilizes all suitable techniques in a compatible manner to maintain pest populations at levels below those causing economic loss [2]. The hazards of chemical control to the environment and to human health have increased emphasis on cultural control methods in IPM systems. Many cultural procedures used for virus control are aimed at eradicating or altering one or more of the primary participants in the transmission process (vector, virus-source plants, and susceptible crops) or preventing them from coming together [3, 4]. Some of these control strategies are intended to maximally reduce the number of inoculative insects in the field or to interfere with the transmission process during any of its phases, thereby reducing the spread of virus in the field. Luring insects away from crops with colored mulches as described by Cohen [5] or repelling them with reflective surfaces are sometimes powerful tools for protecting field crops from virus epidemics [6, 7]. However, cultural controls for protecting Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
indoor crops are relatively limited and mainly consist of mechanical barriers such as 50-mesh screens [8, 9]. This chapter describes some aspects of ultraviolet (UV) vision in insects and the use of this knowledge to create an innovative approach to control. This new approach, already implemented in Israeli agriculture, alters UV vision to prevent insects from invading greenhouses. Moreover, by altering the normal behavioral paradigm of insect vectors by changing their UV vision, one can reduce plant virus transmission.
//. Structure and Function of ttie Insect Compound Eye Light is the major means by which insects communicate with their environment and orient themselves to hosts. The characteristics dictating various behavioral reactions of insects to light are intensity, spectral composition, polarization, and other physical attributes of light. Insects perceive light signals via their compound eyes. Located on the sides of the head, the eyes are connected to the visual centers of the brain, where light-induced electrochemical impulses are processed and translated, usually to motor responses. A.
Morphology of the Compound Eye
The structural unit of the compound eye is the ommatidium. The number of ommatidia varies from less than 20 to over 30,000, depending on the species. The dioptric apparatus is located on the distal part of the ommatidium and consists of a corneal lens and underlying crystalline cone. The retinula, or photosensitive part of the ommatidium, consists of seven or eight elongated pigmented or retinula cells located beneath the crystalline cone. The pigmented cells connect the optic lobe of the protocerebrum via nerve fibers originating from their proximal end. The retinula cells are arranged around the rhabdom of the ommatidium and form a central axis and site of photoreception [10]. Some retinula cells can serve as UV receptors. Sensitivity to UV is determined by a special UV visual pigment, UV rhodopsin. At least two spectral classes of retinula cells occur in flies, pure UV receptors and UV receptors that are also sensitive to wavelengths in the visible spectrum [11]. Diurnal or day-active insects have apposition compound eyes. In apposition vision there is no sharing of light among adjacent ommatidia. The crystalline cone and retinula of each ommatidium are in close contact. Additionally, both the comeagenous cells surrounding the crystalline cone and the retinula cells surrounding the rhabdom of each ommatidium are densely and evenly pigmented. These anatomical features of the apposition eye prevent light that enters the diop-
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trie apparatus of one ommatidium from escaping that ommatidium and secondarily stimulating any adjacent ommatidia. Since there is no sharing of light, each ommatidium forms its own individual elemental image. Sensory information gathered by ommatidia is transmitted for analysis to the paired optic lobes, one on each side of the protocerebrum of the supraesophageal ganglion or brain [12]. B.
Light Signals and Insect Responses to Light
Insects are strongly dependent on color vision. True color vision has been defined as the ability of an animal to distinguish spectral colors on the basis of wavelength differences, independent of brightness. On the other hand, wavelength-specific behavior occurs when different behavioral patterns of an animal result from markedly different spectral sensitivities. This implies that photoreceptors with different spectral sensitivities can trigger different behavioral reactions without being integrated into the central nervous system [13]. From a biological point of view, the spectral-specific responses of insects are affected by three main parameters that characterize light: (1) hue, determined by the dominant wavelength (Knax) remitted by a surface; (2) color saturation or purity of hue, which represents the distance (in percent) between white (0% purity) and any spectral color (100% purity); and (3) brightness or light intensity, which represents overall reflection. Intensity affects response when a particular response is associated with the peak intensity of a dominant wavelength [14, 15]. Color vision is either dichromatic (as in dragonflies and beetles) or trichromatic (as in bees). Behavioral and corresponding electrophysiological experiments have shown that trichromatic vision, analogous to human vision, is mediated by three types of photoreceptors (wavelength presented parenthetically): UV (320 nm), blue (440 nm), and green (540 nm) [11]. The colors insects can see will depend, therefore, on the selective absorption of radiation from the entire visible-light spectrum, including UV (invisible "black light" to humans). Some insects can distinguish flowers with the same color in the visible spectrum by analyzing differences in their UV reflection [16,17]. Differences in UV reflection and absorption by different parts of the flower serve as visual cues helping bees to reach pollen and nectar [11]. Differences in the reflection of UV from female and male wings occur in many diurnal butterflies. Sometimes these unique patterns of UV reflection serve as sex markers, assisting communication between the sexes. Nocturnal light is rich in UV, and the fact that the wings of many nocturnal moths reflect UV facilitates the sexes getting together [16]. C.
Light Signals and Orientation
Any light beam consists of a mixture of linearly polarized waves oscillating in all possible transversal planes. There are ways to selectively separate lightwaves that are oscillating in the same plane or direction, yielding a beam of polarized light.
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In contrast to the human eye, insects possess a special polarization analyzer, which responds to changes in the plane of polarized light. Natural light is polarized (up to 70 or 80%) in different directions. The state of light polarization changes during the day with corresponding changes in the position of the sun. Both compound eyes and ocelli (simple eyes) are sensitive to polarized light. The wavelength peaks of polarization sensitivities vary for different insects: 433-435 nm for the field cricket Gryllus campestris and 455 nm for the housefly Musca domestica. Honey bees, Aphis mellifera, do not react to polarization across the whole visible spectrum but only to the shorter-wavelength, high-frequency portion of the light spectrum, particularly UV wavelengths in the range of 345 to 350 nm [18]. The sensitivity of the compound eye to polarized light allows insects to orient themselves to their surroundings and distinguish objects. Most insects have a strong attraction to polarized UV light. The total energy of UV light needed to attract insects is always significantly lower than that needed in other wavelengths. The high attractiveness of reflected UV radiation remains unexplained [16].
///.
Ultraviolet-Dependent, Vision-Related Behavior
As discussed earlier, the UV component of the light spectrum plays an important role in the ecological behavior of insects, including orientation, navigation, feeding, and interaction between the sexes. Understanding how these behavior types are affected by the light spectrum in economically important insects is prerequisite to designing innovative, light-based control strategies. A.
Effect of Ultraviolet on Whiteny Flight Behavior
Mound [19] suggested that Bemisia tabaci is attracted by two wavelength groupings of transmitted light, the blue-UV and yellow portions of the spectrum. He concluded that B. tabaci is attracted to either yellow or UV, but not to both at the same time. He indicated that sensitivity to shortwave radiation provides a strong stimulus for individuals to fly toward the sky on a sunny day [19]. Attraction to UV induces migratory behavior, whereas yellow radiation induces vegetative behavior and possibly assists in host selection. No olfactory reactions have been noted for B. tabaci. Moericke [20] demonstrated that Aleyrodes brassicae is able to distinguish between UV absorbing white and an identical white surface that reflects UV Vaishampayan et al [14] observed a strong positive response of the greenhouse whitefly, Trialeurodes vaporariorum (Westwood), to surfaces with maximum reflectance or transmittance in the yellow-green region (520-610 nm) and a moderately positive response to UV between 360 and 380 nm. Light in the blue-violet region seemed to inhibit the response, and red (610-700 nm) might also be moderately inhibitory. The first steps in host selection (orientation and
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landing) are mediated in the greenhouse whitefly mainly, if not exclusively, by a response to reflected yellow light in the 520 to 610 nm range [21]. Coombe [22] found that the greenhouse whitefly took off more readily and walked faster under 400 nm light than under 500 nm light. He confirmed the suggestions of Mound [19] that the two types of radiation are complementary and involved in the balance between UV-induced migratory behavior and yellowinduced alighting (landing) behavior. B.
Effect of UV on Aphid Flight Behavior
Take off and flight activities of aphids are increased in the presence of shortwave light. Aphids taking off from a plant are strongly attracted to ultraviolet light. After flying for varying periods of time, aphids enter the alighting or searching phase, during which they are repelled by the shortwave light of the sky and attracted to long-wave light reflected from plants [23]. Whether landing aphids are repelled or not by shortwave light is under debate. White surfaces that reflect shortwave light or aluminum sheets that reflect most of the light spectrum, including U y inhibit aphid landing. The latter is confirmed by the correlation between UV reflection and repellence of aphids from reflective surfaces [23, 24]. C. Effect of UV on Thrips Flight Behavior Based on the behavioral response of the western flower thrips, Frankliniella occidentalis, to UV and to blue and yellow traps, Vernon and Gillespie [25] suggested that the flower thrips has a photosystem similar to bees, with three photoreceptors tuned to 350 to 360 nm in the UV, 440 to 450 nm in the blue, and 540 to 570 nm in the yellow wavelengths. However, based on results obtained from electroretinograms ofE occidentalis exposed to flashes of light ranging from 365 to 620 nm, one peak of efficiency occurs at 365 nm and a second in the green-yellow region at about 540 nm. The latter suggests that the flower thrips has only two types of photoreceptors, one sensitive to UV and another to green-yellow wavelengths. There is no physiological evidence for a third photopigment sensitive to blue wavelengths. Thrips use peak sensitivity to green-yellow in long-range orientation to plants and to flowers, thus discriminating among U y green-yellow, and blue spectral hues [26]. Reflectance of UV wavelength (350-390 nm) is an important inducer of thrips landing on a host. If UV reflectance is very high, anthophilous (flower-feeding) thrips are repelled from a surface of otherwise attractive colors, whereas grassfeeding thrips are not affected [26]. There is a significant increase in thrips capture when a filter that eliminates wavelengths shorter than 370 nm is placed over a highly reflective UV lead-white trap. These results indicate that a highly reflective UV surface, or more specifically the spectrum between 350 and 370 nm is repellent [15]. In contrast to these findings, Costa and Robb [27] have demon-
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YEHEZKEL ANTIGNUS , MOSHE LAPIDOT, AND SHLOMO COHEN
strated in choice experiments that thrips are attracted to UV-containing light. A possible explanation for this discrepancy is the role of UV intensity in the reaction of thrips to this part of the spectrum.
IV
Ultraviolet' Vision-Based Management Strategies
Manipulation of the visual behavior of insects has been used successfully in the past in IPM programs aimed at protecting crops from insects and insect-borne virus diseases. Control measures include the use of colored and reflective plastic mulches and spraying with whitewashes [3,4]. The possibility of insect control by manipulating UV vision was discovered only recently [28,29]. This approach and probable mechanisms that may explain it are described below. A.
Light Traps for Nocturnal Insects
The concept of using light traps to eradicate harmful insects is an old one. The limits of light traps in the past were associated with the inefficiency of kerosene and acetylene light sources. Significant progress was achieved with the introduction of electrical power to rural areas. Most light traps consist of a mercury lamp seated on a mechanical, aerodynamic, or electrical killing apparatus. This type of trap can attract insects from distances of a few meters to several hundred meters, depending on the power of the lamp. In the United States, traps consist of a fluorescent lamp emitting "black light." The light source is a low-pressure mercury tube supplied with a filter transparent to the near ultraviolet at 365 nm. This type of trapping proved useful in combating insects living in hidden places [16, 30]. Light traps are sometimes used in Israel to protect greenhouse-grown roses from moths. B.
Use of Ultraviolet-Absorbing Films to Protect Greenhouse Crops
Worldwide, growing of crops under plastic has steadily increased during the last decade, with nearly 1 million acres now under plastic in the Mediterranean area [31]. Plastic-covered greenhouses physically isolate and protect plants while providing an environment in which temperature and humidity as well as light intensity and quality can be optimally controlled. Greenhouse photoselective (UV-absorbing) cladding materials were found useful not only for horticultural aspects but also for combating fungal diseases [32, 33]. The first evidence for UV-absorbing films reducing insect invasion of greenhouse came from Japan. Nakagaki et al [29, 34] reported fewer Aphis gossypii and Trialeurodes vaporariorum whiteflies on tomatoes grown in a plastic house treated to exclude UV light than on plants grown in an ordinary plastic house.
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400
600 Wavelength (nm)
Wavelength (nm)
Fig, 1 Light transmission spectra of standard and UV-absorbing plastic sheets. Four different plastics were tested: (A) IR, a regular non-UV-absorbing polyethylene (Ginegar Plastic Products, Israel); (B) Solarig, a UV-absorbing polyethylene (Palrig, Neot Mordechai, Israel); (C) IR-Veradim, a UVabsorbing polyethylene (Ginegar Plastic Products, Israel); (D) Rav-Hozek, a UV-absorbing PVC (Erez, Thermoplastic Products, Israel).
Thrips {E occidentalis and Scirtothrips dorsalis) also occurred in lower numbers under UV-absorbing plastic, as did the leafminer Liriomyza bryoniae. In a series of experiments carried out in Israel during the last 5 years, it was shown that photoselective greenhouse cladding materials can efficiently filter the UV spectrum from light, thus reducing insect numbers and spread of insect-borne plant viruses [28]. Spectrally modified polyethylene is produced commercially by adding UVabsorbing chemicals to the raw material. Polyethylene is the most popular greenhouse cladding material because of its relative low price. Polyvinylchloride (PVC) and polycarbonate also have UV-absorbing properties. All the aforementioned materials block light in the UV range (200^00 nm) but not that in the visible range (400-700 nm) (Fig. 1) [28]. The protective activity of the UV-absorbing films was tested by monitoring and comparing the invasion rates of insect pests into experimental tunnels and greenhouses covered with conventional polyethylene films versus UV-absorbing films [28]. In small (6 x 6 m) experimental "walk-in" tunnels, the number of whiteflies trapped on sticky yellow plates under a UV-absorbing film was one-fourth to one-tenth that of the number recorded under regular sheeting (Fig. 2). In another experiment carried out in the same structures, the number of ^. gossypii aphids recorded under regular films was about 100 times as high as the number recorded under UV-absorbing sheets (Fig 3). In the same experiment, the UV-absorbing films produced a tenfold reduction in invasion of E
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YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
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occidentalis thrips (Fig. 4) [28]. Trials conducted with green herbs such as basil, chives, mint, cherville, and sage grown in commercial walk-in tunnels (5 X 50 m) yielded similar results. Only one-tenth as many B. tabaci adults were found in Sage tunnels covered with UV-absorbing films as were found in ones covered with regular film [35]. In tunnels where chives were cultivated under UV-absorbing sheets, E occidentalis thrips numbered 0 to 10 per trap. Contrastingly, in tunnels covered with ordinary film, thrips populations reached levels that warranted spraying with chemicals. In mint tunnels covered with conventional sheets, the population of Liriomyza trifolli leafminers varied from 75 to 220 per trap, whereas under the UV-excluding film, the numbers of leafminers per trap were between 10 and 45 (Fig. 5). At the time of harvesting, the crop grown under ordinary film was not marketable owing to the severity of leafminer damage, whereas the mint crop grown under UV filtration was not affected [35]. In another set of experiments conducted in the same structures, UV-absorbing films dramatically reduced the infestation of mint with the nocturnal moths Spodoptera lituralis and Laphygma sp. These moths infested and destroyed mint grown under conventional polyethylene films [36]. Reducing pest populations in these commercial walk-in tunnels also reduced the need for chemical pesticides. The number of insecticide applications under the UV-absorbing sheets was reduced by 50 to 80% (Table I) [35]. Similar protective effects against whiteflies
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YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
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Table I Effect of Standard and UV-Absorbing Polyethylene Cladding on Insecticide Usage Insecticide usage" Herb*
IR standard sheeting
504 UV-absorbing sheeting
Sage Sage Sage Mint Chives Chervil Basil Basil Basil
Confidor Biobeat Evisek Vertimek Marshal Vertimek Vertimek Vertimek Biobeat
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" Insecticides were applied according to the buildup of insect populations inside tunnels. * Herbs were grown in commercial walk-in tunnels.
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(D CO
Days After Planting Fig, 6 Tomato yellow leaf curl virus (TYLCV) disease incidence in tomato plants grown in commercial walk-in tunnels covered with either standard (IR) or UV-absorbing (IR-V) polyethylene sheets. Means + SEM differ significantly at P < .05 when analyzed by one-way analysis of variance.
and thrips were obtained in commercial walk-in tunnels in which the cut flower lisianthus (Eustoma grandiflora) was grown [37]. The protective effect of UV-absorbing films is not limited to insect pests but is also highly effective in reducing the spread of insect-borne plant viruses. An extraordinary reduction in spread of whitefly-borne viruses was recorded in tomatoes, cucumbers, and melons grown in walk-in tunnels and greenhouses covered with UV-absorbing films [28, 38, 39]. Of tomatoes grown in walk-in tunnels covered with a UV-absorbing film, without any insecticide application, 1% were infected with tomato yellow leaf curl virus (TYLCV), compared with 80% of control plants grown under regular film (Fig. 6) [38]. Ultraviolet-absorbing sheets proved even more effective in reducing the spread of TYLCV in tomato than in protecting the plants from whitefly invasion. The latter suggests that the film somehow reduces the whitefly's virus transmission efficiency. The incidence of cucurbit yellowing stunt disorder virus (CYSDV) disease in cucumbers was only one-seventh of that for plants grown under UV-absorbing film versus regular film [39]. Preliminary observations also indicate the film's effectiveness in reducing the spread of nonpersistent, aphid-borne zucchini yellow mosaic virus (ZYMV). The efficiency of protection is clearly dependent on the capacity of the plastic to absorb UV, thus PVC sheets proved superior to polyethylene films in this
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YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
to
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90
100
110
120
130
Days after planting Fig, 7 Trapping of leafminers on sticky yellow traps in commercial greenhouses covered with either standard (IR) or UV-absorbing (IR-V) polyethylene sheets.
respect [28, 38]. There also is a positive correlation between the length of a tunnel and its protective capacity: the longer the tunnel, the lower the proportion of unfiltered sunlight entering the structure and the greater the protection. The geometry of a plastic house is another important determinant of the protective efficiency of UV filtration. Walk-in tunnels allow maximal UV filtration, since the plastic film covers most of the outer surface of the structure except for the nonsheeted, screen entrance to provide ventilation. However, most vegetables are grown in greenhouses with vertical sidewalls. The sidewalls are not covered during daytime and allow unfiltered light to enter. In a series of experiments conducted in commercial greenhouses, it was confirmed that filtration of light by the UV-absorbing plastic roof cover is sufficient to inhibit the invasion of insects such as whiteflies, thrips, and leafminers (Fig. 7) [40]. In these same experiments, it was shown that TYLCV disease incidence in greenhouses covered with UV-absorbing film was only one-fifth of that in control greenhouses covered with conventional film. To obtain adequate protection, the use of a UV-absorbing roof should always be combined with 50-mesh screening on vertical sidewalls. A combination of a UV-absorbing roof and 30-mesh screening on greenhouse sidewalls failed to provide protection against whitefly invasion and the spread of TYLCV [40]. The implementation of UV-blocking cladding materials in the greenhouse industry depends not only on its contribution to pest management but also on the
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absence of any negative effects either on plants [41] or beneficial insects (e.g., pollinators and natural enemies) that are artificially introduced into greenhouses [42]. No significant differences were found in grovs1:h, yield, maturation time, or fresh or dry weight of plant parts in tomatoes grown in greenhouses under conventional and UV-absorbing plastics. Moreover, physiological disorders were reduced by 38% under the UV-blocking plastic compared to the standard structure [43]. In further tests, pepper and cucumber yield and quality were not affected by filtration of UV [44]. Similar results were obtained when the above-mentioned parameters were tested in Israel [40, 47]. However, UV elimination was harmful to a certain variety of lisianthus owing to the violet pigmentation of the flower of this cultivar [37]. Bumblebees are part of greenhouse technology [45], and like other insects, they are exposed to the effects of UV elimination. When the activities of bumblebees under conventional and UV-blocking films are compared in experimental minigreenhouses, a delay in the startup of the hive is noticed under UV-absorbing sheets [46]. However, no significant differences in this and other activities of bumblebees, such as foraging, occur in similar comparisons conducted in standard-size greenhouses [47]. In addition, preliminary results indicate normal activity of the parasitic wasp Aphidius colemani in a UV-deficient environment (Antignus, unpublished). C. Ultraviolet-Absorbing, Insect-Proof Screening Insect-proof screens are used in Israel routinely in tomato crops to protect them from whiteflies and spread of TYLCV Physical barriers formed by 50-mesh screens represent a major element of IPM, contributing to a reduction in the use of poisonous pesticides in indoor-grown vegetables, flowers, and fresh herbs [48]. These screens alone cannot seal the greenhouse completely, not even from infiltration by large insects such as whiteflies, leafminers, and aphids. To improve these protective limitations, it is suggested that nets with a double insect exclusion mechanism be used, based on both their physical and optical properties. Fifty-mesh screens with absorbency properties in the UVA and UVB range, "bionet screens," were developed and found to have an improved blocking effect against a wide range of insects. Bionets are significantly better than conventional 50-mesh screens in protecting tomatoes from infestation with B. tabaci (Fig. 8), the spread of TYLCV (Figs. 9 and 10), red spider mites, and leafininers (L. trifolii), as well as in protecting cucumbers from A. gossypii aphids (Fig. 11). However, 50mesh bionet screens failed to prevent invasion and buildup of the western flower thrips, E occidentalis [49]. Ultraviolet-absorbing screens with a lower density (16and 30-mesh) were not superior to conventional screens of the same mesh size. (A positive correlation was found between the density of the screen and its UV filtration capacity.) None of the bionet screens of 16- and 30-mesh densities was able to reduce the transmission of UV light below the putative critical level required for
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YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
600 -/T
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100 -.
80 J 60 J
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Days after planting Fig, 9 Tomato yellow leaf curl virus (TYLCV) disease incidence of tomato plants grown in walk-in tunnels covered with either standard or UV-absorbing (bionet) 50-mesh screens. Means ± SEM with different letters differ significantly at P < .05 when analyzed by the Student r-test.
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Fig, 10 Tomato plants grown in walk-in tunnels under UV-absorbing 50-mesh bionets (A) show a remarkable decline in infection by tomato yellow leaf curl virus (TYLCV) in comparison with plants grown under standard 50-mesh nets (B).
interference with the insect's visual behavior. The protection mechanism of these nets is a mechanical one and not sufficient to provide practical protection [49]. D.
Putative Mechanisms for the Protective Effects of Ultraviolet Filtration
The infestation of greenhouses with insect pests is associated with the immigration patterns of their populations in the open environment. In earlier sections of this chapter we discussed the adaptation of the insect eye to sense short-wavelength light and the importance of this part of the spectrum as a determinant of temporal and spatial patterns of insect movement and dispersal. The rhythm of activity of insects during the day is dictated by a physiological clock, which is synchronized with the motion of the sun. As mentioned earlier, the state of light polarization during the day depends on the position of the sun in the sky [16]. Changes in insect flight activity during the day may result from their analyzing and responding to changes in UV light polarization. Whitefly flight activity, for instance, is limited to the early hours of the day between 8 and 10 A.M. [50]. Ultraviolet light stimulates aphids and whiteflies to take off and fly toward the sky [19, 23]. Ultraviolet-absorbing cladding material prevents the reflection of UV from greenhouses covered with these materials. Under these circum-
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YEHEZKEL ANTIGNUS, MOSHE LAPIDOT, AND SHLOMO COHEN
200 (Q
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Days after planting Fig. 11 Comparison of standard 50-mesh and UV-absorbing (bionet) screens for their effectiveness in protecting cucumber crops grown in walk-in tunnels from infestation with aphids. Random sampling of 10 leaves from each tunnel monitored the population size of aphids. Means ± SEM with different letters differ significantly at P < .05 when analyzed by the Student ^test.
Stances, the "UV compass" of immigrating insects is directing them to fly to UV-reflecting zones away from the UV-absorbing plastic houses. This hypothetical model of the protective mechanism is supported by the significantly lower numbers of whiteflies that are trapped over greenhouse walls covered with UVabsorbing films or UV-absorbing nets [51] as compared with standard nonabsorbing films and nets. The sky and sun are the main sources of UV by day, whereas the sky and moonlight are the major sources at night. Some nocturnal moths react to the UV in moonlight in the same manner as do diurnal insects to that in sunlight [16]. The low moth infestation rates for crops under UV-absorbing films supports this [35]. The reduction in whitefly population density found under the UV-absorbing plastics cannot by itself explain the dramatic reduction of TYLCV infection [28,40]. It is suggested that the UV-deficient environment created under UV-absorbing films reduces insect activity, diminishing the efficiency of viruliferous insects to transmit virus. Indeed, the reports of Coombe [22] on the high activity of whiteflies under UV and our data on the reduced flight activity of aphids and whiteflies in a UV-deficient environment support this hypothesis [51]. However, the mechanism of pest reduction under UV-blocking plastics is mainly associated with the reduced rates of insects entering these plastic houses [27]. Once insects penetrate the UV-deficient environment, their reproduction and development are normal [28; Antignus et al, unpub-
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lished]. In summary, a twofold mechanism is suggested for the anti-insect and antiviral activity of UV-absorbing films: (1) Ultraviolet absorbancy by greenhouse cladding materials alters the flight orientation of insect pests toward UV-reflecting sites away from the greenhouses. As a consequence, fewer insects invade UV-nonreflective structures. (2) The elimination of UV from light transmitted through UVabsorbing films alters the normal behavior of invading insects, reducing flight activity, feeding behavior, or both. Under these conditions, the virus transmission efficiency of the vector is reduced.
VC Concluding Remarks A greenhouse represents a significantly modified environment compared with the natural habitat. Factors such as temperature, humidity, light spectrum, light intensity, light diffusion, and air movement are all-important, not only for plant development but also as determinants of insect population dynamics. Both the attraction of insects to greenhouse crops and the flux of insect pests and virus diseases within greenhouses is affected by an array of factors not yet studied. Any change in light quality or intensity in a greenhouse associated with a UV-absorbing cladding material may affect the crop (e.g., interference with flower pigmentation) as well as beneficial insects (e.g., pollinators and natural enemies) in the greenhouse. Despite our positive experience with the implementation of UVblocking materials for the management of insect pests in greenhouse crops, many questions regarding the roles of factors such as those cited above must be answered before this procedure is generally adopted for every greenhouse crop. Technological ability to design films with specific spectral properties opens the way to a wider use of optics to interfere with vision-related behavior of insects. However, a better understanding of the visual cues operating between insects and plants is required. In the near future, greenhouse plastic films with spectral properties specifically designed for particular crops, climates, pests, and pathogens will likely be available.
References 1. Perring, T.M., Gruenhagen, N.M., and Farrar, C.A. (1999). Management of plant viral diseases through chemical control of insect vectors. Annu Rev. Entomol. 44,457-481. 2. Dent, R.D. (1995). Introduction. In "Integrated Pest Management" (D. Dent, ed.), pp. 1-8. Chapman and Hall, London. 3. Harrison, B.D. (1984). Progress and problems in the control of arthropod-, nematode-, and seedborne plant viruses. In "Control of Virus Diseases" (E. Kurstak, ed.), pp. 265-299. Marcel Dekker, New York.
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4. Antignus, Y. (1999). Cultural control of insect transmitted viruses. In "Current Trends in Epidemiology and Virus Control in Horticultural Crops" (I.M.A. Gomez, ed.), pp. 79-89. Fundacion para la Investigacion Agraria en la Provincia de Almeria. 5. Cohen, S. (1982). Control of whitefly vectors of viruses by color mulches. In "Pathogen, Vectors and Plant Diseases, Approaches to Control" (K.F. Harris and K. Maramorosch, eds.), pp. 45-56. Academic Press, New York. 6. Harpaz, I. (1982). Nonpesticidal control of vector-borne viruses. In "Pathogens, Vectors and Plant Diseases, Approaches to Control" (K.E Harris and K. Maramorosch, eds.), pp. 1-21. Academic Press, New York. 7. Raccah, B. (1986). Nonpersistent viruses: Epidemiology and control. In "Advances in Virus Research" (K. Maramorosch, F.A. Murphy, and A.J. Shatkin, eds.), pp. 387-429. Academic Press, New York. 8. Berlinger, M.J., Mordecchi, S., and Leeper, A. (1991). Application of screens to prevent whitefly penetration into greenhouses in the Mediterranean Basin. lOBC/WPRS Bull. 14(5), 105-110. 9. Cohen, S., and Berlinger, M.J. (1986). Transmission and cultural control of whitefly-bome viruses. Agr. Ecosyst. Environ. 17, 89-97. 10. Goldsmith, T.H., and Bernard, G.D. (1974). The visual system of insects. In "The Physiology of Insecta" (M. Rockstein, ed.), pp. 166-263. Academic Press, New York and London. 11. Stark, W.S., and Tan, K.E.W.P (1982). Ultraviolet light: Photosensitivity and other effects on the visual system. Photochem. Photobiol. 36, 371-380. 12. Mazokhin-Porshnykov, G.A. (1969). Structure of faceted eyes and visual centers. In "Insect Vision" (T.H. Goldsmith, ed.), pp. 1-19. Plenum Press, New York. 13. Coombe, P.E. (1981). Wave length specific behavior of the whitefly Trialeurodes vaporariorum (Homoptera: Aleyrodidae). J. Comp. Physiol. 144, 83-90. 14. Vaishampayan, S.M., Kogan, M., Waldbauer, G.P., and Wooley, J.T. (1975). Spectral specific responses in the visual behavior of the greenhouse whitefly, Trialeurodes vaporariorum (Homoptera: Aleurodidae). Entomol. Exp. Appl. 18, 344-356. 15. Terry, L.I. (1997). Host selection, communication and reproductive behavior. In "Thrips as Crop Pests" (T. Lewis, ed.), pp. 65-118. CAB International, Oxford, New York. 16. Mazokhin-Porshnykov, G.A. (1969). Light as mean of recognition and orientation. In "Insect Vision" (T.H. Goldsmith, ed.), pp. 213-252. Plenum Press, New York. 17. White, R.H., Stevenson, R.D., Bennet, R.R., Cutler, D.E., and Haber, WA. (1994). Wavelength discrimination and role of ultraviolet vision in the feeding behavior of hawkmoths. Biotropica, 26, 427-435. 18. Seliger, H.H., Lall, A.B., and Biggley, WH. (1994). Blue through UV polarization sensitivities in insects. Optimizations for the range of atmospheric polarization conditions. J. Comp. Physiol. 175, 475-486. 19. Mound, L.A. (1962). Studies on the olfaction and color sensitivity of Bemisia tahaci (Genn.) (Homoptera, Aleurodidae). Entomol. Exp. Appl. 5, 99-104. 20. Moericke, V. (1955). Neue Untersuchungen iiber das Farbensehen der Homopteren. Proc. Conf. Potato Diseases 2nd Meeting, Lisse-Wageningen, Netherlands, 1954, 55-69. 21. Vaishampayan, S.M., Waldbauer, G.P., and Kogan, M. (1975). Visual and olfactory responses in orientation to plants by the greenhouse whitefly, Trialeurodes vaporariorum (Homoptera: Aleurodidae). Entomol. Exp. Appl. 18, 412^22. 22. Coombe, P.E. (1982). Visual behavior of the greenhouse whitefly, Trialeurodes vaporariorum. Physiol. Entomol. 7, 243-251. 23. Kring, J.B. (1972). Flight behavior of aphids. Annu. Rev. Entomol. 17, A6\-A91.
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24. Kring, J.B. (1970). Response of aphids to color and light. From theory to practical application. Frontiers Plant Sci. 23, 6-7. 25. Vernon, R.S., and Gillespie, D.R. (1990). Spectral responsiveness of FrankUniella occidentalis (Thysanoptera: Thripidae) determined by trap catches in greenhouses. Environ. Entomol. 19, 1229-1241. 26. Matteson, N., Terry, I., Ascoli, C.A., and Gilbert, C. (1992). Spectral efficiency of the western flower thrips, FrankUniella occidentalis. J. Insect Physiol. 38, 453-459. 27. Costa, H.S., and Robb, K.L. (1999). Effects of ultraviolet-absorbing greenhouse plastic films on flight behaviour of Bemisia argentifolii (Homoptera: Aleyrodidae) and FrankUniella occidentalis (Thysanoptera: Thripidae). J. Econ. Entomol. 92, 557-562. 28. Antignus, Y., Mor, N., Ben-Joseph, R., Lapidot, M., and Cohen S. (1996). UV-absorbing plastic sheets protect crops from insect pests and from virus diseases vectored by insects. Environ. Entomol. 25, 919-924. 29. Nakagaki, S., Sekiguchi, K., and Onuma, K. (1982). The growth of vegetable crops and establishment of insect and mite pests in a plastic greenhouse treated to exclude near UV radiation. (2) EstabHshment of insect and mite pests. Bull. Ibaraki-Ken Hort. Exp. Sta. 10, 39-47 (in Japanese). 30. Glick, RA., and Hollingsworth, J.R (1954). Response of the pink bollworm moth to certain ultraviolet and visible radiation. J. Econ. Entomol. 47, 81-86. 31. Castilla, N., and Jarret, R (1995). Protected cultivation in the Mediterranean area. Plasticulture 107, 13-20. 32. Elad, Y. (1997). Effect of solar light on the production of conidia by field isolates of Botrytis cinerea and on several diseases of greenhouse-grown vegetables. Crop Prot. 16, 635-642. 33. Reuveni, R. (1997). Control of downy mildew in greenhouse-grown cucumbers using blue photoselective polyethylene sheets. Plant Dis. 81, 999-1004. 34. Nakagaki, S., Amagai, H., and Onuma, K. (1984). The growth of vegetable crops and establishment of insect and mite pests in a plastic greenhouse treated to exclude near UV radiation. (4) Establishment of insect pest on tomatoes. Bull. Ibaraki-Ken Hort. Exp. Sta. 12, 89-94 (in Japanese). 35. Antignus, Y, Lapidot, M., Hadar, D., MessikaY, and Cohen, S. (1997). The use of UV-absorbing plastic sheets to protect crops against insects and spread of virus diseases. In "CIR\ Proceedings, International Congress for Plastics in Agriculture" (S. Ben-Yehosua, ed.), pp. 23-33. Laser Pages Publishing Ltd., Jerusalem. 36. Messika, Y, Antignus, Y, Lapidot, M., Ben Yakir, D., Chen, M., and Zimmerman, C. (1999). Effects of UV-absorbing polyethylene films on spice crops infestation with insect pests and on insecticide application regime under these conditions. Gan Sadeh Vameshek 9, 53-55 (in Hebrew). 37. Messika, Y, Nishri, Y, Gokkes, M., Lapidot, M., and Antignus, Y (1998). UV-absorbing films and Aluminet screens—an efficient control mean to block the spread of insect and viral pests in Lisianthus. Dapey Meida, The Flower Grower Magazine 13, 55-57 (in Hebrew). 38. Antignus, Y, Cohen, S., Mor, N., Messika, Y, and Lapidot, M. (1996). The effects of UV-blocking greenhouse covers on insects and insect-borne virus diseases. Plasticulture 112, 15-20. 39. Mizrahi, S., Sacs, Y, Mor, N., Elad, Y, Reuveni, R., and Antignus, Y (1998). Comparative study on the protection effects of commercial polyethylene films with different absorption spectra against insect, fungal and viral pests. Gan Sadeh Vameshek 5, 33-37 (in Hebrew). 40. Antignus, Y, Lachman, O., Leshem, Y, Matan, E., Yehezkel, H., and Messika, Y (1999). Protection efficiency of UV-absorbing films in greenhouses with vertical walls. In "Summary of Research Projects and Field Experiments in Tomato Crops for 1999." Bull, of Israeli Extension Service, pp. 29-39.
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41. Jansen, M.A.K., Gaba, V, and Greenberg, B.M. (1998). Higher plants and UV-B radiation: Balancing damage, repair, and acclimation. Trends Plant Sci. 3, 131-135. 42. Van Lenteren, J.C. (1995). Integrated pest management in protected crops. In "Integrated Pest Management" (D. Dent, ed.), pp. 311-343. Chapman and Hall, London. 43. Amagai, H., Onuma, K., and Nakagaki, S. (1984). The growth of vegetable crops and establishment of insect and mite pests in a plastic greenhouse treated to exclude near UV radiation. (3) Growth of tomatoes. Bull. Ibaraki-Ken Hort. Exp. Sta. 12, 81-88 (in Japanese). 44. Onuma, K., and Nakagaki, S. (1982). The growth of vegetable crops and establishment of insect and mite pests in a plastic greenhouse treated to exclude near UV radiation. (1) The growth of pepper and cucumber. Bull Ibaraki-Ken Hort. Exp. Sta. 10, 31-38 (in Japanese). 45. Pressman, E., Shaked R., Rosenfeld, K., and Hefetz, A. (1999). A comparative study of the efficiency of bumble bees and electric bee in pollinating unheated greenhouse tomatoes. J. Hort. Sci. Biotech. 74, 101-104. 46. Steinberg, S. Prag, H., Gouldman, D., Antignus, Y., Pressman, E., Asenheim, D., Moreno, Y, and Schnitzer, M. (1997). The effect of ultraviolet-absorbing plastic sheets on pollination of greenhouse tomatoes by bumblebees. In "Proceedings of the International Congress for Plastics in Agriculture (CIPA), Israel. 47. Seker, I. (1999). Studies on the effects of UV-absorbing films on the pollination activity of bumble bees in greenhouse tomatoes. In "Summary of Research Projects and Field Experiments in Tomato Crops for 1999." Bull Israeli Extension Service, pp. 41-53. 48. Ausher, R. (1997). Implementation of integrated pest management in Israel. Phytoparasitica 25, 119-141. 49. Antignus, Y, Lapidot, M., Hadar, D., Messika Y, and Cohen, S. (1998). UV absorbing screens serve as optical barriers to protect vegetable crops from virus diseases and insect pests. J. Econ. Entomol. 91, 1401-1405. 50. Cohen, S., and Melamed-Madjar, V (1978). Prevention by soil mulching of the spread of tomato yellow leaf curl virus transmitted by Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae) in Israel. Bull. Entomol. Res. 68, 465-470. 51. Antignus, Y, Nestel, D., Cohen, S., and Lapidot, M. (2001). Ultraviolet-deficient greenhouse environment affects whiteflies attraction and flight-behavior. Environ. Entomol. 30, 394-399.
CHAPTER 18
Bionomics of Micrutalis malleifera Fowler and Its Transmission of Pseudo-Curly Top Virus JAMES H . TSAI
/. Introduction Diseases caused by geminiviruses are of major economic importance in tropical and subtropical regions of the world [1]. A disease of tomato, Lycopersicon esculentum Mill, resembling beet curly top disease, was first reported to occur in South Florida in the 1950s [2, 3]. Simons and Coe [4] were first to distinguish this disease from that caused by leafhopper-bome beet curly top geminivirus on the basis of the disease agent's transmission by a treehopper, Micrutalis malleifera FOWIQT. Hence the disease was named pseudo-curly top. The identity of the pseudo-curly top geminivirus (PCTV) was not confirmed until the late 1980s [5, 6]. Tomato pseudo-curly top only occurs on the east and west coasts of south Florida and has caused sporadic disease on tomato for the last five decades. Although in certain years the incidence of PCTV could be over 50% [7], the epidemiology and geographical distribution of PCTV and its relationship with other geminiviruses have not been studied for lack of a simple and accurate diagnostic test.
//. Biology of Pseudo-Curiy Top Disease A.
Disease Symptoms and Host Range
In general, PCTV symptoms are very similar to those caused by beet curly top virus. Young plants infected by PCTV are severely stunted because of shortened Virus-Insect-Plant Interactions Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. ISBN 0-12-327681-0
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Fig. 1 Pseudo-curly top symptoms in nightshade, Solarium nigrum L., showing chlorosis of the leaf edge, leaf curling, and shoot proliferation.
intemodes and reduced leaf size. In contrast, plants infected late in their growth may only show symptoms on some branches or parts of branches. Common symptoms include vein clearing, chlorosis of the leaf edge, and leaf curling and cupping, as well as apical shoot proliferation (Figs. 1 and 2). Infected plants are often stunted and rarely set fruit. Symptoms on nightshade. Solarium nigrum L., a common weed host, include vein clearing and chlorosis followed by downward and upward leaf curlings [8]. The limited host range of PCTV includes tomato, nightshade, jimsonweed (Datura stramonium L.), tobacco (Nicotiana glutinosa L), chickweed (Stellaria medea L.), eggplant (Solanum melangena L.), lettuce {Lactuca sativa L.), and ragweed {Ambrosia sp.) [6,9, 10]. The host range of beet curly top geminivirus (BCTV) is much wider than that of PCTV, including more than 300 species in 44 plant families [11]. Undoubtedly, some host species overlap with the host range of PCTV, but it has been shown repeatedly that sugar beet. Beta vulparis L., is not a host of PCTV [6, 12]. B.
Characteristics of Pseudo-Curly Top Virus
Pseudo-curly top virus is considered a geminivirus, based on the formation of telltale nuclear inclusions in plant cells, serological analysis, particle morphology, and genomic sequence data. In both leaf and flower tissues of tobacco plants inoculated with PCTV and stained with azure-A, nuclear inclusions known to contain geminivirus-like particles
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Fig, 2 Pseudo-curly top symptoms in tomato, Lycopersicon esculentum Mill., showing chlorosis of the leaf edge and downward leaf curling.
andring-shapedfibrillar bodies are observable by light microscopy. The same is true of ultrathin sections examined by transmission electron microscopy (TEM). These characteristic inclusions are also evident in plant tissues infected with other geminiviruses [5]. Virions of PCTV have been detected by double-antibody sandwich enzyme-linked immunosorbent assay (DAS-ELISA) in lettuce, eg^lant, nightshade, and tomato, as well as in the treehopper vector with use of a polyclonal antiserum prepared to partially purified PCTV A relationship between PCTV and BCTV is demonstratable by indirect ELISA assays. The DNA nature of PCTV's genome was confirmed by the digestibility of nucleic acid from partially purified PCTV by DNase I but not RNase A. The geminate morphology of its virion is observable by TEM [6]. DNA sequence analysis of PCTV indicates that the genome has some features in common with the leafhopper-bome subgroup I and whiteflyborne subgroup III geminiviruses [13]. Studies also indicate that the coat protein of PCTV although distinct from that of other geminiviruses, exhibits features more akin to the leafhopper-bome than the whitefly-bome geminiviruses [13].
UL Vector Biology A.
Life History
To date, PCTV is the only plant virus reported to be transmitted by a treehopper. Simons [10] conducted the initial study of the life history of M malleifera and
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found that the breeding hosts were Hmited to Solanaceae, including eggplant, nightshade, and ground cherry {Physalis spp.). Based on greenhouse studies, he reported that the nymphs underwent only four molts to reach adulthood. The average durations for stadia 1 through 4 on eggplant were 6, 4, 5, and 8 days, respectively. Both sexes required 38 days for development of immature stages. However, when this treehopper was studied by Tsai [14; unpublished data] at 25 ± 1 °C in growth chambers using eggplant, nightshade, and ground cherry {R floridana Rydb.) as rearing hosts, all nymphs underwent five molts. The average durations of stadia 1 through 5 were 4.6 ± 1.2, 4.3 ± 1.6, 4.4 ± 1.1, 5.4 ± 1.2, and 7.8 ± 0.7 days, respectively, on eggplant; 4.8 ± 1.0, 4.5 ± 0.8,4.6 ± 1.0, 5.2 ± 0.8, and 7.6 ± 0.9 days, respectively, on nightshade; and 4.6 ± 0.2,4.3 ± 0.1,4.4 ± 0.2, 5.4 ± 0.2, and 7.8 ±0.1 days, respectively, on ground cherry [14; Tsai, unpublished data]. The average adult longevities for female and male were, respectively, 56.1 and 37.2 days on eggplant, 33.7 and 30.6 days on nightshade, and 12.0 and 10.6 days on ground cherry. Eggs were deposited mainly on terminal stems and petioles but were often found on eggplant leaves. Eggs are imbedded under the epidermis with a portion of egg exposed at the surface. At 25 ± 1°C, the average egg incubation period was 13.6 days (range, 12-16 days). The mean preoviposition period was 3.6 days. The oviposition period was 51.3 days on eggplant. The average number of eggs per day per female was 2.04. The average lifetime number of eggs laid per female was 55 [14; Tsai, unpublished data]. Both nymphs and adults are very docile and can be handled with ease for rearing and experimental purposes. B.
Morphology 1.
EGG
The mean length of an egg is 0.9 ± 0.02 mm (range, 0.7-1.0 mm), and its mean width is 0.3 ± 0.0 mm (range, 0.2-0.3 mm). It is white and translucent, with one end blunt and the other end pointed sideways. 2.
FIRST INSTAR
The body of the first instar (Fig. 3) has a tubular shape, a mean length of 1.3 ± 0.02 mm (range, 1.0-1.7 mm), and a width of 0.4 ± 0.01 mm (range, 0.05-0.5 mm). It is creamy white in color. The dorsum bears a pair of well developed spines projectingft-omthe vertex of the head. One pair of spines projects posteriorly fi-om the dorsum of each thoracic segment and from the dorsum of each of the abdominal segments 3, 4, 5, 6, 7, and 8. a. Head The head has a triangular clypeus, which is densely covered with setae. The eyes are prominent, project laterally, and are pigmented black. The vertex bears one pair of well developed spines, and the rostrum projects caudally
18.
MiCRUTALIS MALLEIFERA
355
FOWLER
V^^ * - - t ^
>'^-, Fig, 3
Lateral view of first instar nymph of the treehopper Micrutalis malleifera Fowler.
between prothoracic legs to the metacoxa. The basal three segments of the antenna are enlarged and its terminal segments are filamentous. b. Thorax, The dorsa of the prothorax, mesothorax, and metathorax each bear one pair of posteriorly projecting spines. The thoracic legs are well developed and sparsely hirsute. c. Abdomen. The dorsa of the abdominal segments 1 and 2 are unarmed. Segments 3 through 8 are each armed with a pair of posteriorly projecting dorsal spines, one on each side of the median line. Venter of abdomen is slightly concave. Last abdominal segment is setaceous with a dorsal pair of posteriorly projecting spines. 3.
SECOND INSTAR
The outline and shape of the second instar (Fig. 4) are similar to those of the first instar except in size and its pale green color. Its mean length is 1.7 ± 0.02 mm (range, 1.5-2.1 mm) and its mean width is 0.6 ± 0.01 mm (range, 0.6-0.7 mm). It has paired dorsal spines on its head, thorax, and abdomen, and the spines bear a few secondary lateral setae. a. Head and Thorax, The head is similar to that of the first instar in outline and shape. There are dense setae on the clypeus, frons, and vertex. The thorax has
356
JAMES H . TSAI
Fig. 4
Lateral view of second instar nymph of Micrutalis malleifera Fowler.
pairs of dorsal spines with secondary setae on the basal portions. All legs are similar in form and size and covered with dense setae. b. Abdomen. The dorsa of abdominal segments 1 and 2 are unarmed. The dorsa of segments 3 throught 8 have posteriorly projecting spines; the spines have secondary setae on the basal half. The last abdominal segment is setaceous, with a dorsal pair of posteriorly projecting spines. 4.
THIRD INSTAR
The outline and shape of the third instar (Fig. 5) are similar to those of the second instar, except for the enlargement of the thoracic lateral plates, which have developed into wing buds. The mean length is 2.3 ± 0.02 mm (range, 1.9-2.5 mm), and the mean width is 0.8 ± 0.02 mm (range, 0.5-1.0 mm). The color is bright green to pale green. There are paired dorsal spines with brown tips on the head, thorax, and abdomen. The dorsal spines bear long secondary setae. The venter of the abdomen is slightly concave. a. Head and Thorax, The clypeus, frons, and eyes are covered with setae. The thoracic segments are enlarged, each with a pair of dorsal spines projecting posteriorly. Wing buds are apparent on the meso- and metathoracic segments. The mesothoracic wing buds cover three-fourths of the metathoracic wing buds, and
18.
MiCRUTALIS MALLEIFERA
FOWLER
357
Fig, 5 Lateral view of third instar nymph of Micrutalis malleifera Fowler.
the metathoracic wing buds cover the first abdominal segment. The prothoracic sclerite does not protrude. b. Abdomen. Abdominal segments 3 through 8 each have a dorsomedial pair of posteriorly projecting spines. The venter of the abdominal segments is slightly concave. The last abdominal segment is setaceous, with a dorsal pair of posteriorly projecting spines. All spines bear long secondary setae. 5.
FOURTH INSTAR
The outline and shape of the fourth instar (Fig. 6) are similar to those of the third instar except in size. The mean length and width are 3.1 ± 0.02 mm (range, 2.9-3.7 mm) and 1.2 ± 0.02 mm (range, 0.9 ± 1.5 mm), respectively. The body is green to pale green in color. The mesothoracic wing buds extend to the front edge of the third abdominal segment. The metathoracic wing buds are well developed and covered by the mesothoracic wing buds. Paired dorsal spines with black tips are prominent. The dorsal spines bear long secondary setae. The venter of the abdominal segments is slightly concave. a. Head. The head has setae on the clypeus, frons, and eyes. Paired dorsal spines on the vertex are smaller than the thoracic and abdominal dorsal spines.
358
JAMES H . TSAI
Fig. 6 Lateral view of fourth instar nymph oiMicrutalis malleifera Fowler.
b. Thorax. The thorax has a prothoracic dorsal sclerite extending caudally. The mesothoracic wing buds cover the metathoracic wing buds and extend to the front edge of the third abdominal segment. c. Abdomen. The dorsa of segments 3 through 8 each bear a pair of prominent posteriorly projecting spines. The spines bear distinct secondary lateral setae. The venter of the abdomen is slightly concave. The last abdominal segment is tubular and black and bears a pair of posteriorly projecting spines. 6.
FIFTH INSTAR
The outline and shape of the fifth instar (Fig. 7) are similar to those of the fourth instar except in size. Mean length is 4.2 ± 0.02 mm (range, 3.5-5.5 mm) and mean width is 1.8 ± 0.04 mm (range, 1.5-2.7 mm). The color is mostly green, but some individuals are grayish brown. The body setae are dense. There are paired dorsal spines on the head, thorax, and abdomen with prominent black tips. The dorsal spines have long secondary setae. The venter of the abdomen is moderately concave. a. Head. The clypeus, frons, and eyes are covered with setae. There are paired dorsal spines on the vertex, which are about one-half the length of the abdominal dorsal spines.
18.
MiCRUTALIS MALLEIFERA
FOWLER
359
Fig, 7 Lateral view of fifth instar nymph of Micrutalis malleifera Fowler.
b. Thorax. The thorax has dense setae on three thoracic segments and legs. There is a prothoracic dorsal sclerite extending beyond the paired dorsal spines of the mesothorax. The mesothoracic wing buds cover the metathoracic wing buds and extend caudally onto three-fourths of the third abdominal segment. c. Abdomen. The abdomen is triangular in cross-section and is densely covered with setae of various lengths. The dorsa of segments 3 through 8 each bear a pair of prominent posteriorly projecting spines. The spines bear distinct secondary lateral setae. The last abdominal segment is greatly elongated and cylindrical. The venter of the abdomen is slightly concave. 7.
ADULT
Adults (Fig. 8) of M malleifera possess characteristics typical of the subfamily Smiliinae, with its prothoracic tibia not foliaceous and its metathoracic tarsi similar in size to the prothoracic and mesothoracic tarsi. The head is about twice as broad as it is long. The adult female is 4.6 ± 0.03 mm (range, 4.0-5.0 mm) in length and 2.5 ± 0.01 mm (range, 2.3-2.8 mm) in width. The male is 4.2 ± 0.02 mm (range, 4.0 ± 4.5 mm) long and 2.2 ± 0.01 mm (range, 2.1-2.5 mm) wide. a. Head. The head has a triangular frontal aspect, with compound eyes projecting laterally; the face is reddish brown. The clypeus is beaklike and round
360
JAMES H . TSAI
Fig. 8
Lateral view of adult Micnitalis malleifera Fowler treehopper.
below. Antennae project on each side of the clypeus. Three basal antennal segments are enlarged, and the distal segments are fused into a filament. The rostrum is visible ventrally, fitting in the median cavity between the coxae, and extending beyond the mesocoxae. b. Thorax, The anterior two thirds of the pronotum are black. The pronotum is finely pubescent, extending to the terminal third of the abdomen. The coxae and legs are reddish brown and finely pubescent. The forewings have three prominent longitudinal veins arising near the base and four apical cells. The forewings extend to the abdominal apex. c. Abdomen, The abdomen is compressed, with genitalia occupying the terminal third; it is largely concealed by forewings.
IV.
Virus Transmission
To date, M malleifera is the only known vector of PCTV, which it transmits in a circulative manner. Tests to detect transovarial transmission of PCTV proved negative [Tsai, unpublished data]. However, transmission of PCTV can be achieved by injecting M malleifera with either partially purified virus prepara-
18.
MICRUTALIS MALLEIFERA
FOWLER
361
tion [6] or crude sap [Tsai, unpublished data]. Both nymphs and adults are efficient vectors of PCTV. The average transmission efficiencies for fourth-instar nymphs were 60.1% and 78.5% following 10- and 24-hour acquisition access periods (AAPs), respectively. The average transmission efficiencies for adults were 49.0% and 70.1%) following 10 and 24-hour AAPs, respectively. The latent period, measured as LP50, was estimated at 13.5 hours in fourth-instar nymphs after a 10-hour AAP, the shortest latent period being 10 hours. Pseudo-curly top geminivirus can readily be transmitted by M. malleifera injected with freshly squeezed sap from the petiole or terminal stem of infected nightshade. The transmission rates have ranged from 30 to 59% depending on the virus titer in the sap inoculum [Tsai, unpublished data]. A transmission rate of 51% was reported when M. malleifera was injected with a partially purified preparation of PCTV [6]. The minimum AAP and inoculation period are less than 1 hour, and both sexes transmit PCTV at about the same rate [9]. The retention period of PCTV in M malleifera ranges from 4 to 23 days depending on the longevity of the vector. In most cases, retention periods coincide with test insect longevities [Tsai, unpublished data].
References 1. Duffus, J.E. (1987). Whitefly transmission of plant viruses. In "Current Topics in Vector Research" (K.F. Harris, ed.), Vol. 4, pp. 73-91. Springer-Verlag, New York. 2. Stoner, W.N., and Hogan, W.D. (1950). Viruses affecting vegetable crops in the Everglades area. Fla. Agr. Exp. Sta. Annu. Rep. No. 558, 206. 3. Giddings, N.J., Bennett, C.W., and Harrison, A.L. (1951). A tomato disease resembling curly top. Phytopathology 41, 415-417. 4. Simons, J.N., and Coe, D.M. (1958). Transmission of pseudocurly top virus in Florida by a treehopper. Virology 6 , 4 3 ^ 8 . 5. Christie, R.G., Ko, N.-J., Falk. B.W., Hiebert, E., Lastra, R., Bird, J., and Kim, K.S. (1986). Light microscopy of geminivirus-induced nuclear inclusion bodies. Phytopathology 76, 124-126. 6. McDaniel, L.L., and Tsai, J.H. (1990). Partial characterization and serological analysis of pseudocurly top virus. Plant Dis. 74,17-21. 7. Zitter, T.A., and Tsai, J.H. (1981). Viruses infecting tomato in southern Florida. Plant Dis. 65, 787-791. 8. Tsai, J.H., and Brown, L.G. (1991). Pseudo-curly top of tomato. Plant Pathology Circular 344,2. Fla. Dept. of Agriculture and Consumer Services, Gainesville, FL. 9. Simons, J.N. (1962). The pseudo-curly top disease in south Florida. J. Econ. Entomol. 55, 358-363. 10. Simons, J.N. (1962). Life-history and behavioral studies on Micrutalis maelleifera, a vector of pseudo-curly top virus. J. Econ. Entomol. 55, 363-365. 11. Bock, K.R. (1982). Geminivirus diseases in tropical crops. Plant Dis. 66,266-270. 12. Simons, J.N. (1980). Membracids. In "Vectors of Plant Pathogens" (K.F. Harris, ed.), pp. 93-96. Academic Press, New York.
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13. Briddon, R.B., Bedford, I.D., Tsai, J.H., and Markham, P.G. (1996). Analysis of the nucleotide sequence of the treehopper-transmitted geminivirus, tomato pseudo-curly top virus, suggests a recombinant origin. Virology 219, 387-394. 14. Tsai, J.H. (1989). Biology and ecology of treehopper transmission of a geminivirus. In "Proceedings of the 6th International Conference on Comparative and Applied Virology" Oct. 15-21, Banff, Alberta, Canada. Symp. Abstr. No. W7-3.
Index
transmission factors action mode, 156-159 bridge hypothesis, 156-157 characterization, 152-156 identification, 150-152 P2 properties, 147, 152-155, 159-160 P3 properties, 147, 155-156 regulation, 159-160 circulative transmission, electrical penetration graph analysis, 74-80 acquisition experiments, 76 direct data, 76-77 inoculation experiments, 76 methods, 74-76 phloem role, 74 statistical data processing, 78-80 cucumber mosaic virus transmission noncirculative virus transmission, 91, 95, 112,126 overview, 167, 175-176 structure, 170-172 transmission mechanisms, 168, 173-175 engineered mutant defects, 174-175 nonpersistent, 173-174 spontaneous mutant defects, 174 vectors, 169-170 viral genome, 168 noncirculative virus transmission electrical penetration graph analysis, 87-103, 120-125 acquisition, 89, 91-92, 97-98, 121, 124-125, 128 efficiency, 95-96 egestion, 92-93, 116, 121-125, 127129 elucidation, 91-94, 115-129 inoculability retention, 97-98 insecticide effects, 98-100
Abutilon mosaic virus, whitefly vector transmission in continental Europe, 282 Abutilon yellows virus, whitefly vector transmission, 295, 304-305 Acalymma vittata, Southern bean mosaic virus transmission, 134-135, 138-140 Accessory salivary gland cells, Luteovirus transmission by aphids, 210-212, 223-226, 228 Agropyron mosaic, Rymovirus association, 30-31 Aleurodes tabaci, see Bemisia tabaci Antixenosis, see Resistance to viruses Aphid cuticular receptors, noncirculative virus transmission, 111, 118-119, 125-127 Aphids, see also Aphis gossypii; Myzus persicae; Rhopalosiphon padi barley yellow dwarf virus transmission acquisition experiments, 76 direct data, 76-77 inoculation, 76, 210-211 methods, 74-76 overview, 207-208, 226-228 phloem role, 74 proteins involved in transmission, 213-225 coat protein, 214-215 nonstructural proteins, 214 readthrough protein, 215-227 specificity, 211-212 statistical data processing, 78-80 symbionin interactions, 210, 225-226 virus acquisition, 208-210 cauliflower mosaic virus transmission, 148-160 classification, 148-149 helper dependency, 144, 149-152
363
364 Aphids (continued) mechanisms, 88-90, 115-117 mineral oil effects, 98-100 optimal acquisition time, 89, 97-98 overview, 87-88, 102-103, 121, 128129 preacquisition starvation effect, 88, 96-97 salivation mechanism, 93-94, 114, 116-117,121,123-128 semipersistent transmission, 89, 100-102, 127-129 site of vims retention, 117-120 virus-vector specificity, 94-96 nonpersistent transmission mechanisms, 115-117,120-127 terminology, 112-115 potyvirus transmission, 195-197 bridge hypothesis, 125, 185, 195-197 direct binding hypothesis, 125, 196-197 mechanisms, 195 specificity, 197-198 HC-Pro-dependent specificity, 118, 197-198 transmissibility variation, 197 types, 197 ultraviolet management strategies ultraviolet-absorbing films, 336-338 vision-related flight behavior, 335 Aphis gossypii diseases transmitted, 112 electrical penetration graph analysis of noncirculative virus transmission, 87-103 mechanisms, 88-90 nonpersistent transmission acquisition, 89, 91-92, 97-98, 124-125 efficiency, 95-96 egestion, 92-93, 116, 121, 122-125 elucidation, 91-94, 115-129 inoculability retention, 97-98 insecticide effects, 98-100 mineral oil effects, 98-100 optimal acquisition time, 89, 97-98 preacquisition starvation effect, 88, 96-97 salivation mechanism, 93-94, 114, 116-117,121,123-128 virus-vector specificity, 94-96 overview, 87-88, 102-103, 121, 128- 129 semipersistent transmission, 89, 100-102, 127-129
INDEX
ultraviolet management strategies ultraviolet-absorbing films, 336-338 vision-related flight behavior, 335 Arthropod vectors, see specific species Aspartid acid-alanine-glycine motif, coat proteins in potyvirus transmission, 125, 186-189,196,200
B Barley yellow dwarf virus aphid vector interactions electrical penetration graph analysis, 74-80 acquisition experiments, 76 direct data, 76-77 inoculation experiments, 76 methods, 74-76 phloem role, 74 statistical data processing, 78-80 inoculation, 76, 210-211 overview, 207-208, 226-228 proteins involved in transmission, 213-225 coat protein, 214-215 nonstructural proteins, 214 readthrough protein, 215-227 specificity, 211-212 symbionin interactions, 210, 225-226 virus acquisition, 208-210 genome organization, 207-208 Bean pod mottle virus, leaf-feeding beetle association, 134-135, 138 Bean yellow mosaic virus, noncirculative transmission, 92 Beetles, see Chrysomelidae; Colorado potato beetle; Leaf-feeding beetles Beet pseudo-yellows virus, whitefly vector transmission, 280-281, 295-297 Beet western yellows virus, aphid vector transmission coat protein role, 214-215 nonstructural protein role, 214 protein identification, 213-214 readthrough protein role, 215-227 conserved domain region, 220-225 gene map, 216 readthrough domain variability, 218-220 Begomoviruses, see also Tomato yellow leaf curl virus family characteristics, 2-3 whitefly vector transmission in continental Europe, 282-285
INDEX
Bemisia tabaci characteristics, 247-248, 268-270, 287 control strategies, 309-324 biological control integrated pest management, 262-264 parasitoids, 318-323 pathogenic fungi, 315-318 foreign exploration, 311-315 apparent field parasitism, 312-315 collections, 311-312 in Spain, 314-315 taxonomic identification, 312 in Thailand, 315 insecticide resistance, 264-265 overview, 309-311, 323-324 ultraviolet-based management, 334-343 cucurbit yellow stunting disorder virus transmission, 281-282, 297-298 economic impact, 248-250 lettuce chlorosis virus transmission, 295, 300-301 lettuce infectious yellows virus transmission, 249, 295, 298-300 pest status, 248-250 population dynamics, 252-268 biological traits, 252-254 life history, 253-254 reproduction, 252-253 ecological traits, 254-259 dispersal, 259 intraplant distribution, 254-259 environmental factors, 260-265 crop irrigation, 261 host plant diversity, 260-261 insecticide resistance, 264-265 natural enemies, 262-264 overwintering, 262 plant nutrition, 261 weather, 261 Hfe tables, 265-268 population models, 265-268 sweet potato chlorotic virus transmission, 295, 304 taxonomy, 250-252 tomato chlorosis virus transmission, 295, 302-303 tomato yellow leaf curl virus infection by sexual transmission, 1-22 begomovirus characteristics, 2-3 endosymbiotic chaperonins role, 4-9 circulative transmission, 4-5, 81 GroEL homologue involvement, 5-7,22
365 virus acquisition, 4, 19 virus transmission, 4 geminivirus characteristics, 2-3 GroEL role transmission, 5-7, 22 viral coat protein interactions, 5-7, 22 mechanisms, 12-18 population infection, 15-16 serial transmission among sexual partners, 13-15 sexual transmission, 12-13, 19-20 tomato infection, 16-18 overview, 1-2, 18-22 tomato yellow leaf curl virus characteristics, 3 virus effects on whiteflies, 9-12 fecundity effects, 10-12 longevity effects, 10-12 long-term viral coat protein retention, 9-10 long-term viral DNA retention, 9-10 ultraviolet-dependent, vision-related flight behavior, 334-335 Bigeminivirus, see Abutilon mosaic virus; Cotton leaf cnmiple virus; Honeysuckle yellow vein mosaic virus; Squash leaf curl virus; Tomato yellow leaf curl virus Black currant reversion disease, Closterovirus association, 31-32 Bridge hypothesis aphid transmission factor action, 156-157 potyvirus transmission by aphids, 185, 195-197 Brome streak mosaic disease, Rymovirus association, 31 Bromovirus, see Cowpea chlorotic mottle virus; Tomato aspermy virus Bunyaviridae, see Tomato spotted wilt virus; Tospovirus Bymovirus, number of viruses, 183
Capsid-protein subunit noncirculative virus transmission, 118-120, 124-127 potyvirus transmission, HC-Pro role, 194 Carmovirus vectors, see Leaf-feeding beetles Cauliflower mosaic virus bimodal transmission patterns, 89, 112 gene expression strategies, 145-147 genome organization, 145-147
366 Cauliflower mosaic virus (continued) overview, 143-148, 160-161 transmission by aphids, 148-160 classification, 148-149 helper dependency, 144, 149-152 transmission factors action mode, 156-159 bridge hypothesis, 156-157 characterization, 152-156 identification, 150-152 P2 properties, 147, 152-155, 159-160 P3 properties, 147, 155-156 regulation, 159-160 Caulimovirus, see Cauliflower mosaic virus; Thistle mosaic disease Cherry mottle leaf disease, Clostewvirus association, 31-32 Chrysomelidae, virus transmission mechanisms, 133-140 beetle-plant interactions, 133-134 overview, 133, 140 regurgitant deposition, 135-136 virus acquisition, 133-134 virus-beetle interactions, 134-135 virus-plant interactions, 136-140 gross-wound inoculation technique, 136-137, 140 ribonuclease role, 137-138, 140 unwounded cell infection, 139-140 virus translocation, 138-139 Cibarium, virus retention site, ingestionegestion theory, 118-120 Circulative viral transmission electrical penetration graph analysis, 69-83 barley yellow dwarf virus transmission by Rhopalosiphon padi, 74-80 acquisition experiments, 76 direct data, 76-77 inoculation experiments, 76 methods, 74-76 phloem role, 74 statistical data processing, 78-80 overview, 69-70, 82-83 techniques, 70-74 vector resistant plants, 81-82 tomato yellow leaf curl virus transmission by Bemisia tabaci, 4-5, 81 Citrus chlorotic dwarf disease, whitefly vector transmission in continental Europe, 284-285 Closteroviruses, see also Abutilon yellows virus; Beet pseudo-yellows vims; Lettuce
INDEX
infectious yellows virus; Tomato infectious chlorosis virus whitefly vector transmission in continental Europe, 280-282 Coat protein cucumber mosaic virus structure, 172 genetically engineered resistance strategies to potato leafroll virus antisense strategy, 238-239 coat protein strategy, 236-238 Luteovirus transmission by aphids, 214-215 potyvirus transmission aspartid acid-alanine-glycine motif, 186-189,196,200 N-terminal role, 186-189 structure, 185-186 transmission biology, 184-185 tomato yellow leaf curl virus sexual transmission by Bemisia tabaci GroEL interactions, 7-9, 22 long-term coat protein retention, 9-10 Coccinellidae, see Leaf-feeding beetles Colorado potato beetle, potato leafroll virus resistance study, 239 Comovirus, see Bean pod mottle virus Cotton leaf crumple virus, transmission by Bemisia tabaci, 249 Cowpea chlorotic mottle virus, structure, 170-172 Crinivirus, whitefly vector transmission abutilon yellows virus transmission, 295, 304-305 beet pseudo-yellows virus transmission, 280-281,295-297 in continental Europe cucurbit yellow stunting disorder virus transmission, 281-282 tomato infectious chlorosis crinivirus transmission, 281 cucurbit yellow stunting disorder virus transmission, 281-282, 297-298 diodia vein chlorosis virus transmission, 305 lettuce chlorosis virus transmission, 295, 300-301 lettuce infectious yellows virus transmission, 249,295, 298-300 overview, 293-294, 305-306 sweet potato chlorotic virus transmission, 295, 304 tomato chlorosis virus transmission, 295, 302-303
INDEX
tomato infectious chlorosis virus transmission, 281, 295, 301-302 virus-vector relationships, 294-296 inoculation feeding period, 296 minimum acquisition access feeding period, 296 persistence, 296 transmission efficiency, 294-296 Cucumber mosaic virus noncirculative virus transmission, 91, 95, 112,126 overview, 167, 175-176 structure, 170-172 transmission mechanisms, 168, 173-175 engineered mutant defects, 174-175 nonpersistent, 173-174 spontaneous mutant defects, 174 vectors, 169-170 viral genome, 168 Cucumovirus, see Peanut stunt virus; Tomato aspermy virus Cucurbit yellow stunting disorder virus ultraviolet-absorbing films effects, 341 whitefly vector transmission, 281-282, 297-298 Curculionidae vectors, see Leaf-feeding beetles Cuticula-borne virus transmission, see Noncirculative virus transmission
D Diodia vein chlorosis virus, whitefly vector transmission, 305 Direct binding hypothesis, potyvirus transmission by aphids, 196-197 Double membrane-bound particles, eriophyid mite-associated diseases fig mosaic disease, 31-34 High Plains disease, 30-32, 3 9 ^ 2 overview, 29-30, 4 2 ^ 4 pigeon pea sterility mosaic disease, 31,36 redbud yellow ringspot disease, 31, 34 rose rosette disease, 31, 34-36 Rymovirus caused diseases, 30-31 thistle mosaic disease, 31, 37-39 tomato spotted wilt tospoviruses compared, 42^3 wheat spot chlorosis disease, 31, 33 wheat spot mosaic disease, 30-31, 33 in woody dicots, 32
367 E Egestion, ingestion-egestion theory, 111-129 electrical penetration graph analysis, 120-122 nonpersistent transmission, 92-93, 116 overview, 111-112,116,129 potential drop analysis, 122-125 phases and subphases, 122-123 virus acquisition, 124-125, 128 virus inoculation, 123-124, 128 terminology, 112-115 watery saliva role, 125-127 Electrical penetration graphs circulative virus transmission analysis, 69-83 barley yellow dwarf virus transmission by Rhopalosiphon padi, 74-80 acquisition experiments, 76 direct data, 76-77 inoculation experiments, 76 methods, 74-76 phloem role, 74 statistical data processing, 78-80 overview, 69-70, 82-83 techniques, 70-74 vector resistant plants, 81-82 noncirculative virus transmission analysis, 87-103,111-129 mechanisms, 88-90, 102-103, 115-117 nonpersistent transmission, 90-100, 111-112 acquisition, 89, 91-92, 97-98, 121, 124-125, 128 efficiency, 95-96 egestion, 92-93, 116, 121-125, 127129 elucidation, 91-94, 115-129 inoculability retention, 97-98 insecticide effects, 98-100 mineral oil effects, 98-100 optimal acquisition time, 89, 97-98 preacquisition starvation effect, 88, 96-97 salivation mechanism, 93-94, 114, 116-117,121,123-128 site of virus retention, 117-120 virus-vector specificity, 94-96, 118119 overview, 87-88, 102-103 semipersistent transmission, 89, 100-102 Emanovirus, see also Pea enation mosaic virus-1 genome organization, 207-208
368 Encarsia, white fly parasitism studies apparent field parasitism, 312-315 establishment methods, 322-323 field evaluation, 321-322 prerelease surveys and evaluations, 318-319 quarantine methods, 319-320 quarantine screening, 320-321 Engineered resistance, see Resistance to viruses Eretmocerus, white fly parasitism studies apparent field parasitism, 312-315 establishment methods, 322-323 field evaluation, 321-322 prerelease surveys and evaluations, 318-319 quarantine methods, 319-320 quarantine screening, 320-321 Eriophyid mites associated diseases, 30-42 double membrane-bound particles association, 32-42 fig mosaic disease, 31-34 High Plains disease, 30-32, 3 9 ^ 2 mechanisms, 29-30, 42-44 pigeon pea sterility mosaic disease, 31, 36 redbud yellow ringspot disease, 31,34 rose rosette disease, 31, 34-36 thistle mosaic disease, 31, 37-39 tomato spotted wilt tospoviruses compared, 42-43 wheat spot chlorosis disease, 31, 33 wheat spot mosaic disease, 30-31, 33 Rymovirus caused diseases, 30-31 in woody dicots, 32 overview, 29-30, 4 2 ^ 4 Extravasation, definition, 115
Fig mosaic disease, eriophyid mite association, 31-34 Frankliniella occidentalis Tospovirus transmission, 51-63 kinetics, 54-56 midgut infection, 56-60 overview, 51-52, 63 salivary gland infection, 60-61 vector description, 52-54 viral protein function, 62-63 virus composition, 52 virus morphology, 52 virus replication in thrips, 56
INDEX
ultraviolet management strategies ultraviolet-absorbing films, 337-339 vision-related flight behavior, 335-336 Fungi, white fly control, 315-318 biotic factors, 316-317 compatibility with other natural enemies, 318 field studies, 317-318 fungal factors, 315-316 physical factors, 316
Garlic mosaic disease, Rymovirus association, 31 Geminivirus, see also Maize streak virus; Tomato yellow leaf curl virus family characteristics, 2-3 whitefly vector transmission in continental Europe, 282-285 Genetically engineered resistance, see Resistance to viruses Graminella nigrifrons, semipersistent transmission of maize chlorotic dwarf virus, 101-102 GroEL protein, tomato yellow leaf curl virus transmission by Bemisia tabaci transmission mechanisms, 5-7, 22 viral coat protein interactions, 7-9, 22 Gross-wound inoculation technique, virus transmission mechanisms by leaf-feeding beetles, 136-137, 140
H Helper component protein noncirculative virus transmission bridge hypothesis, 118-119, 125-127, 156-157 cauliflower mosaic virus transmission by aphids, 144, 149-152, 156 dependency, 144, 149-152, 170 mechanisms, 118-120, 124-129 potyvirus transmission characterization, 189-190 functional domains, 190-193 aphid transmission function, 191-193 mutations, 191-194 proteolytic function, 190-191 HC-Pro domain site, 184 HC-Pro role aphid binding domain, 194
INDEX
binding domains, 193-194 bridge hypothesis, 118-119, 125-127, 185, 195-197 capsid-binding domain, 194 direct binding hypothesis, 125, 196-197 interaction models, 199 other functions, 193 requirement for virus retention, 193 structure-function relationship, 193-194 transmission function, 191-193 transmission mechanisms, 118-119, 125-127, 195 virus-aphid specificity, 197-198 purification, 189 transmissibility loss, 189 transmission biology, 185 High Plains disease, eriophyid mite association, 30-32, 3 9 ^ 2 Honeysuckle yellow vein mosaic virus, whitefly vector transmission in continental Europe, 283 Hordeum mosaic disease, Rymovirus association, 31 Hybrigeminivirus, see Tomato pseudo-curly top virus
Ingestion-egestion theory, 111-129 electrical penetration graph analysis, 120-122 ingestion-salivation hypothesis compared, 121,128-129 nonpersistent transmission, 92-93, 111-112, 116 overview, 111-112, 116, 121,128, 129 potential drop analysis, 122-125 phases and subphases, 121-123, 128 virus acquisition, 121, 124-125, 128 vims inoculation, 121, 123-124, 128 terminology, 112-115 watery saliva role, 125-129 Ingestion-salivation hypothesis electrical penetration graph analysis, 120-122 ingestion-egestion theory compared, 121, 128-129 overview, 116-117, 129 potential drop analysis, 122 virus retention site, 117-120
369 cibarium, 118-120 common duct, 117-118 maxillary food canal, 118-120 Insecticides, see also Pest control nonpersistent transmission effects, 98-100 resistance in Bemisia tabaci, 264-265 Insect vectors, see specific species Integrated pest management, see Pest control Ipomoea yellow vein virus, whitefly vector transmission in continental Europe, 283-284 Ipomovirus, number of viruses, 183
Leaf-feeding beetles, virus transmission mechanisms, 133-140 beetle-plant interactions, 133-134 overview, 133, 140 regurgitant deposition, 135-136 virus acquisition, 133-134 virus-beetle interactions, 134-135 virus-plant interactions, 136-140 gross-wound inoculation technique, 136-137, 140 ribonuclease role, 137-138, 140 unwounded cell infection, 139-140 virus translocation, 13 8-139 Leafhoppers, see Graminella nigrifrons Lettuce chlorosis virus, whitefly vector transmission, 295, 300-301 Lettuce infectious yellows virus, whitefly vector transmission, 249,295, 298-300 Light traps, ultraviolet-based pest management strategies, 336 Liriomyza trifolli, ultraviolet management strategies, ultraviolet-absorbing films, 339-440 Luteovirus, see also Barley yellow dwarf virus; Beet western yellows virus; Potato leafroll virus aphid interactions, 207-228 inoculation, 210-211 overview, 207-208, 226-228 proteins involved in transmission, 213-225 coat protein, 214—215 nonstructural proteins, 214 readthrough protein, 215-227 specificity, 211-212 symbionin interactions, 210, 225-226 virus acquisition, 208-210
370 M Machlomovirus vectors, see Leaf-feeding beetles Maize chlorotic dwarf vims direct binding hypothesis support, 196 semipersistent transmission by Graminella nigrifrons, 101-102, 127 Maize streak virus, electrical penetration graph analysis, 81 Maple mosaic disease, whitefly vector transmission in continental Europe, 284-285 Meloidae vectors, see Leaf-feeding beetles Metopolophium dirhodum, virus specificity, 211 Micrutalis malleifera, tomato pseudo-curly top virus transmission, 351-361 disease biology, 351-353 characteristics, 352-353 disease symptoms, 351-352 host range, 351-352 life history, 353-354 morphology, 354-360 egg, 354 first instar, 354-355 second instar, 355-356 third instar, 356-357 fourth instar, 357-358 fifth instar, 358-359 adult, 359-360 overview, 351 transmission mechanisms, 360-361 Mineral oils, nonpersistent transmission effects, 98-100 Mosaic viruses, see specific viruses Myzus persicae cauliflower mosaic virus transmission, 148-149 noncirculative viral transmission analysis electrical penetration graph analysis, 90-91,96-99 mechanisms nonpersistent transmission, 115-116 terminology, 112-115 potato leafroU virus transmission, 233, 236 tobacco vein mottling virus transmission, 197-198
Nicotiana clevelandii, coat protein study, 214-215
INDEX
Noncirculative virus transmission, see also Cauliflower mosaic virus bridge hypothesis, 125, 156-157 electrical penetration graph analysis mechanisms, 88-90, 120-125 nonpersistent transmission acquisition, 89, 91-92, 97-98, 124-125 efficiency, 95-96 egestion, 92-93, 116, 121, 122-125 elucidation, 91-94, 115-129 inoculability retention, 97-98 insecticide effects, 98-100 mineral oil effects, 98-100 optimal acquisition time, 89, 97-98 preacquisition starvation effect, 88, 96-97 salivation mechanism, 93-94, 116-117, 121, 123-128 virus-vector specificity, 94-96 overview, 87-88, 102-103, 121, 128-129 potential drop analysis, 122-125 semipersistent transmission, 89, 100-102, 127-129 ingestion-egestion theory, 111-129 electrical penetration graph analysis, 120-125 nonpersistent transmission, 92-93, 115117 overview, 111-112, 116, 121, 128-129 potential drop analysis, 122-125 phases and subphases, 122-123 virus acquisition, 124-125, 128 virus inoculation, 123-124, 128 terminology, 112-115 watery saliva role, 125-127 ingestion-salivation hypothesis electrical penetration graph analysis, 120-122 overview, 116-117, 129 potential drop analysis, 121-122 virus retention site, 117-120 cibarium, 118-120 common duct, 117-118 maxillary food canal, 118-120 nonpersistent transmission, see Nonpersistent virus transmission semipersistent transmission egestion role, 127-129 electrical penetration graph analysis, 89, 100-102 Nonpersistent virus transmission characteristics, 169
INDEX
cucumber mosaic virus transmission by aphids overview, 167, 175-176 structure, 170-172 transmission mechanisms, 168, 173-175 engineered mutant defects, 174-175 spontaneous mutant defects, 174 working model, 173-174 vectors, 169-170 viral genome, 168 electrical penetration graph analysis acquisition, 89, 91-92, 121-125, 127-129 efficiency, 95-96 egestion, 92-93, 116 elucidation, 91-94, 115-129 inoculability retention, 97-98 insecticide effects, 98-100 mineral oil effects, 98-100 optimal acquisition time, 89, 97-98 preacquisition starvation effect, 88, 96-97 salivation mechanism, 93-94, 116-117, 121, 123-128 virus-vector specificity, 94-96, 118-119 ingestion-egestion role, 92-93, 116 ingestion-salivation role, 93-94, 116-117 mechanical contamination hypotheses, 115-116
O Oat necrotic mottle disease, Rymovirus association, 31 Onion mite-borne latent disease, Rymovirus association, 31 Overwintering, Bemisia tabaci population dynamics, 262
Paecilomyces fumosoroseus, white fly control, 312,315-318 biotic factors, 316-317 compatibility with other natural enemies, 318 field studies, 317-318 fungal factors, 315-316 physical factors, 316 Parabemisia myricae characteristics, 285-287 citrus chlorotic disease transmission, 285 Peach mosaic disease, Closterovirus association, 31-32
371 Pea enation mosaic virus-1, genome organization, 207-208 Peanut stunt virus, genome organization, 168 Pea seed-borne mosaic virus, bimodal transmission patterns, 89 Pest control Bemisia tabaci, 309-324 biological control integrated pest management, 262-264 parasitoids, 318-323 pathogenic fungi, 315-318 foreign exploration, 311-315 apparent field parasitism, 312-315 collections, 311-312 in Spain, 314-315 taxonomic identification, 312 in Thailand, 315 insecticide resistance, 264-265 overview, 309-311, 323-324 ultraviolet-based management, 334-343 ultraviolet vision interference in insects, 331-348 insect eye structure and function, 332-334 compound eye morphology, 332-333 light signalss, 333-334 orientations, 333-334 visual responsess, 333 overview, 331-332, 347 ultraviolet-based management strategies, 336-347 light traps, 336 putative mechanisms, 345-347 ultraviolet-absorbing films, 336-343 ultraviolet-absorbing insect-proof screening, 343-345 ultraviolet-dependent, vision-related behavior, 334-336 aphid flight behavior, 335 thrip flight behavior, 335-336 whitefly flight behavior, 334-335 Pigeon pea sterility mosaic disease, eriophyid mite association, 31, 36 Plum pox virus, nonpersistent transmission, 183 Polerovirus, see also Potato leafroll virus genome organization, 207-208 Potato leafroll virus genetically engineered resistance, 233-242 approaches, 236-240 coat protein antisense strategy, 238-239
372 Potato leafroll virus (continued) coat protein strategy, 236-238 putative movement protein strategy, 239-240 replicase strategy, 239 ribozyme strategy, 240 genome organization, 235-236 overview, 233-235, 242-243 resistance mechanisms, 240-242 genome organization, 207-208 Potato virus Y noncirculative transmission electrical penetration graph analysis, 95, 98-101, 120-125 mechanisms nonpersistent transmission, 115-117, 120-125,183, 185, 189-192 terminology, 112-115 viral resistance mechanisms in transgenic plants, 240 Potyviruses, see also Bean yellow mosaic virus; Pea seed-borne mosaic virus; Plum pox virus; Potato virus Y; Sweet potato chlorotic virus; Tobacco etch virus; Tobacco vein mottling virus; Zucchini yellow mosaic virus associated diseases, 30-31 coat protein aspartid acid-alanine-glycine motif, 125, 186-189, 196,200 N-terminal role, 186-189 structure, 185-186 transmission biology, 184-185 helper proteins characterization, 189-190 functional domains, 190-193 aphid transmission function, 118-119, 125-127, 191-193 HC-Pro mutations, 191-194 proteolytic function, 190-191 purification, 189 transmissibility loss, 189 transmission biology, 185 number of viruses, 183 overview, 181, 198-200 transmission biology, 182-185 coat protein, 184-185 helper proteins, 185 non-aphid-transmissible strains, 184 transmission mode, 182-183 vectors, 182
INDEX
virion deficiency effects, 184 viruses, 183 transmission by aphids, 195-197 bridge hypothesis, 125, 185, 195-197 direct binding hypothesis, 125, 196-197 mechanisms, 195 specificity, 197-198 HC-Pro-dependent specificity, 197-198 transmissibility variation, 197 types, 197 Preacquisition starvation effect, electrical penetration graph analysis of noncirculative viral transmission, 88, 96-97 Pseudo-curly top virus, 351-361 biology, 351-353 characteristics, 352-353 disease symptoms, 351-352 host range, 351-352 overview, 351 vector biology, 353-360 life history, 353-354 morphology, 354-360 egg, 354 first instar, 354-355 second instar, 355-356 third instar, 356-357 fourth instar, 357-358 fifth instar, 358-359 adult, 359-360 virus transmission, 360-361 Putative movement protein, genetically engineered resistance strategy to potato leafroll virus, 239-240
Readthrough protein, Luteovirus transmission by aphids, 215-227 gene map, 216 readthrough domain conserved region, 220-225 variability, 218-220 Redbud yellow ringspot disease, eriophyid mite association, 31, 34 Replicase, genetically engineered resistance strategy to potato leafroll virus, 239 Resistance to insecticides, Bemisia tabaci, 264-265 Resistance to viruses genetically engineered resistance to potato leafroll virus, 233-242
INDEX
approaches, 236-240 coat protein antisense strategy, 238-239 coat protein strategy, 236-238 putative movement protein strategy, 239-240 replicase strategy, 239 ribozyme strategy, 240 genome organization, 235-236 overview, 233-235, 242-243 resistance mechanisms, 240-242 mechanisms, 81-82 Rhopalosiphon maidis, virus specificity, 211 Rhopalosiphon padi, barley yellow dwarf virus transmission electrical penetration graph analysis, 74-80 acquisition experiments, 76 direct data, 76-77 inoculation experiments, 76 methods, 74-76 phloem role, 74 statistical data processing, 78-80 virus specificity, 211 Ribonuclease, virus transmission role in leaffeeding beetles, 137-138, 140 Ribozyme, genetically engineered resistance strategy to potato leafroll virus, 240 RNA viruses, see specific viruses Rose rosette disease, eriophyid mite association, 31, 34-36 Ryegrass mosaic disease, Rymovirus association, 30-31 Rymovirus associated diseases, 30-31 number of viruses, 183
Salivation accessory salivary gland cells, Luteovirus transmission by aphids, 210-212, 223-226, 228 ingestion-salivation hypothesis electrical penetration graph analysis, 120-122 overview, 116-117, 129 potential drop analysis, 121-122 semipersistent transmission, 127-129 virus retention site, 117-120 cibarium, 118-120
373 common duct, 117-118 maxillary food canal, 118-120 watery saliva role, 125-127 Schizaphis graminum, virus specificity, 211 Semipersistent transmission electrical penetration graph analysis, 89, 100-102 ingestion-egestion theory, 127-129 Sexually transmitted viruses, tomato yellow leaf curl virus transmission by Bemisia tabaci, 1-22 begomovirus characteristics, 2-3 endosymbiotic chaperonins role, 4-9 circulative transmission, 4-5, 81 GroEL homologue involvement, 5-7, 22 virus acquisition, 4, 19 virus transmission, 4 geminivirus characteristics, 2-3 GroEL role transmission, 5-7, 22 viral coat protein interactions, 7-9, 22 mechanisms, 12-18 population infection, 15-16 serial transmission among sexual partners, 13-15 sexual transmission, 12-13, 19-20 tomato infection, 16-18 overview, 1-2, 18-22 tomato yellow leaf curl virus characteristics, 3 virus effects on whiteflies, 9-12 fecundity effects, 10-12 longevity effects, 10-12 long-term viral coat protein retention, 9-10 long-term viral DNA retention, 9-10 Shallot mite-borne latent disease, Rymovirus association, 31 Sitobion avenae, virus specificity, 211 Sobemovirus, see Southern bean mosaic virus Southern bean mosaic virus, Acalymma vittata association, 134-135, 138-140 Spartina mottle disease, Rymovirus association, 31 Spodoptera lituralis, ultraviolet management strategies, ultraviolet-absorbing films, 339 Squash leaf curl virus, transmission by Bemisia tabaci, 4, 249 Sweet potato chlorotic virus, whitefly vector transmission, 295, 304
374 Symbionin, Luteovirus transmission by aphids, 210,225-226
Thistle mosaic disease, eriophyid mite association, 31, 37-39 Thrips Tospovirus transmission, 51-63 kinetics, 54-56 midgut infection, 56-60 overview, 51-52, 63 salivary gland infection, 60-61 vector description, 52-54 viral protein function, 62-63 virus composition, 52 virus morphology, 52 virus replication in thrips, 56 ultraviolet management strategies ultraviolet-absorbing films, 337-339 vision-related flight behavior, 335-336 Tobacco etch virus nonpersistent transmission, 183, 186-188 viral resistance mechanisms in transgenic plants, 240 Tobacco mosaic virus, leaf-feeding beetle association, 136, 138-139 Tobacco vein mottling virus, transmission by Myzus persicae, 197-198 Tomato aspermy virus, genome organization, 168 Tomato chlorosis virus, whitefly vector transmission, 295, 302-303 Tomato infectious chlorosis virus, whitefly vector transmission, 281, 295, 301-302 Tomato pseudo-curly top virus, 351-361 biology, 351-353 characteristics, 352-353 disease symptoms, 351-352 host range, 351-352 overview, 351 vector biology, 353-360 life history, 353-354 morphology, 354—360 egg, 354 first instar, 354-355 second instar, 355-356 third instar, 356-357 fourth instar, 357-358 fifth instar, 358-359 adult, 359-360
INDEX
virus transmission, 360-361 Tomato spotted wilt virus double membrane-bound particle diseases compared, 4 2 ^ 3 thrip vector association, 51-63 composition, 52 morphology, 52 overview, 51-52, 63 thrip vectors, 52-53 description, 52-54 kinetics, 54-56 midgut infection, 56-60 salivary gland infection, 60-61 viral protein function, 62-63 virus replication in thrips, 56 Tomato yellow leaf curl virus ultraviolet-absorbing films effects, 341-344 whitefly vector transmission, 1-22 endosymbiotic chaperonins role, 4-9 circulative transmission, 4-5 GroEL homologue involvement, 5-7, 22 vims acquisition, 4, 19 virus transmission, 4 geminivirus characteristics, 2-3 GroEL role transmission, 5-7, 22 viral coat protein interactions, 7-9, 22 mechanisms, 12-18 population infection, 15-16 serial transmission among sexual partners, 13-15 sexual transmission, 12-13, 19-20 tomato infection, 16-18 overview, 1-2, 18-22 transmission in continental Europe, 284 virus characteristics, 3 virus effects on whiteflies, 9-12 fecundity effects, 10-12 longevity effects, 10-12 long-term viral coat protein retention, 9-10 long-term viral DNA retention, 9-10 Tospovirus, see also Tomato spotted wilt virus anatomical perspective of transmission, 51-63 composition, 52 morphology, 52 overview, 51-52, 63 thrip vectors, 52-53 description, 52-54
375
INDEX
kinetics, 54-56 midgut infection, 56-60 salivary gland infection, 60-61 viral protein function, 62-63 virus replication in thrips, 56 Transcription, Caulimovirus gene expression strategies, 145-147 Transgenic virus resistance, see Resistance to viruses Transmission factors, cauliflower mosaic virus transmission by aphids action mode, 156-159 bridge hypothesis, 156-157 characterization, 152-156 identification, 150-152 P2 properties, 147, 152-155, 159-160 P3 properties, 147, 155-156 regulation, 159-160 Trialeurodes vaporariorum abutilon yellows virus transmission, 295, 304-305 beet pseudo-yellows virus transmission, 280-281,295-297 characteristics, 285 maple mosaic disease virus transmission, 285 tomato infectious chlorosis virus transmission, 281, 295, 301-302 ultraviolet management strategies ultraviolet-absorbing films, 336 vision-related flight behavior, 334-335 Tymovirus vectors, see Leaf-feeding beetles
U Ultraviolet pest management, 331-348 insect eye structure and function, 332-334 compound eye morphology, 332-333 light signals, 333-334 orientation, 333-334 visual responses, 333 overview, 331-332, 347 ultraviolet-based management strategies, 336-347 hght traps, 336 putative mechanisms, 345-347 ultraviolet-absorbing films, 336-343 ultraviolet-absorbing insect-proof screening, 343-345 ultraviolet-dependent, vision-related behavior, 334-336
aphid flight behavior, 335 thrip flight behavior, 335-336 whitefly flight behavior, 334-335
Virus resistance, see Resistance to viruses Virus transmission factors, see Transmission factors
W Waikavirus, see Maize chlorotic dwarf virus Weather, Bemisia tabaci population dynamics, 261 Wheat spot chlorosis disease, eriophyid mite association, 31, 33 Wheat spot mosaic disease, eriophyid mite association, 30-31, 33 Wheat streak mosaic, Rymovirus association, 30-31 Whitefly, see also Bemisia tabaci; Trialeurodes vaporariorum ultraviolet-dependent, vision-related flight behavior, 334-335 virus transmission abutilon mosaic virus transmission, 282 abutilon yellows virus transmission, 295, 304-305 beet pseudo-yellows virus transmission, 280-281,295-297 citrus chlorotic dwarf disease transmission, 284-285 cucurbit yellow stunting disorder virus transmission, 281-282, 297-298 diodia vein chlorosis virus transmission, 305 honeysuckle yellow vein mosaic virus transmission, 283 Ipomoea yellow vein virus transmission, 283-284 lettuce chlorosis virus transmission, 295, 300-301 lettuce infectious yellows virus transmission, 249, 295, 298-300 maple mosaic disease transmission, 284-285 overview, 279, 287, 293-294, 305-306 sweet potato chlorotic virus transmission, 295, 304
376 Whitefly (continued) tomato chlorosis virus transmission, 295, 302-303 tomato infectious chlorosis virus transmission, 281, 295, 301-302 tomato yellow leaf curl virus transmission, 284 virus-vector relationships inoculation feeding period, 296 minimum acquisition access feeding period, 296
INDEX
persistence, 296 transmission efficiency, 294-296 vector species, 285-287
Zucchini yellow mosaic virus nonpersistent transmission, 183, 187, 191-192 ultraviolet-absorbing films effects, 341