Global Plant Genetic Resources for Insect-Resistant Crops

GLOBAL PLANT GENETIC RESOURCES FOR INSECT-RESISTANT CROPS

Edited by

Stephen L. Clement and Sharron S. Quisenberry

CRC Press Boca Raton New York

© 1999 by CRC Press LLC

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Library of Congress Cataloging-in-Publication Data McLachlan, Alan Molecular biology of the hepatitis B virus / Alan McLachlan p. cm.

Includes bibliographical references and index.

ISBN 0-8493-2695-8

1. Hepatitis B virus. 2. Biology—molecular. I. McLachlan, Alan. II. Title. [DNLM: 1. Hepatitis B virus. QW 710 G289h]

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Foreword

Masa Iwanaga In spite of the successes of the “Green Revolution”, food security is assuming an importance of ever-increasing dimensions. Most projections estimate a growth in world population over the next quarter century to 8.5 billion, signifying an additional 3 billion people to feed. At the same time, an overall decline is being registered in the per capita surface area available for food production. It is imperative to find sustainable, cost-effective means of augmenting food supplies, primarily through reliable improvement of crop yields, to meet the inevi­ table increased demand for resources. At present, despite all of this century’s improvements in production technology, 800 million people are estimated to be undernourished. The majority of these people reside in developing countries, where almost four-fifths of the world’s population live and where, by the year 2025, it is thought that 97% of the global population will compete for food supplies, with social, environmental, economic, and political repercussions worldwide. Therefore, it is essential that the problem of food security be recognized as a challenge facing all of humanity, and that concerted global efforts in agricultural research be intensified. Both national governments and international bodies must collaborate in increasing productivity on all available agricultural land while employ­ ing environmental safeguards to protect natural resources (i.e., soil, water, plant genetic resources) for the future. Of these natural resources, genetic resources offer us the greatest benefits in terms of return on scientific, technological, and economic inputs and, therefore, require the most focused attention of researchers. The safe conservation and sustainable use of plant genetic resources are the keys to ensuring food security and, by extension, poverty alleviation and the promotion of peaceful development. The prevention of genetic erosion of these resources and, even more importantly, the conservation of genetic diversity in the world’s plant crops, are essential to future food security. Food production can be increased by the extension of arable land. Advances in this direction, however, appear to have reached the limits of our present scientific and technological capabilities. Further expansion of agricultural areas can only be made at unacceptable environmental costs. A second means of increasing food production would be to intensify cultivation and increase yields from lands which are presently agriculturally viable. This was the objective of agricultural policies and research programs in the last few decades, when the development and spread of high-yielding cultivars of major crops resulted in large increases in production. These gains, however, came only at the expense of reduced sustainability. Improved or high-yielding cultivars alleviated some problems but it soon became apparent that they also introduced constraints that created new

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problems. The evolution of plants and insects are closely intertwined, with local adaptation of landraces incorporating, through a lengthy selection process, a natural resistance to impediments to their growth in the local environment. This balance between pest organisms and host plants was disturbed by the spread of monoculture of improved cultivars and the resulting reliance on a series of chemical pesticides. Monoculture of single high-yielding cultivars made crops more vulnerable, in the long run, to the resilience of endemic pests and diseases, which adapted to the expanded and homogenized host environment. This led to the increasing dependence of farmers on the use of chemical means to counter these sources of crop losses. But, over time, pest organisms — with their short reproductive cycles — are able to evolve resistance to pesticides. This leads to an increase in pest infestations, increased use of pesticides in an escalating “treadmill” of action and reaction, and higher production costs and damage to the environment and human health. The problem is often compounded by pesticides inadvertently killing the natural enemies of the target pests. Reduction of food supplies through damage by pathogens, animal pests, and weeds — apart from abiotic factors such as weather fluctuations — is serious, accounting by some estimates to anywhere from 30 to 70% of total production losses, depending on the crop and region. In value terms, total crop losses may approach $490 billion (U.S.). Insects are the major culprits, especially in developing countries where tropical conditions often foster large populations and where, unfortunately, crop losses are most damaging and difficult to compensate. In relative terms, insects are responsible for greater crop damage worldwide than disease pathogens or weeds. Insect pests have been estimated to cause $90.5 billion in damages to eight principal food and cash crops (rice, wheat, barley, maize, potato, soybean, cotton, and coffee) versus $76.9 billion in damage by disease pathogens and $76.3 billion by weeds.1 Apart from direct damage to plants by insects through consumption of plant parts, insects also serve as vectors for viruses which cause a number of plant diseases. Damage occurs not only in the field, but also under post-harvest storage conditions and during use. In addition to expansion of agricultural production areas and intensification of production, a third means of increasing food production tackles the problem of reducing these crop losses. It involves the protection of food yields through means which break or circumvent the “pesticide treadmill” pattern and which seek to restore a sustainable balance. This is accomplished by integrated pest management (IPM), which aims to protect crop yields by containing losses from pest attacks and at the same time lowering economic and environmental risks. It has been suggested that by adopting IPM approaches, global crop yields could be doubled. Approaches include: • introducing the natural enemies (parasites, predators, and diseases) of the pest species; • improved monitoring and predictions of pest populations and movements; • judicious and reduced use of pesticides; • intercropping to avoid monoculture;

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• introduction of pest-resistant cultivars; • insect control through the employment of pheromones, repellents, intro­ duction of sterility, etc.; and • culturally acceptable, cost-effective, and non-toxic methods for protecting stores of harvested crops from damage. Host plant resistance is a basic component of IPM. Plant breeding for pest resistance has been estimated to be responsible for a full third of recent yield increases. Resistant cultivars can be developed through breeding programs and resistance qualities can be conferred in vitro on plants that are subject to damage by insects either in the field or in post-harvest storage. This is the key area where plant genetic resources researchers are equipped to provide a foundation for food security. The number of genebanks housing accessions of plant genetic resources germ­ plasm has increased in the last 20 years from 10 to the more than 1,300 collections that are registered in FAO’s WIEWS (World Information and Early Warning System) database. Approximately 6.1 million accessions are stored worldwide in ex situ germplasm collections, including 527,000 in field genebanks. Data are not available on how many accessions maintained by genebanks have been used in breeding programs or have contributed to improved cultivars. However, there is a consensus that the conserved germplasm has not been fully used and that there is a need in most countries to ensure better use of plant genetic resources (including underutilized species) through plant breeding for more pest-resistant cultivars. Highly promising areas in which genetic resources research is successfully focusing, such as molecular techniques for identifying genes which determine desirable characteristics and in vitro methods for rapid propagation and selection, merit increased emphasis in future IPM programs. In addition, the deployment of productive diversity in crop plants is a cornerstone of the holistic IPM approach, requiring interactive expertise in several fields to bring about increased plant resistance. Agrobiodiversity contributes to reduced crop losses through: • bolstering intraspecific diversity to put in place a broader range of traits to which insect pests must adapt, thus making more difficult the cyclical interplay between effect and response in continuing pest-host adjustments; • introduction and maintenance of interspecific diversity to effect a complex ecosystem where competition between species will reduce rapid develop­ ment of simple single-plant host/single-insect pest relationships through a portfolio of crop species subject to a range of pests; • intercropping to provide barriers to movement and spread of pests; and • use of “time” controls, i.e., different cultivars of a crop can be planted in alternating years to slow down the process of pest adaptation, and plant breeding can result in crop cultivars with traits that allow them to be planted or harvested earlier or later to break synchronization with the pest’s reproductive cycle.

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Numerous obstacles limit the effective use of plant genetic resources. They include lack of characterization and evaluation data, lack of documentation and information, poor coordination of policies at the national level, poor linkages between the genebank and the users of the germplasm, and lack of plant breeding programs. This situation is improving, however, and exciting successful examples of effec­ tive use of genetic diversity for the development of insect-resistant crops abound. This book provides many such examples, from which we can learn valuable lessons for the effective use of conserved germplasm. The widely applicable lesson is that close, effective, and long-term collaboration is required among different players, including germplasm curators, plant breeders, molecular biologists, entomologists, ecologists, and social scientists. As plant yields are increased, however, there is a corresponding rise in their vulnerability to pests and pathogens, making vigilance and collaboration by plant genetic resources scientists and entomologists vital to minimize crop losses. Linkage of efforts at national, regional, and international levels is also critical to deal with globally important pest problems, and thus to increase the world’s food security by maximizing existing resources and production potential.

REFERENCE 1. Oerke, E.C., H.W. Dehne, F. Schonbeck, and A. Weber, Crop Production and Crop Protection: Estimated Losses in Major Food and Cash Crops, Elsevier Science B.V., Amsterdam, 1994.

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Preface

Social and natural scientists, the general public, and policymakers affiliated with government and non-governmental organizations have become more aware of the importance of plant genetic resources to world agriculture and food security over the last four decades of the 20th century. Erosion of biodiversity in agriculture resulting from widespread cropping of high-yielding, genetically uniform cultivars, coupled with the continuous and permanent loss of natural plant biodiversity, brought about by habitat destruction and environmental degradation, has resulted in this heightened interest in germplasm conservation. In light of these developments, the question becomes: Can we sustain human society, albeit one that owes its origins to the emergence of land plants, without the vast genetic diversity that has enabled plants to surmount insect pests, diseases, and droughts over the ages? An ample supply of genetically diverse germplasm is vital to sustain future agricultural pro­ ductivity worldwide. What makes this book unique among a considerable body of literature on the importance and use of genetic resources to world agriculture is the entomological focus of the assembled chapters. The chapters are grouped into six topic groups: foreword, cereal crops, legume crops, vegetable crops, root and tuber crops, and basic research and biotechnology. An important concept that is addressed in several places in the book is the dependency of modern agriculture on chemical pest control. Modern crop cultivars without the pest-resistant genes or alleles that occur in the wild types and related species of crops are often highly susceptible to insect attack and damage, a point emphasized in the Foreword to this volume. These resistance genes frequently have been lost under intense selection for traits such as high yield and improved human nutritional value. If we hold to the widely believed importance of developing agricultural production systems less dependent on chemical pesticides, researchers will have to look to the world’s plant germplasm conservation sites as a major source of pest-resistant genes for crop improvement programs. The message conveyed by each of the crop-based chapters in this book is that germplasm stocks have been indispensable to entomologists, plant breeders, and other agricultural scientists for insect resistance evaluations for crop improvement. Sometimes, fewer than 10 accessions have been screened for insect resistance; however, many evaluations have included hundreds if not thousands of accessions of a particular collection. Insect- and mite-resistant material has been identified (Chapters 1 through 11), although a higher frequency of usable resistance has been found in the germplasm stocks of some crops (see chapters on rice, wheat, barley, sorghum, common bean, soybean, and alfalfa) than in other crops (see the chapters on grain legumes, vegetable crops, potato, and sweetpotato). As pointed out by some authors in this book, the breeding of advanced lines with resistance genes from unadapted germplasm has been constrained by biological, technological, and social

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factors. Overcoming these constraints is contingent on interdisciplinary research teams in national and international programs, i.e., plant breeders, molecular biolo­ gists, entomologists, plant pathologists, and social scientists, working together to apply modern plant breeding and new biotechnological innovations to develop insectresistant cultivars. This book is also about the use and potential of germplasm stocks for basic research in insect-plant interactions, which the penultimate chapter covers, and the applications of biotechnology to the use of natural plant genes for insect-resistant crops, which the final chapter covers. Progress in developing insect-resistant crops relies on indepth knowledge of the myriad biotic factors that mediate insect-plant interactions, and conserved germplasm has much to offer the entomologist and ecologist with an interest in this field of study. Finally, the resource that plant and microbial germplasm bring to plant biotechnology for insect resistance is incalcu­ lable when one considers that scientists have barely scratched the “biodiversity surface” for resistance genes and products. This book is the product of the efforts, cooperation, and understanding of world leaders in the conservation and use of global plant genetic resources for sustainable agricultural production. We are grateful to the authors for their contributions. We also express our indebtedness to colleagues who provided critical reviews of each chapter. Pioneer Hi-Bred International and Zeneca Ag Products provided partial funding for a symposium at the XX International Congress of Entomology, August 1997, on which this book was based. Finally, we owe a special debt of gratitude to Leslie Elberson and Marilyn Weidner for their professional and tireless efforts in assisting with the preparation of this book for publication. Stephen L. Clement Pullman, Washington Sharron S. Quisenberry Lincoln, Nebraska

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Editors

Stephen L. Clement is the research entomologist, Plant Germplasm Introduction and Testing Research Unit, U.S. Department of Agriculture (USDA), Agricultural Research Service (ARS), Washington State University, Pullman, WA, and affiliate professor of entomology, Department of Plant, Soil and Entomological Sciences, University of Idaho, Moscow, and scientist, Department of Entomology, Washington State University, Pullman. He obtained his Ph.D. in entomology in 1976 from the University of California, Davis, and was assistant professor of entomology, Ohio Agri­ cultural Research and Development Center, The Ohio State University (1976-1980), and research entomologist in USDA-ARS laboratories in Albany, California (1980) and Rome, Italy (1981-1986), before assuming his present position in 1986. Over the past 25 years Dr. Clement has written broadly on insect biology and behavior, insect pest management, plant resistance to insects, and biological weed control. His research projects have concentrated on rice, corn, rangeland weeds, artichoke, alfalfa, grain legumes, cool-season perennial grasses, fungal endophytes of grasses, and wheat. He is the author or coauthor of more than 115 professional papers, abstracts, and book chapters. Clement has presented or copresented more than 85 research papers before national and international audiences. Dr. Clement is a member of the Entomological Society of America, Crop Science Society of America, Kansas Entomological Society, International Organization of Biological Control, Pisum Genetics Association, and the honorary society Sigma Xi. He has been the recipient of research grants from Sigma Xi, U.S. Environmental Protection Agency, and USDA agencies (Cooperative State Research, Education, and Extension Service; Foreign Agricultural Service, Scientific Cooperation Program). Sharron S. Quisenberry is professor and head of the Department of Entomology, University of Nebraska, Lincoln, (1994-present) and courtesy professor in the Departments of Agronomy, Curriculum and Instruction, and Center for Biotechnol­ ogy, University of Nebraska-Lincoln. She obtained her Ph.D. in entomology (1980) from the University of Missouri, Columbia, and was assistant professor of entomol­ ogy at Iowa State University (1980-1982), assistant to professor of entomology at Louisiana State University (1982-1991), and professor and division chair of ento­ mology at the University of Idaho (1991-1994). Dr. Quisenberry has published in the areas of plant resistance to insects, insect/plant interactions, insect biology, insect pest management, and livestock insects related to insect/animal interactions and insecticide resistance. Her research projects have concentrated on insects related to wheat, rice, bermudagrass, tall fescue, alfalfa, and livestock. Dr. Quisenberry is the author or coauthor of more than 165 professional papers, and presented or copresented more than 90 papers at international, national, and regional professional meetings. She has received research

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grants from National Science Foundation, USDA (e.g., Cooperative State Research; National Research Initiative; Integrated Pest Management; Special Grants), Murdock Trust, and Ford Foundation. Dr. Quisenberry holds a U.S. Patent and germplasm registration with colleagues. Dr. Quisenberry is a member of the Entomological Society of America (ESA) and was the recipient of the ESA Southeastern Branch J. E. Bussart Memorial Award for Research (1991). She is a Board Certified Entomologist and member of Gamma Sigma Delta and Sigma Xi honor societies. Other professional societies include the American Agronomy Society, American Association for the Advancement of Sci­ ence, American Association of University Professors, Crop Science Society of America, and the Council for Agriculture Science and Technology.

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Contributors

Silvia Arnone Research Agronomist ENEA, Dipartimento Innovazione Settore Biotecnologie e Agricoltura CRE Casaccia Rome, Italy

Susan E. Cowgill, Ph.D. Doctor Centre for Plant Biochemistry and Biotechnology University of Leeds Leeds, United Kingdom

James D. Barbour, Ph.D. Assistant Professor Parma Research and Extension Center University of Idaho Parma, Idaho

Massimo Cristofaro Research Entomologist ENEA, Dipartimento Innovazione Settore Biotecnologic e Agricoltura CRE Casaccia Rome, Italy

David J. Boethel, Ph.D. Professor Department of Entomology Louisiana State University Baton Rouge, Louisiana Cesar Cardona, Ph.D. Entomologist Centro Internacional de Agricultura Tropical (CIAT) Cali, Colombia Edward E. Carey, Ph.D. Sweetpotato Breeder International Potato Center (CIP) Nairobi, Kenya Stephen L. Clement, Ph.D. Research Entomologist USDA/ARS Western Regional Plant Introduction Station Washington State University Pullman, Washington Wanda Collins, Ph.D. Deputy Director General — Research International Potato Center (CIP) Lima, Peru

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Francois du Toit, Ph.D. Research Station Manager Pannar (PTY) LTD Bainsvlei South Africa Sanford D. Eigenbrode, Ph.D. Assistant Professor Department of Plant, Soil and Entomological Sciences University of Idaho Moscow, Idaho Kathy L. Flanders, Ph.D. Assistant Professor and Extension Entomologist Department of Entomology Auburn University Auburn, Alabama Angharad M. R. Gatehouse, Ph.D. Doctor Department of Biological Sciences Science Laboritories University of Durham Durham, United Kingdom

Elvis A. Heinrichs, Ph.D. Adjunct Professor Department of Entomology University of Nebraska Lincoln, Nebraska Masa Iwanaga, Ph.D. Deputy Director General (Programme) International Plant Genetic Resources Institute (IPGRI) Rome, Italy Julia Kornegay, Ph.D. Director of Research Fairchild Tropical Garden Miami, Florida George R. Manglitz, Ph.D. Professor Emeritus Department of Entomology University of Nebraska Lincoln, Nebraska Il-Gin Mok, Ph.D. Plant Breeder

International Potato Center (CIP)

Bogor, Indonesia

Dolores W. Mornhinweg, Ph.D. Geneticist USDA/ARS Plant Science and Water Conservation Research Laboratory Stillwater, Oklahoma

Gary C. Peterson, Ph.D. Professor Agriculture Research and Extension Center Texas A&M University Lubbock, Texas David R. Porter, Ph.D. Research Geneticist USDA/ARS Plant Science and Water Conservation Research Laboratory Stillwater, Oklahoma Sharron S. Quisenberry, Ph.D. Professor and Head Department of Entomology University of Nebraska Lincoln, Nebraska Edward B. Radcliffe, Ph.D. Professor Department of Entomology University of Minnesota St. Paul, Minnesota C. Michael Smith, Ph.D. Professor Department of Entomology Kansas State University Manhattan, Kansas

Kanayo F. Nwanze, Ph.D. Director General West Africa Rice Development Association (WARDA) Bouaké, Côte d’Ivoire

Edgar L. Sorensen, Ph.D. Research Geneticist (Retired) USDA/ARS Department of Plant Pathology Kansas State University Manhattan, Kansas

Bonnie B. Pendleton, Ph.D. Postdoctoral Research Associate Department of Entomology Texas A&M University College Station, Texas

George L. Teetes, Ph.D. Professor Department of Entomology Texas A&M University College Station, Texas

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Paul Thompson, Ph.D. Professor Department of Plant Sciences Mississippi State University Pontotoc, Mississippi

Susanne Weigand Legume Entomologist (Retired) International Center for Agricultural Research in the Dry Areas (ICARDA) Aleppo, Syria

James A. Webster, Ph.D. Laboratory Director and Research Entomologist USDA/ARS Plant Science and Water Conservation Research Laboratory Stillwater, Oklahoma

Zhang Da Peng, Ph.D. Sweetpotato Breeder International Potato Center (CIP) Lima, Peru

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Table of Contents

FOREWORD Masa Iwanaga SECTION I: CEREAL CROPS Chapter 1 Germplasm Evaluation and Utilization for Insect Resistance in Rice Elvis A. Heinrichs and Sharron S. Quisenberry Chapter 2 The Value of Conserved Wheat Germplasm Evaluated for Arthropod Resistance C. Michael Smith, Sharron S. Quisenberry, and Francois du Toit Chapter 3 Insect Resistance in Barley Germplasm David R. Porter, Dolores W. Mornhinweg, and James A. Webster Chapter 4 Genetic Diversity of Sorghum: A Source of Insect-Resistant Germplasm George L. Teetes, Gary C. Peterson, Kanayo F. Nwanze, and Bonnie B. Pendleton SECTION II: LEGUME CROPS Chapter 5 Bean Germplasm Resources for Insect Resistance Cesar Cardona and Julia Kornegay Chapter 6 Assessment of Soybean Germplasm for Multiple Insect Resistance David J. Boethel Chapter 7 Germplasm Resources, Insect Resistance, and Grain Legume Improvement Stephen L. Clement, Massimo Cristofaro, Susan E. Cowgill, and Susanne Weigand

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Chapter 8 Alfalfa Germplasm Resources and Insect Resistance George R. Manglitz and Edgar L. Sorensen SECTION III: VEGETABLE CROPS Chapter 9 Vegetable Crops: Search for Arthropod Resistance in Genetic Resources James D. Barbour SECTION IV: ROOT AND TUBER CROPS Chapter 10 Utilization of Sweetpotato Genetic Resources to Develop Insect Resistance Wanda W. Collins, Edward E. Carey, Il-Gin Mok, Paul Thompson, and Zhang Da Peng Chapter 11 The Potato: Genetic Resources and Insect Resistance Kathy L. Flanders, Silvia Arnone, and Edward B. Radcliffe SECTION V: BASIC RESEARCH AND BIOTECHNOLOGY Chapter 12 Plant Genetic Resources for the Study of Insect-Plant Interactions Sanford D. Eigenbrode and Stephen L. Clement Chapter 13 Biotechnological Applications of Plant Genes in the Production of InsectResistant Crops Angharad M. R. Gatehouse

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Section I Cereal Crops

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1

Germplasm Evaluation and Utilization for Insect Resistance in Rice E. A. Heinrichs and Sharron S. Quisenberry

CONTENTS I. Introduction II. Germplasm Conservation III. Germplasm Evaluation and Utilization A. West Africa B. Asia C. Latin America D. North America IV. Biotechnology as a Tool to Enhance Resistance A. Tissue and Anther Culture B. Wide Crosses C. Transgenic Plants D. DNA Markers E. DNA Fingerprinting of Insects V. Integration of Resistant Cultivars A. Plant Resistance and Biological Control B. Plant Resistance and Cultural Control C. Plant Resistance and Chemical Control VI. Resistant Cultivars in IPM Systems VII. Conclusions References

I.

INTRODUCTION

The diversity and complexity of the constraints limiting food crop production call for a well-focused and strategic approach to agricultural development. Changing pest populations caused by the immigration of new pests, introduction of new crops, and cropping intensification attest to the need for dynamic crop breeding programs.22 Crop cultivars developed in such breeding programs that have resistance to the various abiotic and biotic stresses such as drought, salinity, soil toxicity, weeds,

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insects, and diseases are major components in integrated pest management (IPM) programs.43 The foundation of IPM programs should be laid with crop cultivars that are not only well adapted to the various abiotic constraints, but also have some level of resistance to one or more insect pests and diseases. The cultivar should be the best from an economic perspective, as conditioned by resistance to key pests and positive or negative interactions with other components of the production system. Development of rice, Orysa sativa L., cultivars having genetic resistance to insects requires an effective program of germplasm screening to identify genotypes that can be used as parents in rice breeding programs. These breeding programs must increase productivity per unit of agricultural land, and this must be accomplished in a way that conserves natural resources and the environment and meets the food requirements of people who are undernourished. The breeding of rice cultivars with multiple resistance to pests and tolerance to abiotic stresses is essential if we are going to experience another “Green Revolution,” which some have termed the “Super Green Revolution.”13 Breeding programs of the first “Green Revolution” involved close collaboration among plant breeders, agronomists, pathologists, and entomologists. The “Super Green Revolution” breeding programs should involve additional disciplines encompassing both the natural and social sciences. The availability of germplasm with a wide range of genetic diversity is essential to the success of these breeding programs.

II.

GERMPLASM CONSERVATION

There is a high level of genetic diversity in rice, Oryza sp., that exceeds that of most other crops. Throughout the long history of rice cultivation, rice strains adapted to the many different agroclimatic regions of the world have been selected by farmers. It has been estimated that there are more than 100,000 strains of rice in the world. Utilization of the genetic diversity which exists in rice has been economically profitable. For example, the contribution of rice landraces from genebanks in South Asia is estimated at $150 to $200 million (U.S.) per year.17 Included in the diversity that exists in the world collection of rice is genetic resistance to numerous rice insect pests. Rice germplasm collections, varying in size from a few hundred to several thousand accessions, are maintained in various countries by national research programs and by International Agricultural Research Centers (IARCs) of the Consultative Group on International Agricultural Research (CGIAR). The global collection consists of approximately 420,000 accessions, including duplicates (Table 1). The largest holdings and the global base collection are at the International Rice Research Institute (IRRI) in the Philippines (19% of the global total). Other large holdings are located in national programs in China (5%), India (5%), the U.S. (National Small Grains Collection, Aberdeen, Idaho, 5%), and at the West Africa Rice Development Association (WARDA) in Côte d’Ivoire (4%).17 The IRRI collection consists of about 77,000 accessions of O. sativa, 2,000 strains of O. glaberrima Steud., and 1,000 wild rices under long-term storage. About 11,000 duplicate or back-up accessions are maintained under long-term storage conditions at the National Seed Storage Laboratory (NSSL), Fort Collins, Colorado. Of the 10 largest rice germplasm collections, 7 are stored under long- or medium-term conditions.17 © 1999 by CRC Press LLC

TABLE 1 Status of Rice (Oryza) Germplasm Collections Genebank Institutea IRRI ICR-SAAS CRRI NSGC WARDA ICGR-CAAS RD NBPGR IITA NIAR NSSL CNPAF Univ. of Kyushu CIRAD IRCT-CIRAD ICRR-JAAS CRIA MARDI CENARGEN BRRI Others Total

Country/ organization CGIAR China India USA CGIAR China Thailand India CGIAR Japan USA Brazil Japan France France China Indonesia Malaysia Brazil Bangladesh

Accessions

Storage facilities (%)

Number

%

LTb

MTc

STd

Others

80634 20000 20000 19646 17440 16885 15350 12872 12315 11559 10833 8998 8000 7306 7131 6900 5917 5333 4734 4265 124223 420341

19 5 5 5 4 4 4 3 3 3 3 2 2 2 2 2 1 1 1 1 30 100

100 0 0 0 0 100 0 100 50 100 100 0 0 0 0 0 0 0 0 0

0 0 100 100 0 0 0 0 50 0 0 100 0 0 0 0 100 100 100 87

0 100 0 0 100 0 0 0 0 0 0 0 0 0 0 100 0 0 0 13

0 0 0 0 0 0 100 0 0 0 0 0 100 100 100 0 0 0 0 0

34

22

13

31

a

IRRI = International Rice Research Institute; ICR-SAAS = Institute of Crop Research-Sichuan Academy of Agricultural Sciences; CRRI = Central Rice Research Institute; NSGC = U.S. National Small Grains Collection; WARDA = West Africa Rice Development Association; ICGR-CAAS = International Center for Crop Germplasm Resources — Chinese Academy of Agricultural Sciences; RD = Rice Division; NBPGR = National Board for Plant Genetic Resources; IITA = International Institute for Tropical Agriculture; NIAR = National Institute of Agrobiological Resources; NSSL = U.S. National Seed Storage Laboratory; CNPAF = Centro Nacional de Pesquisa de Arroz e Feijão; CIRAD = Centre de Coopération Internationale en Recherche Agronomique pour le Developpement; IRCT-CIRAD = Institute de Rechenchers du Coton et des Textiles Exotiques — Centre de Coopération Internationale en Recherche Agronomique pour le Developpement; ICRR-JAAS = Institute of Crop Resources Research-Jiangsu Academy of Agricultural Sciences; CRIA = Central Research Institute for Agriculture; MARDI = Malaysian Agricultural Research and Development Institute; CENARGEN = Centro Nacional de Recursos Geneticos e Biotecnologia; BRRI = Bangladesh Rice Research Institute. b Long-term. c Medium-term. d Short-term. Data from FAO (Food and Agriculture Organization of the United Nations), The State of the World’s Plant Genetic Resources for Food and Agriculture, Rome, 1996. (Ref. 17)

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Estimates are that 95% of landraces and 10% of the wild species have been collected.17 On a global basis, accessions consist of 1% wild relatives, 25% landraces, 9% advanced cultivars/breeding lines, and 65% mixed categories or unknown material. For safety, 77% of O. sativa, 54% of O. glaberrima, and 65% of wild relatives have been duplicated at IRRI. The various rice germplasm collection and conservation programs have been successful on a global basis and have provided a large number of accessions for use in insect resistance screening and breeding programs.

III.

GERMPLASM EVALUATION AND UTILIZATION

Rice research programs in several regions of the world have active projects for the development of insect resistant rice cultivars (Table 2). Resistance to more than 30 insect species has been identified through the screening of germplasm collections and numerous insect-resistant cultivars are being used as components in pest management systems worldwide.21

A. WEST AFRICA West Africa dominates the sub-Saharan rice sector with 60% of the rice area and 56% of the production.42 Driven by urbanization and changing food preferences, rice consumption in sub-Saharan Africa has increased at more than 5% per annum since 1970 and rice is emerging as a commodity of strategic significance.71 Rice yields in West Africa average only 40% of the world imports, are rising at an annual rate of 6.8% since 1970, and are projected by the Food and Agriculture Organization (FAO) to increase to 4 mt by the year 2000 costing more than $1 billion (U.S.) in scarce foreign exchange. Rice availability and rice prices are major factors in the welfare of the poorest West African consumers who are the least food secure. Weeds, insects, and diseases are major constraints to rice production in West Africa. The most important insects are root feeders such as termites, several species of stem borers, the African rice gall midge, Orseolia oryzivora Harris and Gagne, foliage feeders including the rice caseworm, Nymphula depunctalis Guenée, and chrysomelid beetles which transmit rice yellow mottle virus (RYMV).23 Because of their importance in all West African rice ecosystems, stem borers have received the greatest emphasis in the development of resistant rice cultivars. Through the screening of rice germplasm by the International Institute of Tropical Agriculture (IITA), Institut de Recherche Agronomique Tropicale (IRAT), WARDA, and the National Cereals Research Institute (NCRI) in Nigeria, cultivars with resistance to the stem borers Diopsis longicornis Macqart, Chilo zacconius Blezynski, Maliarpha separatella Ragonot, and Sesamia calamistis (Hampson) have been identified.42 Screening of rice cultivars for resistance to D. longicornis has been conducted at IITA in Nigeria,3-5,65 at Bouaké, Côte d’Ivoire,57 and in Senegal.69 Cultivars with moderate levels of resistance were from Asia, Africa, and South America. In addition, nine tropical O. glaberrima (Tog) cultivars from Liberia were selected. In his report on screening at IITA, Akinsola2 lists 35 cultivars as moderately resistant to D. longicornis (Table 3). A review of the literature indicates 54 O. sativa and 10 tropical O. glaberrima cultivars as moderately resistant to D. longicornis.23

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TABLE 2 Status of Screening and Breeding for Varietal Resistance to Rice Insect Pests

Common name White stem borer Pink borer Stalk-eyed fly African rice gall midge Asian rice gall midge Brown planthopper Whitebacked planthopper Green leafhopper Zigzag leafhopper White leafhopper Blue leafhopper Striped stem borer Yellow stem borer Whorl maggot Seedling fly Armyworm Thrips Rice bug Black bug Caseworm Leaffolder

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Scientific name Maliarpha separatella Sesamia calamistis Diopsis longicornis Orseolia oryzivora Orseolia oryzae Nilaparvata lugens Sogatella furcifera Nephotettix virescens Recilia dorsalis Cofana spectra Empoascanara maculifrons Chilo suppressalis Scirpophaga incertulas Hydrellia philippina Atherigona exigua Mythimna separata Stenchaetothrips biformis Leptocorisa oratorius Scotinophara latiuscula Nymphula depunctalis Cnaphalocrocis medinalis

Region West West West West Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia Asia

Africa Africa Africa Africa

Screening methods developed

Resistance sources identified

Resistant cultivar released

Genes for resistance identified

+ + + + + + + + + + + + + + + + + + + + +

+ + + + + + + + + + + + + + + – + + + + +

– + – – + + + + + – – + + + – – + – + – –

– – – – + + + + + – – – – – – – – – – – –

TABLE 2 (continued) Status of Screening and Breeding for Varietal Resistance to Rice Insect Pests

Common name Leaffolder Hispa Bloodworm Rice delphacid Sugarcane borer Lesser cornstalk borer Whorl maggot Fall armyworm Rice water weevil Rice weevil Maize weevil Lesser grain borer Angoumois grain moth

Scientific name Marasmia patnalis Dicladispa armigera Chironomus tepperi Tagosodes orizicolus Diatraea saccharalis Elasmopalpus lignosellus Hydrellia griseola Spodoptera frujiperda Lissorhoptrus oryzophilus Sitophilus oryzae Sitophilus zeamais Rhyzopertha dominica Sitotroga cerealella

Region

Screening methods developed

Resistance sources identified

Resistant cultivar released

Genes for resistance identified

Asia Asia Australia Latin America Latin America Latin America Latin America North America North America Worldwide Worldwide Worldwide Worldwide

+ + + + + + + + + + + + +

+ + + + + + + + + + + + +

– – – + + – + – – – – – –

– – – – – – – – – – – – –

Modified from Heinrichs, E.A., Rice insects: the role of host plant resistance in integrated pest management systems, Korean J. Appl. Entomol., 31:256-275, 1992. (Ref. 21)

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TABLE 3 Rice Cultivars Tested and Identified with Moderate Resistance to Stem Borers at the International Institute of Tropical Agriculture, Ibadan, Nigeria Stem borer Diopsis longicornis Chilo zacconius Maliarpha separatella Sesamia calamistis

Accessions tested (no.)

Moderately resistant accessions (no.)

Type of test

2,036 600 2,000 800

35 10 5 6

Field and screenhouse Field and screenhouse Field Field and screenhouse

Data from Akinsola, E.A., The Biology and Ecology of Rice Stem Borers in Nigeria, Ph.D. Dissertation, University of Ibadan, Ibadan, 1979. (Ref. 1)

Screening studies for C. zacconius resistance were started at IITA in 1973 when three Asian cultivars, “H8,” “Malagkit Sung Song,” and “Ratna,” were selected out of 400 cultivars screened.34 In 1975, 200 cultivars were screened in the field and then retested in the screenhouse at IITA where “TOs 2513” and “Taitung 16” were selected.65 Ukwungwu67 evaluated 60 cultivars previously identified as having resistance to certain insects and found “Lac 23” (white) from Liberia and “IR 4625-123-1-2” as having moderate resistance. Akinsola2 reported 10 moderately resistant accessions out of 600 tested at IITA (Table 3). The literature lists 22 cultivars from 9 countries (mostly Asian) as having moderate resistance.23 Field screening for M. separatella resistance at Badeggi, Nigeria,1 at IITA in Ibadan, Nigeria,65 and in Man, Côte d’Ivoire,57 identified 14 accessions as having moderate resistance. A total of 26 accessions has been reported as having moderate resistance.23 Resistant donors have been used as parents in the WARDA breeding program and, in 1991, 10 lines with resistance under high insect pressure were reported by WARDA breeders located at IITA.23 Only 10 cultivars have been selected for resistance to S. calamistis.23 Of these, “Sikasso” from Nigeria and “Taitung 16” from Taiwan have been used as parents in the breeding program.2 The African rice gall midge has only recently become a pest in West Africa, especially in Burkina Faso and Nigeria.68 No resistant sources have been identified although extensive screening of Asian germplasm resistant to the Asian rice gall midge, O. oryzae (Wood-Mason), has been conducted. In 1993, the WARDA/Commonwealth Agricultural Bureaux (CABI) Gall Midge Project began operation to identify, among other objectives, gall midge-resistant germplasm for use in the breeding program. Although cultivars with resistance to C. zacconius and S. calamistis have been released (Table 2), resistant cultivars still occupy a small percentage of the rice area in West Africa. Commercial cultivars with moderate resistance to D. longicornis are “BG 90-2,” “IR 8,” “Faro 188A,” “Iguape Cateto,” and “I Kong Pao.”23 “BG 90-2” is widely grown in West Africa, especially in the irrigated Sahel area of Mali. The widely adopted traditional upland cultivar “Lac 23” has moderate resistance to C. zacconius.

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The cultivars with resistance to M. separatella and S. calamistis can be used as donor parents in the breeding of new stem borer resistant cultivars, although resistance levels are only moderate. Identification of germplasm with higher levels of resistance to the various stem borer species and other major insect pests is needed. Once identified, resistant germplasm can be used in the development of high yielding, multiple pest-resistant rice cultivars that are also tolerant to the abiotic constraints in West Africa.

B. ASIA Sources of resistance to more than 30 insect species have been identified in the world rice collections (Table 2).21 There are numerous rice breeding programs in Asia that include resistance to insects as a breeding objective, including the national programs and the IRRI program. These programs have made significant progress in the development and release of pest-resistant cultivars. The All India Coordinated Rice Improvement Project (AICRIP) through the screening of rice germplasm at 20 sites has identified germplasm resistant to seven rice insect pests.59 More than 40 cultivars with multiple resistance to as many as four insect species have been commercially released.59 The first hybrid rice developed by Chinese scientists was released in 1976 and by 1991 the area under hybrid rice cultivation reached 17.6 million ha (55% of China’s total rice area) and contributed to 66% of the country’s rice production.76 Chinese hybrids have been adapted for the tropics and national programs in India, Indonesia, Malaysia, Philippines, and Vietnam. They have 15% yield advantage compared with open pollinated cultivars, possess good grain quality, and have multiple disease and insect resistance.6 The Korean national rice breeding program has emphasized resistance to planthoppers and leafhoppers in the development of multiple pest-resistant cultivars. One cultivar, “Namyeongbyeo,” is resistant to five diseases, namely bacterial leaf blight Xanthomonas pv. oryzae (Ishyama) Dye, black-streaked dwarf virus (fijivirus), blast Pyricularia oryzae Cavara, dwarf virus (reovirus), and stripe virus (tenuivirus); three hopper species, namely brown planthopper Nilaparvata lugens (Stål), green leafhopper Nephotettix cincticeps (Uhler), and smaller brown planthopper Laodelphax striatella (Fallen); and a nematode Aphelencoides besseyi Christie. “Namyeongbyeo” yields in multi-location trials were 5.69 t/ha, 10% higher than the highest yielding check.64 The IRRI rice improvement program emphasizes the development of germplasm with multiple resistance to key diseases and insects.29,39 Screening procedures for numerous rice insect species have been developed.28 Emphasis in screening and breeding activities has been on the major insect pests including the Asian gall midge, brown planthopper, green leafhopper, yellow stem borer Scirpophaga incertulas (Walker), and the striped stem borer Chilo suppressalis (Walker) (Table 2). Cultivars with multiple resistance to these pests and the diseases blast, bacterial leaf blight, sheath blight Rhizoctonia solani Kuhn, tungro virus (spherical form = ribotungrovirus; bacilliform = badnavirus), and grassy stunt virus (tenuivirus), have been developed.24 The widespread adoption of these cultivars has helped stabilize rice production in Asia.39

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In Asia, where rice is mostly grown under irrigated conditions, modern cultivars have rapidly spread. Ninety-four percent of the rice hectarage is planted to modern cultivars in the Philippines. Furthermore, modern cultivars occupy 100% of the rice area in China and Japan.40 Most of the cultivars planted have resistance to at least one insect. Cultivars developed at IRRI have had a significant economic and social impact in Asia.40 Cultivar “IR36,” with multiple resistance to insects and diseases, was once grown on about 16 million ha in Asia, but has been replaced by newer cultivars which have multiple resistance to at least seven key diseases and insects. The availability of rice cultivars with multiple resistance to insects and diseases has minimized the need for pesticides and has promoted the adoption of IPM practices. Indonesia’s national IPM policy resulted in the banning of 57 rice insecticides.24

C. LATIN AMERICA Rice is a major food crop in Latin America where pest control expenses contribute to increasing costs and decreasing profitability of rice production.73 Crop protection costs in Colombia account for 22% of total production costs and insecticides represent 35% of the total pest control costs in Latin America (Rice Program Database, Centro Internacional de Agricultura Tropical [CIAT], Cali, Colombia). Cost of rice insecticides in Latin America is $170 million (U.S.) per year. CIAT scientists predict that with improved management, including host plant resistance, insect control costs can be cut to 0.2% of total production costs which is equal to the value of 8 kg/ha of paddy rice. Insect-resistant cultivars are one component in an overall strategy for developing commercially acceptable rice cultivars in Latin America. Screening has identified cultivars with resistance to the “sogata” planthopper Tagosodes orizicolus (Muir), sugarcane borer Diatraea saccharalis (F.), whorl maggot Hydrellia griseola (Fallen),73 and the lesser cornstalk borer Elasmopalpus lignosellus (Zeller).28 Major emphasis in pest resistance breeding in Latin America is on the “sogata” planthopper and the rice hoja blanca virus (RHBV) (tenuivirus) that this insect vectors. The uncertainty of RHBV epidemics in the Andean, Central American, and Caribbean countries causes farmers to apply “insurance sprays” even when the problem is not apparent. It is estimated that about half of the lowland farmers in these areas apply one insecticide spray for planthopper-RHBV each cropping season, costing approximately $15 million (U.S.) annually.74 Numerous sources of resistance against the planthopper have been identified in CIAT screening. Planthopper-resistant cultivars were released in 1970 (“CICA 4”) and the level of resistance was increased in “CICA 8” by 1978, based on the “Tetep” cultivar source.74 Most of the rice cultivars, including “CICA 4” and “CICA 8,” grown in Latin America are tolerant to the planthopper and outbreaks on these cultivars have been rare. These cultivars tolerate natural pest fluctuations and allow beneficial arthropods to maintain planthopper populations at sub-economic levels. Surveys of commercial fields throughout Colombia indicated that no insecticide applications were necessary when tolerant cultivars were planted even though the maximum planthopper densities exceeded the action threshold. Because of cultivar

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tolerance, direct damage by the planthopper was insignificant and biological agents were allowed to regulate population densities below the action threshold.73 More than 70% of the major commercial cultivars in Latin America have resistance to the planthopper, 50% have resistance to the whorl maggot, and 80% have resistance to the sugarcane borer. Some cultivars have occupied large areas, such as “J 104” (110,000 ha in Cuba) and “Oryzica 1” (260,000 ha in Colombia).73 Economic analyses indicate that the breeding and utilization of planthopperRHBV resistant cultivars is highly profitable. An impact assessment of CIAT research projects in the years 1997 to 2010 for breeding planthopper-RHBV resistant rice cultivars indicated a net present value of $616 million (U.S.).56 In a subsequent impact assessment, this resistance was predicted to provide a high internal rate of return of 63%.33

D. NORTH AMERICA Rice is an important crop in the economies of Arkansas, California, Louisiana, Mississippi, Missouri, and Texas in the U.S. Although the U.S. only grows 1% of the world’s rice production, yields are high (6.2 t/ha) and the U.S. is a major rice exporter.36 The rice water weevil, Lissorhoptrus oryzophilus Kuschel, is the major rice pest in North America causing significant damage to rice root systems.60 It ranges from Central America to Canada and has recently been introduced into Japan from the U.S.,66 and has since spread to Korea41 and Taiwan.58 Screening of germplasm in the U.S. has emphasized this pest but some germplasm screening for the fall armyworm, Spodoptera frugiperda (J.E. Smith),53 and studies on the effect of cultivar height on the least skipper, Ancyloxypha numitor (F.),63 also have been conducted. Because all cultivars currently grown in the U.S. are susceptible to larvae of the rice water weevil, research has attempted to identify sources of resistance in rice lines and cultivars.62 Based on the field screening technique of Bowling,10 more than 8,000 USDA Rice World Collection lines have been screened for larval resistance. Two lines designated WC1711 and CI11048 were identified in California20 and three lines (WC1403, WC1349, WC1815) were identified in Louisiana as having moderate tolerance to the rice water weevil.19 These lines have been used as parents in the breeding program in Louisiana. Further screening in Louisiana has identified several promising breeding lines. Field screening in 1990 identified AL6029, LA2218, TX22041, URN199, and URN200 as having tolerance, and TX12685 and TX13079 as having low levels of antixenosis.47 Greenhouse evaluations in 1991 and 1992 identified the breeding lines 8723518, 8725417, and 8725454 as having resistance based on larval numbers on the roots.50 In a 1991-1992 field evaluation of 40 breeding lines, lines 8720906 and 8721937 exhibited moderate levels of tolerance and lines 872239 and 8721317 had low levels of tolerance.51 These lines supported high larval populations, but yields did not decline or were higher when compared with treated controls. This germplasm also recovered from root pruning damage caused by larval feeding. Pantoja et al. screened about 5,000 rice accessions to the fall armyworm in the greenhouse in Louisiana.53 Moderate resistance to larval defoliation was detected in

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plant introductions (PI) 160842, 346830, 346833, 346840, and 346853 from the U.S. National Small Grains Collection. Moderate levels of resistance were also detected in U.S. cultivars “Honduras,” “Newrex,” and “Chinese,” and in O. glaberrima Steud accessions 101800, 102554, and 369453 from the IRRI collection. However, no commercial cultivars with resistance to this pest have been released (Table 2). The need for cultivars with higher levels of resistance continues in spite of the extensive screening of germplasm for rice water weevil and fall armyworm resistance. This need has increased with the banning of effective insecticides for water weevil control in the U.S.51 On an international basis, the need has become increasingly acute with the spread of the rice water weevil to Japan, Korea, and Taiwan where this pest causes serious economic damage. Biotechnology techniques have been used to enhance levels of resistance to the rice water weevil.14

IV.

BIOTECHNOLOGY AS A TOOL TO ENHANCE RESISTANCE

Several biotechnological approaches are employed in the development of insectresistant rice cultivars. Tissue and anther culture techniques are utilized to increase the level of resistance, and wide hybridization and transformation contribute to the generation of novel genetic variation by increasing the gene pool available to breeders. DNA markers of rice and DNA fingerprinting help to make selection, phenotypic analysis, and biotype studies more efficient and powerful.9,46

A. TISSUE

AND

ANTHER CULTURE

Tissue culture and anther culture techniques have been used in Louisiana in attempts to develop germplasm with higher levels of rice water weevil resistance than has been found in conventionally bred lines. In a field evaluation of 66 tissue culture lines in 1990, lines 244 and 2232 showed antixenotic resistance and line 112 tolerance to the rice water weevil.48 Both were regenerated from tissue culture of the susceptible cultivar “Tebonnet.” The anther culture lines 952836 and 953527 exhibited moderate levels of resistance to the rice water weevil in field evaluations of 43 anther culture lines in 1990 and 1991.49

B. WIDE CROSSES Wild species have provided resistance sources where screening of the O. sativa germplasm collection has failed to identify adequate levels of resistance to certain rice insects.32,44,75 Although wild species are often incompatible with O. sativa in conventional breeding, they can be used as donor parents using wide hybridization and embryo rescue techniques.11 Rescue of hybrid embryos through tissue culture and backcrossing to O. sativa allows recovery of new lines which contain segments of the wild genome introgressed into the O. sativa background.8 In studies at IRRI, genes from seven wild rice species have been transferred to O. sativa for resistance to the brown planthopper, whitebacked planthopper Sogatella furcifera (Horvath), and yellow stem borer (Table 4). IRRI breeding lines possessing a gene for brown planthopper resistance from O. officinalis were tested in the Mekong Delta, Vietnam,

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TABLE 4 Status of the Transfer of Genes for Insect Resistance from Wild Rice Species to Oryza sativa by Wide Hybridization and Embryo Rescue Resistance to: Brown planthopper

Whitebacked planthopper Yellow stem borer

Donor species (Genome) O. O. O. O. O. O. O. O.

officinalis (CC) minuta (BBCC) latifolia (CCDD) australiensis (EE) granulata (unknown) officinalis (CC) brachyantha (FF) ridleyi (unknown)

Status Completed Completed Completed Completed Material under test Completed Material under test Material under test

From Brar, D.S. and G.S. Khush, Wide hybridization for enhancing resistance to biotic and abiotic stresses in rainfed lowland rice, In: Proc. Int. Rice Res. Conf., International Rice Research Institute, Los Baños, 1995. (Ref. 11)

where they were more resistant than test cultivars with known resistance genes.54 Current work at IRRI concentrates on the transfer of yellow stem borer resistance from O. brachyantha to O. sativa because only moderate levels of resistance to the stem borer have been identified in O. sativa germplasm.8

C. TRANSGENIC PLANTS The transgenic approach to enhancing resistance to rice insects is attractive because it provides access to non-rice genes, allows purified rice genes to be returned to rice after modifications that give enhanced performance not attainable through mutation and recombination in vivo, and allows the addition of specific characters into rice without the linkage drag and the requirement for backcrossing that accompany sexual hybridization.8 Scientists at IRRI and at other locations linked by the Rockefeller Foundation Biotechnology Network are incorporating novel genes for resistance into rice through transformation. The genes include the Bacillus thuringiensis Berliner (Bt) genes cryIA(b) and cryIA(c) under the control of various promoters and several proteinase inhibitor genes. The rice insects controlled by these genes include the yellow stem borer, striped stem borer,8 and the leaffolder Cnaphalocrocis medinalis (Guenée).18 Genetic engineering of rice with toxin genes from Bt has the potential to provide effective and environmentally safe control of several rice insect pests. Of major concern, however, is its potential lack of stability.9 Numerous strategies have been proposed to delay the evolution of rice pest resistance to Bt toxins in transgenic rice plants.9,12 One strategy is the use of spatial refuges in association with the combination of multiple Bt toxins that bind to different receptors. Proposed spatial refuges consist of tissue specific expression, within field mixtures of Bt and non-Bt rice plants, and field-to-field mixtures of Bt and non-Bt rice. Cohen et al.12 reported that the genetic engineering of rice is progressing rapidly and Bt genes may soon be in agronomic © 1999 by CRC Press LLC

backgrounds suitable for use by farmers. They caution that genetic engineering is proceeding at a faster pace than the development of resistance management strategies.

D. DNA MARKERS Genetic (DNA) markers are being used to accelerate breeding for resistance to the brown planthopper and the Asian gall midge, both major pests of rice in Asia. These markers are being used to cope with the problem of breeding rice cultivars with resistance to insect biotypes. The use of DNA markers enables the breeding program to progress during seasons when there is inadequate insect pressure for field screening, permits breeding to be conducted in countries where the insect does not occur, and allows the pyramiding of genes for more stable resistance. DNA marker-aided selection is being used in breeding for gall midge resistance in China and India as part of a collaborative program coordinated by IRRI. The objective in China is to transfer the gall midge-resistant gene Gm6(t) to the parental lines used in hybrid rice production, while in India the objective is to transfer the resistant gene Gm2 to new germplasm developed for the rainfed lowlands.8

E. DNA FINGERPRINTING

OF INSECTS

Biotypes are a major constraint to the breeding for gall midge resistance in Asia.30 DNA fingerprinting, using restriction fragment length polymorphism (RFLP), random amplified polymorphism DNA (RAPD), and amplified fragment length polymorphism (AFLP) markers, is being used as a tool to study the genetic diversity of this insect in Cambodia, India, Indonesia, Laos, Myanmar, Nepal, Sri Lanka, and Thailand.8,16 The AFLP technique gives the highest degree of reproducible variation between different gall midge isolates and is sufficiently sensitive to be applied to individual insects in a population.70

V.

INTEGRATION OF RESISTANT CULTIVARS

Rice IPM programs employ cultural controls, pesticides, natural enemy conservation, and pest-resistant cultivars as tactics in the management of rice insect pests. The extent to which each component is integrated into the system is dependent on the nature of the pest complex and the socioeconomic conditions existing in the country.61 Where insect-resistant cultivars are available, they are often used as a basic component in the management of rice pests.22 An advantage of using host plant resistance is that, in general, it is compatible with other control tactics.

A. PLANT RESISTANCE

AND

BIOLOGICAL CONTROL

Complexes of biological control agents have been identified for most major rice insect pests and studies indicate that they have the potential to regulate rice insect populations.52,55 In addition to the joint pest controlling roles of host-plant resistance and biological controls, there exists the longer-term significance of their interaction in delaying or preventing the development of biotypes capable of overcoming previously resistant rice cultivars.72 Insect-resistant cultivars may have an adverse effect on © 1999 by CRC Press LLC

natural enemies by reducing prey density, but generally have a positive effect on natural enemies, especially parasites and predators, by minimizing the need for the application of toxic insecticides. Research on the interactions between plant resistance and biological control has increased in recent years and has shown the positive effects of combining the two tactics.61 The combination of the two control tactics may be additive, or even synergistic, in their effect on decreasing pest populations. Because the use of this combination requires minimal training, it is especially attractive in countries where extension services are not able to train farmers in IPM principles and practices. Examples of how these two tactics can be effectively integrated follow. The green leafhopper is an important rice pest in Asia because it is a vector of the pathogen causing rice tungro virus (RTV). Combinations of green leafhopperresistant cultivars and predation by the mirid bug, Cyrtorhinus lividipennis Reuter, have a cumulative effect on green leafhopper populations. In greenhouse studies at IRRI, the number of green leafhoppers only reached six on the leafhopper resistant rice cultivar “IR29” with the predator, and 31 without the predator, while there were 91 and 220 leafhoppers, respectively, on susceptible “IR22” with and without the predator.45 The cumulative effect was evident in the green leafhopper mortality rates. Predatory activity by the mirid bug increased leafhopper mortality on all cultivars by approximately 30%; from 66 to 92% on highly resistant “IR29,” 34 to 56% on moderately resistant “IR8,” and 7 to 40% on susceptible “IR22.” The cumulative effect of antibiosis and predation resulted in 52% greater leafhopper mortality in the highly resistant “IR29” as compared with susceptible “IR22.”45 Insect-resistant rice cultivars can enhance the activity of predators resulting in a synergistic effect of the two control tactics. Cyrtorhinus lividipennis predation rate increased when the prey, brown planthopper nymphs, fed on the resistant rice cultivar “IR36.”37 This may be caused by increased movement of planthopper nymphs on resistant plants, facilitating detection of prey.

B. PLANT RESISTANCE

AND

CULTURAL CONTROL

The integration of insect-resistant cultivars with cultural management can be a powerful tool in managing pests.43 Early maturing cultivars and planting date are cultural practices employed to evade rice insect attack. In field studies in the Philippines, brown planthopper populations and planthopper:predator ratios on early maturing rice cultivars were significantly lower than those on later maturing cultivars because the insect was not able to complete as many generations on the early maturing cultivars as compared with long duration cultivars.25 A shift of planting dates minimized damage to several stem borer species in India, Indonesia, and Malaysia.38 This strategy could be integrated with plant resistance. However, caution should be taken before using pest evasion tactics because they may have negative attributes, such as yield loss caused by planting at a sub-optimal date. Nitrogen fertilizer is a major component for producing high yields of modern rice cultivars. High plant nitrogen generally favors insect pest populations and is manifested in greater pest survival, increased feeding rate, increased fecundity, and faster growth.15 In a greenhouse study, nitrogen fertilizer was shown to favor population growth of brown planthoppers on rice, even on a resistant cultivar.27 However, brown

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planthopper population growth was least at the higher levels of cultivar resistance. Thus, when high nitrogen levels are needed to maximize rice production, planting of resistant cultivars will minimize brown planthoppers populations.

C. PLANT RESISTANCE

AND

CHEMICAL CONTROL

Control of rice insects with insecticides has been shown to be more effective when the host plant is an insect-resistant cultivar as compared with a susceptible cultivar. Depending on the insect pest complex, the level of insect pressure, and the degree of plant resistance, insecticides in combination with an insect-resistant cultivar can increase grain yields above that of a resistant cultivar alone.31 A cumulative effect was observed when plant resistance and insecticides were integrated in the control of the green leafhopper (vector of RTV) in field studies in the Philippines.31 The cumulative effect was based on the percentage of RTV-infected plants in the various cultivars tested and was dependent on the level of green leafhopper resistance in the cultivar and insecticide rate. Cultivar “IR28,” with a high level of green leafhopper resistance, had a low pest population and a low incidence of RTV at all insecticide rates including the untreated control. Conversely, the cultivar with high susceptibility to green leafhopper (“IR22”) had a high level of RTV regardless of the rate of insecticide applied. However, the moderately resistant cultivar “IR36” showed a cumulative effect. Resistance of “IR36” to the green leafhopper alone reduced RTV incidence 50% below that of the susceptible cultivar “IR 22.”31 When used in combination with insecticide, there was a rate effect with RTV incidence decreasing from 42% at the 0.0 kg/ha rate to 30% and 11% at the 0.25 and 0.5 kg/ha insecticide rate, respectively. Grain yields were related to percent RTV incidence. This study indicated the need for high levels of resistance to the vector insect in controlling RTV and the cumulative combination effect of moderate resistance and insecticides to the control of the virus vector.31 Results of a greenhouse study indicated that insecticides cause higher mortality to planthoppers feeding on resistant rice rather than on susceptible rice.26 Both the whitebacked planthopper and the brown planthopper were killed at low insecticide rates when feeding on resistant rice cultivars. Mortality rates of biotype 2 brown planthoppers, reared on the moderately resistant “ASD7,” were 2 to 12 times higher than those reared on susceptible “IR26” when treated with spray applications of various insecticides.26 In the same study, LD50 rates of the whitebacked planthopper were three times as high on susceptible “TN1” than hoppers feeding on moderately resistant “N22.”26 Plant resistance is important in minimizing the extent of insecticide-induced brown planthopper resurgence. In field studies at IRRI, brown planthopper populations on rice plants treated with a resurgence-inducing insecticide only reached 10 insects per hill on a resistant rice cultivar, whereas populations on a susceptible cultivar reached 1,100 per hill.7 Thus, when insecticide is needed to control a defoliator, the level of insecticide-induced brown planthopper resurgence can be reduced, or even eliminated, by the planting of a hopper-resistant cultivar.

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VI.

RESISTANT CULTIVARS IN IPM SYSTEMS

A major portion of the world collection has been evaluated for resistance to rice insects. Many resistant accessions have been identified and some have been used as donor parents to produce high yielding rice cultivars with multiple resistance to pests and tolerance to abiotic stresses. The increased pest problems accompanying the “Green Revolution” have sparked a movement toward an integrated approach to the management of rice insect pests, especially in Asia and Latin America. Pest-resistant cultivars have served as key components in IPM programs and in the development of sustainable rice production systems. Insect-resistant rice cultivars have been successfully used in IPM programs throughout Asia and other regions of the world. The Korean rice IPM program uses insect resistance as a major component.41 The cultivar “Namyeongbyeo,” with high yield potential, is resistant to five diseases, three insects, and a nematode.64 The Sri Lankan cultivar “BG367-3” is resistant to blast, brown spot Cochliobolus miyabeanus Ito and Kuribayashi, tungro virus, leaffolder, Asian gall midge, green leafhopper, leaf miner Dicladispa armigera (Oliver), and stem borer Sesamia inferens (Walker). The AICRIP has identified germplasm resistant to seven insect species.59 Use of these sources as gene donors in the Indian rice improvement program has produced more than 40 commercial cultivars with multiple insect resistance, some with resistance to as many as four pest species. Insect-resistant cultivars have minimized the need for pesticides and promoted the adoption of IPM practices and thus stabilized yields at higher levels than previously achieved. Because of insect resistance, early sprays of insecticides are seldom necessary and pests are maintained at sub-economic levels by biological control agents. Indonesia established a national IPM policy based on the integration of plant resistance, biological control, and cultural practices. The policy has led to an upward trend in rice yields and reduced insecticide use resulting in improved health of the growers and improved environmental quality.40

VII.

CONCLUSIONS

In spite of remarkable progress in increasing rice production through genetic improvement and expansion of crop area, the global pursuit for food self-sufficiency continues as the world population rapidly increases. Population growth in rice growing countries results in 80 to 100 million additional people to feed each year.35 Rice production must approximately double by the year 2020 to feed the world population, which will approach 8 billion. Unless technology rapidly advances, lack of suitable land will cause farmers to till highly erodible and marginal land to meet food demands. Insect-resistant rice cultivars integrated with other pest management tactics contribute to crop pest management strategies that are environmentally and economically acceptable. These tactics are a vital component in technology packages that will increase the productivity of existing land resources. Although significant progress has been achieved in the development and commercial use of insect-resistant rice cultivars, plant resistance has the potential to play an even greater role in rice production systems in the future. Because of various

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constraints in the breeding of insect-resistant cultivars, there are still numerous insect species for which resistant cultivars have not been developed and for which plant resistance as a management tactic has not been used. Biotechnology tools, including wide hybridization, transformation, DNA markers, and DNA fingerprinting, are being used to overcome constraints of breeding rice cultivars with resistance to insects and diseases. These approaches will contribute to the breeding of resistant cultivars for improved crop protection, biodiversity conservation, and profitability.

REFERENCES 1. Akinsola, E. A., The Biology and Ecology of Rice Stem Borers in Nigeria, Ph.D. Dissertation, University of Ibadan, Ibadan, 1979. 2. Akinsola, E. A., Varietal resistance to stem borer pests of rice management in West Africa: a review, Insect Sci. Applic., 8:771-776, 1987. 3. Alam, M. S. and Y. Efron, Resistance of rice to the stalk-eyed fly Diopsis macrophthalma, Int. Rice Comm. Newsl., 35:40-45, 1986. 4. Alghali, A. M., Relative susceptibility of some rice cultivars to the stalk-eyed fly, Diopsis thoracica West., Insect Sci. Applic., 4:135-140, 1983. 5. Alghali, A. M. and E. O. Osisanta, The effects of some rice cultivars on the biology of the stalk-eyed fly Diopsis thoracica West. (Diptera: Diopsidae), Insect Sci. Applic., 3:163-166, 1982. 6. Anonymous, Hybrid seed controversy in India, In: Biotechnology and Development Monitor, No. 19, University of Amsterdam, Amsterdam, 1994, 9. 7. Aquino, G. B. and E. A. Heinrichs, Brown planthopper populations on resistant cultivars treated with a resurgence-causing insecticide, Int. Rice Res. Newsl., 45:12, 1979. 8. Bennett, J., M. B. Cohen, S. K. Katiyar, B. Ghareyazie, and G. S. Khush, Enhancing insect resistance in rice through biotechnology, In: N. Carozzi and M. Koziel, Eds., Advances in Insect Control: The Role of Transgenic Plants, Taylor Francis, London, 1997, 75. 9. Bottrell, D. G., R. M. Aguda, F. L. Gould, W. Theunis, C. G. Demayo, and V. F. Magalit, Potential strategies for prolonging the usefulness of Bacillus thuringiensis in engineered rice, Korean J. Appl. Entomol., 31:247-255, 1992. 10. Bowling, C. C., Tests to determine varietal reaction to rice water weevil, J. Econ. Entomol., 56:893-894, 1963. 11. Brar, D. S. and G. S. Khush, Wide hybridization for enhancing resistance to biotic and abiotic stresses in rainfed lowland rice, In: Proc. Int. Rice Res. Conf., International Rice Research Institute, Los Baños, 1995. 12. Cohen, M. B., R. M. Aguda, A. M. Romena, G. K. Roderick, and F. L. Gould, Resistance management strategies for Bt rice: what have we learned so far? In: G. S. Khush, Ed., Rice Genetics III: Proc. 3rd Int. Rice Genetics Symp., International Rice Research Institute, Los Baños, 1996, 749. 13. Conway, G., U. Lele, J. Peacock, and J. Pineiro, Sustainable Agriculture for a Food Secure World, a Vision for the Consultative Group on International Agricultural Research, Consultative Group on International Agricultural Research, Washington D.C., 1994. 14. Croughan, T. P. and J. F. Robinson, Application of tissue culture techniques to the development of insect resistant rice, In: E. A. Heinrichs and T. A. Miller, Eds., Rice Insects: Management Strategies, Springer-Verlag, New York, 1991, 275.

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15. Dale, D., Plant mediated effects of soil mineral stresses on insects, In: E. A. Heinrichs, Ed., Plant Stress-Insect Interactions, Wiley, New York, 1988, 35. 16. Ehtesham, N. Z., J. S. Bentur, and J. Bennett, Characterization of a DNA sequence that detects repetitive DNA elements in the Asian rice gall midge (Orseolia oryzae) genome: potential use in DNA fingerprinting of biotypes, Gene 153:179-183, 1995. 17. FAO (Food and Agriculture Organization of the United Nations), The State of the World’s Plant Genetic Resources for Food and Agriculture, Rome, 1996. 18. Fujimoto, H., K. Itoh, M. Yamamoto, J. Kyozuka, and K. Shimamoto, Insect resistance generated by introduction of a modified α-endotoxin gene of Bacillus thuringiensis, Biotechnology 11:1151-1155, 1993. 19. Gifford, J. R. and G. B. Trahan, Rice water weevil and rice stalk borer resistance, In: 67th Ann. Progress Rep., Rice Exp. Stn., Louisiana Agricultural Experiment Station, Baton Rouge, 1975, 125. 20. Grigarick, A. A., M. O. Way, and S. L. Clement, Results of rice cultivar tolerance tests to the rice water weevil in California, In: Proc. 16th Rice Tech. Working Group, Lake Charles, 1976, 63. 21. Heinrichs, E. A., Rice insects: the role of host plant resistance in integrated pest management systems, Korean J. Appl. Entomol., 31:256-275, 1992. 22. Heinrichs, E. A., Development of multiple pest resistant crop cultivars, J. Agric. Entomol. 11:225-253, 1994. 23. Heinrichs, E. A., Rice Insects of West Africa: A Review of the Literature, West Africa Rice Development Association, BP 2551, Bouaké, 1996. 24. Heinrichs, E. A. and A. A. Adesina, The contribution of multiple pest resistance to tropical crop production, In: B. R. Wiseman and J. A. Webster, Eds., Proc. 1994 Nelson Memorial Symp. on Host Plant Resistance, Thomas Say Pubs., Entomological Society of America, Lanham, in press. 25. Heinrichs, E. A., G. B. Aquino, S. L. Valencia, S. DeSagun and M. B. Arceo, Management of the brown planthopper, Nilaparvata lugens (Homoptera: Delphacidae), with early maturing rice cultivars, Environ. Entomol., 15:93-95, 1986. 26. Heinrichs, E. A., L. T. Fabellar, R. P. Basilio, Tu-Cheng Wen, and F. Medrano, Susceptibility of rice planthoppers Nilaparvata lugens and Sogatella furcifera (Homoptera: Delphacidae) to insecticides as influenced by level of resistance in the host plant, Environ. Entomol., 13:455-458, 1984. 27. Heinrichs, E. A. and F. G. Medrano, Influence of nitrogen fertilizer on the population development of brown planthopper, Int. Rice Res. Newsl., 10:20-21, 1985. 28. Heinrichs, E. A., F. G. Medrano, and H. R. Rapusas, Genetic Evaluation for Insect Resistance in Rice, International Rice Research Institute, Los Baños, 1985. 29. Heinrichs, E. A., F. G. Medrano, L. Sunio, H. Rapusas, A. Romena, C. Vega, V. Viajante, D. Centina, and I. Domingo, Resistance of IR cultivars to insect pests, Int. Rice Res. Newsl., 7:9-10, 1982. 30. Heinrichs, E. A. and P. K. Pathak, Resistance to the gall midge, Orseolia oryzae (Wood-Mason) in rice, Insect Sci. Applic., 1:121-132, 1981. 31. Heinrichs, E. A., H. R. Rapusas, G. B. Aquino, and F. Palis, Integration of host plant resistance and insecticides in the control of Nephotettix virescens (Homoptera: Cicadellidae), a vector of rice tungro virus, J. Econ. Entomol., 79:437-443, 1986. 32. Heinrichs, E. A., V. D. Viajante, and A. M. Romena, Resistance of wild rices, Oryza spp. to the whorl maggot Hydrellia philippina Ferino, Environ. Entomol., 14:404-407, 1985. 33. Hertford, R., Strengthening ecoregional research for agricultural development in the American lowland tropics: a framework and discussion paper, CIAT, Cali, 1994.

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34. IITA (International Institute of Tropical Agriculture), Cereal Improvement Program Rep. 1973, Ibadan, 1974. 35. IRRI (International Rice Research Institute), Facts About IRRI, Los Baños, 1990. 36. IRRI (International Rice Research Institute), IRRI Rice Facts, Los Baños, 1995. 37. Kartohardjono, A. and E. A. Heinrichs, Populations of the brown planthopper, Nilaparvata lugens (StDl) (Homoptera: Delphacidae), and its predators on rice cultivars with different levels of resistance, Environ. Entomol., 13:359-365, 1984. 38. Khan, M. Q., Control of paddy stem borer by cultural practices, In: The Major Insect Pests of the Rice Plant, Johns Hopkins Press, Baltimore, 1967, 369. 39. Khush, G. S., Multiple disease and insect resistance for increased yield stability in rice, In: L. R. Pollard, Ed., Progress in Irrigated Rice Research, IRRI, Manila, 1989, 79. 40. Khush, G. S., Modern cultivars--their real contribution to food supply and equity, Geojourn., 35:275-284, 1995. 41. Lee, S. C., Toward integrated pest management of rice in Korea, Korean J. Appl. Entomol., 31:205-240, 1992. 42. Matlon, P., M. Becker, M. Dingkuhn, E. Heinrichs, M. Jones, K. Miezan, K. Sahrawat, and A. Sy, Achievements in African rice research, Paper IRC:94/81(b), In: Eighteenth Session Int. Rice Commission, FAO, Rome, 1994. 43. Maxwell, F. G., Use of insect resistant plants in integrated pest management programmes, FAO Plant Prot. Bull., 39:139-146, 1991. 44. Medina, E. B., A. M. Romena, and E. A. Heinrichs, Screening wild rices for resistance to Marasmia patnalis Bradley, Int. Rice Res. Newsl., 11:12, 1986. 45. Myint, M. M., H. R. Rapusas, and E. A. Heinrichs, Integration of varietal resistance and predation for the management of Nephotettix virescens (Homoptera: Cicadellidae) populations on rice, Crop Prot., 5:259-265, 1986. 46. Nair, S., J. S. Bentur, U. P. Rao, and M. Mohan, DNA markers tightly linked to a gall midge resistance gene (Gm2) are potentially useful for marker-aided selection in rice breeding, Theor. Appl. Genetics, 91:68-73, 1995. 47. N’Guessan, F. K. and S. S. Quisenberry, Screening selected lines for resistance to the rice water weevil (Coleoptera: Curculionidae), Environ. Entomol., 23:665-675, 1994. 48. N’Guessan, F. K., S. S. Quisenberry, and T. P. Croughan, Evaluation of rice tissue culture lines for resistance to the rice water weevil (Coleoptera: Curculionidae), J. Econ. Entomol., 87:504-513, 1994. 49. N’Guessan, F. K., S. S. Quisenberry, and T. P. Croughan, Evaluation of rice anther culture lines for tolerance to the rice water weevil (Coleoptera: Curculionidae), Environ. Entomol., 23:331-336, 1994. 50. N’Guessan, F. K., S. S. Quisenberry, and S. D. Linscombe, Investigation of antixenosis and antibiosis as mechanisms of resistance in rice to the rice water weevil (Coleoptera: Curculionidae), J. Entomol. Sci., 29:259-263, 1994. 51. N’Guessan, F. K., S. S. Quisenberry, R. A. Thompson, and S. D. Linscombe, Assessment of Louisiana rice breeding lines for tolerance to the rice water weevil (Coleoptera: Curculiondae), J. Econ. Entomol., 87:476-481, 1994. 52. Ooi, P. A. C. and B. M. Shepard, Predators and parasitoids of rice insect pests, In: E. A. Heinrichs, Ed., Biology and Management of Rice Insects, Wiley Eastern, New Delhi, 1994, 585. 53. Pantoja, A., C. M. Smith, and J. F. Robinson, Evaluation of rice germ plasm for resistance to the fall armyworm (Lepidoptera: Noctuidae), J. Econ. Entomol., 79:1319-1323, 1986.

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54. Phuong, L. T., L. M. Chau, and M. B. Cohen, Resistance of cultivars derived from a wide cross with Oryza officinalis to the brown planthopper in the Mekong Delta, Vietnam, Int. Rice Res. Newsl., in press. 55. Rombach, M. C., D. W. Roberts, and R. M. Aguda, Pathogens of rice insects, In: E. A. Heinrichs, Ed., Biology and Management of Rice Insects, Wiley Eastern, New Delhi, 1994, 613. 56. Sanint, L. R., Summary of technology surpluses (1994-2010) associated with new rice technologies at CIAT, Rice Program Rep., CIAT, Cali, 1993. 57. Sauphanor, B., Résistance Variétale du riz aux Insectes. Rapport d’activités: 2ème Semestre 1981 et 1er Semestre 1982, Mimeo, IRAT, Bouaké, 1982. 58. Shih, H. P., The newly found rice water weevil (Lissorhoptrus oryzophilus Kuschel) on rice plant in Taiwan, Bull. Taoyuan Dist. Agric. Improve. Stn., 7:61-67, 1991. 59. Singh, J., G. S. Dhaliwal, and G. S. Sidhu, Advances in insect resistance in rice, In: G. S. Dhaliwal and V. K. Dilawari, Eds., Advances in Host Plant Resistance to Insects, Kalyani Publishers, New Delhi, 1993, 31. 60. Smith, C. M., The rice water weevil, Lissorhoptrus oryzophilus Kuschel, In: K. G. Singh, Ed., Exotic Plant Quarantine Pests and Procedures for Introduction of Plant Materials, ASEAN (PLANTI), Selangor, 1983. 61. Smith, C. M., Integration of rice insect control strategies and tactics, In: E. A. Heinrichs, Ed., Biology and Management of Rice Insects, Wiley Eastern, New Delhi, 1994, 681. 62. Smith, C. M. and J. F. Robinson, Evaluations of rice cultivars grown in North America for resistance to the rice water weevil, Environ. Entomol., 11:334-336, 1982. 63. Smith, C. M. and J. F. Robinson, Effect of rice cultivar height on infestation by the least skipper, Ancyloxypha numitor (F.) (Lepidoptera: Hesperiidae), Environ. Entomol., 12:967-969, 1983. 64. Sohn, J. K., S. K. Lee, J. C. Koh, H. Y. Kim, S. J. Yang, H. G. Hwang, D. Y. Hwang, and G. S. Chung, A new rice cultivar with multiple resistance to diseases and insect pests “Namyeongbyeo,” Res. Rep. Rural Development Administration (Crops), 29:18-28, 1987 (in Korean with English abstract). 65. Soto, P. E. and Z. Siddiqi, Screening for resistance to African rice insects, Paper presented at the WARDA varietal improvement seminar, West Africa Rice Development Association, Bouaké, 1976, 13. 66. Tsuzuki, H. and K. Isogawa, The initial occurrence of a new rice insect pest, the rice water weevil, in Aichi Prefecture in Japan, Shokubutsu-Boeki Plant Protection, 30:341, 1976, (in Japanese). 67. Ukwungwu, M. N., Susceptibility of Rice Cultivars to the Stem Borer, Chilo zacconius Bleszynski (Lepidoptera: Pyralidae), Ph.D. Dissertation, University of Ibadan, Ibadan, 1983. 68. Ukwungwu, M. N., M. D. Winslow, and V. T. John, Severe outbreak of rice gall midge (GM) in the Savannah zone, Nigeria, Intl. Rice Res. Newsl., 14:36-37, 1989. 69. Vercambre, B., Diopsis thoracica West (Dipt. Diopsidae), Importants ravageurs du riz en Afrique de l’Ouest. Données bio-ècologiques et application à la lutte intégrée, Agronomie Trop., 37:89-98, 1982. 70. Vos, P., R. Hoges, M. Bleeka, M. Reijans, T. van de Lee, M. Homes, A. Freijters, J. Pot, T. Peleman, M. Kaiper, and M. Zabeau, AFLP: a new concept for DNA fingerprinting, Nucleic Acids Res., 23:4407-4414, 1995. 71. WARDA (West Africa Rice Development Association), Rice Trends in Sub-Saharan Africa, WARDA, BP 2551, Bouaké, 1993.

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72. Way, M. J. and K. L. Heong, The role of biodiversity in the dynamics and management of insect pests of tropical irrigated rice — a review, Bull. Entomol. Res., 84:567587, 1994. 73. Weber, G. and O. Parada, Development of integrated pest management systems for rice in Latin America, In: E. A. Heinrichs, Ed., Biology and Management of Rice Insects, Wiley Eastern, New Delhi, 1994, 733. 74. Winslow, M. D., New breakthroughs and present accomplishments in rice research in Latin America and the Caribbean, Paper IRC:94/8-1(C), In: Eighteenth Session Int. Rice Commission, FAO, Rome, 1994. 75. Wu, Jung-Tsung, E. A. Heinrichs, and F. G. Medrano, Resistance of wild rices, Oryza spp. to the brown planthopper, Nilaparvata lugens (Homoptera: Delphacidae), Environ. Entomol., 15:648-653, 1986. 76. Xizhi, L. and C. X. Mao, Hybrid Rice in China — a Success Story, Asia-Pacific Association of Agricultural Research Institutions, FAO Regional Office, Bangkok, 1994.

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2

The Value of Conserved Wheat Germplasm Evaluated for Arthropod Resistance C. Michael Smith, Sharron S. Quisenberry, and Francois du Toit

CONTENTS I. Introduction A. Origin and Domestication of Wheat B. Importance as a Food Crop C. World Production II. Development of Arthropod-Resistant Wheat Germplasm A. Germplasm Resources B. Genetic Recombination and Selection C. Identification of Cereal Resistance Genes III. The Value of Wheat Resistance to Arthropods A. Economic Value B. Ecological and Humanitarian Values IV. Future Directions for Arthropod Resistance Research V. Conclusions Acknowledgments References

I. A. ORIGIN

AND

INTRODUCTION

DOMESTICATION

OF

WHEAT

The origin of Triticum spp. wheat traces back to antiquity in several different languages. The Chinese mai, the Sanskrit sumara and godhuma, the Hebrew chittah, and the Egyptian br are all synonymous of the word wheat.26 Prehistoric wheat has been unearthed in Aggtelek, Hungary, where it is believed to have been grown and consumed during the Stone Age.

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Archeological records indicate that T. monococcum L. and T. turgidum L., both ancestral species of modern bread wheat (see below), were domesticated in the Near East 9,000 years ago.126 Triticum turgidum dicoccum Schrank, another ancestral wheat species, was grown in the Near East as long as 9,500 years ago.164 Approximately 6,000 years ago, some tetraploid and diploid wheats hybridized to form currently grown hexaploid wheats (see below). This occurrence greatly broadened the growing range of wheat.164 The Chinese cultivated wheat approximately 5,000 years ago, viewing it as a gift from heaven. The historian Berosus is quoted as saying that wheat grew wild in the valleys between the Tigrus and Euphrates Rivers, the cradle of modern civilization.26 Cultivated wheats now exist in an allopolyploid series, ranging from diploid to hexaploid, in the A, B, and D genomes (sets of seven chromosomes). The analyses of these genomes pioneered by Kihara73 have been used extensively to develop a knowledge of the evolution and phylogenetic relationships among members and relatives of the tribe Triticeae. Diploid wheat, T. monococcum L. var. urartu (2N=2X=14, AA), is the probable origin of the A genome in polyploid wheats.84 The source of the B genome, although not completely defined, is thought to be T. speltoides (Tausch) Gren. Ex Richter (2N=2X=14, BB).84 The source of the D genome in polyploid wheat is T. tauschii (Coss.) Schmal. (2N=2X=14, DD).64 The development of modern bread wheat occurred as a result of at least two major evolutionary events. The first was the creation of the tetraploid wheat Triticum turgidum L. var. durum (2N=4X=28, AABB), which resulted from a cross between T. monococcum and an unknown B genome donor. Subsequently, a cross between T. turgidum (2N=4X=28, AABB) and T. tauschii (2N=2X=14, DD) created what is now common bread wheat, Triticum aestivum L. (2N=6X=42, AABBDD). The majority of wild and cultivated wheats are genetically either AA, AABB, or AABBDD, and can be classified as T. monococcum (AA), T. turgidum (AABB), or T. aestivum (AABBDD). The chromosomes of hexaploid wheat exist in seven homeologous groups. Each group has three pairs of chromosomes, one each from the A, B, and D genomes.

B. IMPORTANCE AS

A

FOOD CROP

Triticum aestivum is a cool-season cereal crop, but it is grown across the widest range of environments of any cereal crop in the world.10 Wheat is the most important cereal crop in the world as it provides more nourishment for the world’s population than any other source of nutrition.61 By the mid-1990s, wheat had become the food source of choice by the majority of the world’s population. Wheat- and grain-based diets are continually shown to be higher in fiber and lower in fats than meat-based diets. Wheat has been a major food crop in many countries for several hundred years, and it is now a competitor with rice and maize as the dominant food crop of the developing world. From 1961 to 1992, wheat experienced a 4.6% increase in use as a food crop among developing countries, compared with a 4.3% increase for maize and a 3.0% increase for rice. Per capita consumption of wheat in developing countries

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rose from 71 kg per year from 1987 to 1989 to 75 kg per year from 1990 to 1992. Wheat use in all countries increased by 3.4% from 1993 to 1994, compared with 3.0% for maize and 2.8% for rice during the same period.29 Wheat is also used as an animal food. About one-fourth of the post-milling grain weight of wheat, which includes the bran and germ, is utilized as human and livestock food supplements. In the vegetative stage, wheat foliage can be directly grazed by livestock. Some low-grade wheat is fermented for alcohol production. Wheat gluten, a component that determines the quality of the flour obtained from the milling of wheat grains, is used to increase the total protein of baked goods, food products, and low-protein flours. Wheat starch is used for laundering, adhesives, wallpaper, and textiles.10

C. WORLD PRODUCTION More land is devoted worldwide to the production of wheat than any other commercial crop. A wheat crop is harvested somewhere in the world in every month of the year, but most harvests occur from April to September in the temperate regions of the world.9 Until the dissolution of the former Soviet Union and the formation of the European Economic Union, the USSR was the world’s leading wheat producer, with the U.S. and the People’s Republic of China ranking second and third, respectively. The People’s Republic of China is the current leading wheat producer, with a 1994/1995 harvest of 106.4 mmt.4 The European Economic Union (82.1 mmt) and the U.S. (63.2 mmt) rank second and third, respectively, in world production. The two other major world wheat producing countries are the former Soviet Union (60.1 mmt) and India (56.8 mmt).4 Because of these production rates, the continent of Asia leads world wheat production at 281.2 mmt, followed by Europe (119.9 mmt), North America (90.3 mmt), Oceania (18.1 mmt), South America (15.9 mmt), and Africa (13.1 mmt). The major world wheat exporting countries are the U.S., which exported 52% (33.1 mmt) of its 1994 wheat crop and the European Economic Union, which exported 20.1 mmt of its 1994 crop (24% of its production). Canadian producers exported 81% (18.7 mmt) and Australia producers 71% (12.8 mmt) of their 1994 wheat production.4 The principal world wheat importer is the former Soviet Union, which purchased 13.5 mmt of wheat and wheat flour in 1994.4 Other significant wheat importing countries include Japan (6.1 mmt), Egypt and Korea (5.9 mmt each), Brazil (5.8 mmt), and the People’s Republic of China (4.3 mmt). The amount of wheat and wheat flour imported by Brazil, Egypt, Japan, and Korea remained relatively constant from 1991 to 1994, while importation dropped significantly in the People’s Republic of China (73%) and the former Soviet Union (43%). Annual increases in world wheat production have fluctuated between 1.5 and 3.0% during the past 40 years, as a result of changes in human and livestock food consumption patterns, increases in the standard of living in developing countries, and the effects of abiotic and biotic stresses.115 One of the major biotic stresses affecting cultivated wheat is the feeding damage from several arthropod pests in

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different production zones throughout the world. In this chapter, we review and discuss the germplasm resources available to improve wheat for arthropod resistance, the resistance genes that have been identified and used, the value of resistance genes, and the future directions for research to improve wheat with arthropod resistance.

II.

DEVELOPMENT OF ARTHROPOD-RESISTANT WHEAT GERMPLASM

A. GERMPLASM RESOURCES Ayad et al.6 compiled information on the 25 major collections of Triticeae germplasm throughout the world. These collections are maintained by state, national, and international institutions. Other major collections of wheat germplasm are maintained in Australia, Germany, Israel, Italy, and Japan. The largest Triticum collections are stored and maintained at the International Maize and Wheat Improvement Center (CIMMYT), El Battan, Mexico (approximately 98,905 accessions),127 the Vavilov All-Union Institute of Plant Industry, Leningrad, Russia, (approximately 70,000 accessions),6 and the U.S. Department of Agriculture, Agricultural Research Service (USDA-ARS), National Small Grains Collection, Aberdeen, Idaho (45,997 accessions).38 The International Plant Genetic Resources Institute, Rome, Italy (formerly International Board of Plant Genetic Resources), in conjunction with the International Center for Agricultural Research in Dry Areas, initiated a database of the number, location, and condition of all existing wild Triticeae accessions.56 Over 22,000 unique accessions are maintained in 33 different countries and in an additional 25 smaller collections. A special collection is the Kansas State University Wheat Genetic Resource Center Collection of approximately 5,000 accessions of Triticeae. The species identity and country of origin are known for nearly all accessions in these collections but, in many cases, precise data on altitude of collection site and ecogeographic characteristics are not known. Only a fraction of these accessions have been evaluated for more than one arthropod pest.

B. GENETIC RECOMBINATION

AND

SELECTION

Knowledge of the genetics of arthropod resistance in wheat is facilitated by exploiting the aneuploid condition in Triticum species. Aneuploidy is an increase or decrease in the plant chromosome number from the normal 2N euploid chromosome number. The change in chromosome number may involve a single chromosome arm (telosome) change or a change in more than one chromosome. Hexaploid wheat exists in several different types of aneuploid configurations, including nullisomic (2N=40), monosomic (2N=41), disomic (2N=42), trisomic (2N=43), tetrasomic (2N=44), and a range of monosomic through tetrasomic combinations with varying numbers of chromosome arms.62 Although the generation of trisomic and tetrasomic stocks of wheat is possible, di-telosomic stocks and nullisomics with compensating tetrasomics (nuilli-tetra) stocks are used more commonly than tri- or tetrasomic stocks because of their greater vigor than nullisomics and improved specificity over monosomic stocks. Nevertheless, monosomic spring wheat

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cultivars susceptible to the Hessian fly, Mayetiola destructor (Say), have been crossed to a disomic line to determine the chromosome(s) responsible for fly resistance.32 Monosomics produced with the “Chinese Spring” wheat cultivar have also been used to determine the chromosome location of genes for resistance to the greenbug Schizaphis graminum (Rondani), and to the Russian wheat aphid Diuraphis noxia (Mordvilko).58,122 The seven homologous sets of hexaploid wheat chromosomes have similar gene contents and can either replace or compensate for each other in combinations ranging from nullisomic to tetrasomic. Translocations involving the homologous wheat chromosome with foreign chromosomes, chromosome arms, or chromosome segments can be useful breeding events in wheat improvement. Success in moving genes from related foreign species into common wheat depends on the evolutionary distance between the two species involved.30 Species in the primary gene pool of common wheat (T. turgidum, T. monococcum, T. tauschii) share homologous chromosomes. The secondary gene pool includes polyploid Aegilops/Triticum species with at least one genome (a set of 7 chromosomes) homologous to T. aestivum or T. durum. This group includes T. timphophevii Zhuk. and species belonging to the Aegilops section Sitopsis.30 The tertiary pool consists of relatives with only a distant association whose chromosomes are not homologous to those of wheat. Meiosis centric-breakage-fusion techniques, ionizing radiation, and induced homologous recombination have been used to move chromosome arms or chromatin from tertiary gene pool donors into wheat.30 Arthropod resistance genes have been transferred from species in the primary and secondary gene pools following crossing and homologous recombination with chromosomes of agronomically superior common wheat cultivars. Backcross breeding is commonly used to select for progeny with both resistance and improved agronomic traits. Backcross breeding involves crossing the resistant progeny from each previous generation back to the same agronomically desirable parent wheat cultivar. Selection is made after each generation for arthropod resistant progeny with superior agronomic performance. Backcross breeding has been used to incorporate arthropod and disease resistance into agronomically superior wheats that were susceptible to these pests.

C. IDENTIFICATION

OF

CEREAL RESISTANCE GENES

The development and use of arthropod-resistant cultivars has long been recognized as a sound approach to wheat protection. One of the earliest uses of arthropod resistance in the U.S. was when resistant wheat cultivars were grown to protect crops in New York from infestations of the Hessian fly in 1788. Today, arthropod-resistant wheat cultivars are grown in Africa, Asia, Europe, and the U.S. Sitobion avenae — Metopolophium dirhodum — Rhopalosiphum padi Complex Resistance. Numerous sources of resistance to one or more of the aphids in the S. avenae (F.)-M. dirhodum (Walker)-R. padi (L.) complex have been identified by European researchers. Research in England identified resistance to S. avenae in the T. aestivum cultivars “Kador” and “Amigo,” and resistance to both S. avenae and M. dirhodum in the T. dicoccum cultivar “Emmer,” and in the T. monococcum cultivar

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TABLE 1 Wheat Genes Governing Resistance to Biotypes of Schizaphis graminum (Rondani) in North America Reaction to aphid biotype Source “Dickinson” “Amigo” “Largo” CI17959 CI17882 GRS 1201

Origin

Gene

B

C

E

F

G

H

I

Triticum turgidium durum Secale cereale T. tauschii T. tauschii T. speltoides S. cereale

gb1 Gb2 Gb3 Gb4 Gb5 Gb6

s r s s s r

s r r r r r

s s r r r r

r s s s s s

s s s s s r

s s r s s s

— — r r — r

r = resistant; s = susceptible.

“Einkorn.”74,136 In the Czech Republic, resistance to all three aphids was identified in “Mironovskaya 808.”52-54 In Poland, “Asta,” “Grava,” and “Sago” are resistant to S. avenae.72 In France, resistance to S. avenae was confirmed in “Mirela,” “Vala,” “Downy,” and “Kornay,” and in T. monococcum.12,19 In Hungary, several cultivars and the U.S. cultivar “Downy” are resistant to R. padi.98 Canadian research on R. padi as a vector of barley yellow dwarf mosaic virus led to the identification of a T. aestivum x Agropyron repens L. Beauv. hybrid with resistance to R. padi.146 Schizaphis graminum Resistance. Six genes have been identified with resistance to biotypes of S. graminum (Table 1). There is one recessive gene, gb1,18 and five dominant genes, designated Gb 2-6.105,148 The first S. graminum biotype, B, was named because of its ability to overcome the resistance of “Dickinson 28A,” a durum wheat that possesses gb1 and resistance to biotype A.163 Biotype B also has the ability to tolerate higher temperatures than biotype A.128 In the late 1960s, S. graminum populations became severely damaging in sorghum, Sorghum bicolor (L.) Moench, producing areas of the southwestern U.S. and the sorghum biotype C developed.40 “Amigo” wheat, which possess the Gb2 gene from rye, Secale cereale L., is resistant to biotypes B and C.123 Increased insecticide usage on sorghum in Texas and Oklahoma caused the development of biotype D which is resistant to the insecticide disulfoton.102,143 During the past 20 years, several additional biotypes of S. graminum have developed. Biotype E, capable of feeding on biotype C resistant “Amigo” wheat, developed in the U.S. high plains of Texas during the 1970s. Resistance to biotypes C and E was documented in the amphiploids T. turgidum/T. tauschii “Largo” (Gb3) and T. durum/T. tauschii CI17959 (Gb4) which also possess resistance to wheat streak mosaic virus.106,149 Specific resistance to biotype E (Gb5) was identified in CI17882, whose lineage involves T. speltoides, the presumed source of resistance.148 Biotype E greenbugs exhibit normal feeding behavior and fecundity on wheat cultivars resistant to biotype B.87 Because biotype E greenbugs have comparatively higher digestive enzyme activity, they are physiologically superior to biotype C.13 Biotype F differs from biotype E in its ability to kill Canada bluegrass, Poa compressa

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TABLE 2 Development of Diuraphis noxia-Resistant Wheat in North America and South Africa, 1986–1996 Source PI137739 PI262660 SQ24 PI372129 PI294994 29 Pls PI149898 PI243781 a

Origin Iran Russia Triticum tauschii Russia Bulgaria Afghanistan, Iran, Russia Russia —

Genes Dn1 Dn2 dn3 Dn4 Dn5 ? —a Dn6

Ref. 20-22, 129 20-22, 129 89 89, 119 76 103 8 119

Resistance controlled by at least two undesignated genes.

L., and it lacks a dorsal stripe. Biotype F is also susceptible to the gb1 gene in “Dickinson 28A” wheat.65 Biotypes G and H differ in their reactions to small grain hosts.107 Biotype G survives on all known sources of resistant wheat, but is avirulent to a recently discovered source of resistance (GRS 1201), a T. aestivum /S. cereale hybrid with the Gb6 gene.104 Biotype H has the same effect on wheat cultivars as biotype E, but is avirulent to the Gb3 gene of “Largo” wheat. Biotypes I and K, new biotypes virulent to S. graminum-resistant sorghums, were detected in Kansas,41,46 and are avirulent on sources of Gb3, Gb4, and Gb6.105 Chapter 4 of this volume is a source of information on S. graminum biotypes in relation to the development of S. graminum-resistant sorghum. Diuraphis noxia Resistance. Six genes have been identified in different wheats for resistance to D. noxia (Table 2). The dominant genes Dn1 and Dn2 were identified in the hexaploid wheat plant introduction (PI) lines 137739 (Iran) and 262660 (Russia) in South Africa,20-22 and confirmed in the U.S.130 The recessive gene dn3 originates from the T. tauschii accession SQ24, a parent in the amphiploid wheat derived from crosses between T. tauschii and T. turgidum.89 Two additional dominant D. noxia resistance genes, Dn4 and Dn6, originate from a Russian T. aestivum (PI372129) and an Iranian T. aestivum (PI243781).90,119 The resistance of a Bulgarian T. aestivum (PI294994) to D. noxia is controlled by the dominant gene Dn5.20,23,78 Twenty-nine PI lines from Afghanistan, Iran, and Russia exhibited moderate to high levels of D. noxia resistance during an evaluation of more than 12,000 wheat accessions from the USDA-ARS National Small Grains Collection.103 One of the lines (PI14898) has been crossed with the susceptible hard red spring wheat “Bobwhite” to produce the germplasms “STARS-9302W” and “STARS-9303W,” which are both moderately resistant to D. noxia.8 Resistance in these germplasm releases appears to be controlled by two genes. Wheat cultivars with D. noxia resistance were grown for the first time in 1996 in the U.S. in Colorado and in the South African Free State province. The South African wheats possess the Dn1 and Dn2 genes and the Colorado cultivar “Halt” © 1999 by CRC Press LLC

contains the Dn4 gene. Additional U.S. cultivars are being bred from germplasm developed in Idaho, which also contain the Dn1 gene from PI137739 and the Dn2 gene from PI262660.137,138 Mayetiola destructor Resistance. The development of M. destructor biotypes has closely paralleled the development of Hessian fly resistant-wheat cultivars, which began in New York in the early 1900s. Hessian fly biotypes were first recognized in 1930.95 Since then, 27 different genes for M. destructor resistance have been identified in common wheat, durum wheat, T. turgidum L. var. durum, goatgrass, T. tauschii, and in S. cereale (Table 3). Genes H1-H5, the H7H8 combination, and H12 are from T. aestivum,33,93,101 while genes H6, H9-H11, H14-H20, and H27 are from T. turgidum.1,15,76,77,91,92,94,99,111,112,125,139-141 Genes H13, H22-24, and H26 are from T. tauschii,17,48,49 and genes H21 and H25 are from S. cereale.50,114 This information has been summarized by Ratcliffe and Hatchett.110 Sixteen M. destructor biotypes have been identified in field populations, and are differentiated by how various combinations of genes for virulence (the ability to overcome cereal resistance) interact with genes for resistance to M. destructor in different resistance sources.113 Virulence in an M. destructor biotype depends on the existence of the homozygous recessive condition in the virulence gene of the fly at a locus corresponding to a specific dominance gene for resistance.31 Thus, in each instance of cultivar resistance (fly avirulence), the particular fly biotype in question lacks the homozygous recessive condition at the virulence gene locus. High temperature diminishes the expression of resistance genes to some M. destructor biotypes, causing them to appear only moderately resistant.76,77,91,109,133,135,140,141,147 An alternative system of biotype nomenclature, based on fly reaction to sets of different resistance genes has been proposed,100 but this system has yet to be put into practice. The U.S. Great Plains biotype, isolated in western Kansas, carries the homozygous recessive condition for virulence only in association with “Turkey,” and lacks this condition when attempting to survive on cultivars with the H3, H5, H6, H7H8 gene combination, and H9-H19.112 Biotype A (Table 3) is similar to the Great Plains biotype, but is homozygous recessive for virulence on “Seneca” (H7H8 gene combination). Biotype B possesses an additional virulence gene capable of breaking the resistance of the H3 gene in “Monon.” Biotype C is now found infrequently in the north central U.S., but has the opposite reaction of biotype B to the H3 and H6 genes in “Monon” and “Knox 62” wheat, respectively. Biotype D, found commonly in the mid-Atlantic and northeastern U.S., is similar to biotype B, but is also virulent to the H6 gene of “Knox 62.”110,112,113 Biotype E in Georgia47 is similar to biotype C, and is nonvirulent to the H7H8 gene combination. Biotypes F and G are present in the south and southeastern U.S. and are also nonvirulent to the H7H8 gene combination.110,112,113 Biotypes J and L developed in response to the resistance from the H5 gene in “Arthur 71” are found in field populations in Indiana.110,112,113,132,134 Biotype L is now widely distributed in the midwest, mid-south, mid-Atlantic, and southeastern regions of the U.S. The H6 gene is resistant to biotype J. Numerous biotype L resistance genes have been identified in the H9, H10, and H12-27 genes.112 Current major U.S. wheat breeding efforts for M. destructor resistance are located at the University of Florida, University of Georgia, Kansas State University, and

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TABLE 3 Interaction of Cereal Genes for Resistance to M. destructor with Virulence Genes in M. destructor Biotypes Resistance gene(s), source, and cultivar, PI, or CI name None, Triticum aestivum, “Turkey” H1&H2, T. aestivum, “Dawson,” CI3342 H3, T. aestivum, Ill. #1, W-38 sel.7, CI12061 H4, T. aestivum, “Java,” CI10051 H5, T. aestivum, “Ribeiro,” PI56206-8 H6, Triticum turgidum, PI94587 H7H8 combination, T. aestivum, “Seneca” & CI12529 H9, T. turgidum, CI17714 H10, T. turgidum, CI17714 H11, T. turgidum, PI94587 H12, T. aestivum, “Lusco” H13, Triticum tauschii, HU2076 H14 & H15, T. turgidum, CI1764 H16, T. turgidum, PI94587 H17, T. turgidum, PI428435 H18, T. turgidum, CI6887 H19, T. turgidum, PI422297 H20, T. turgidum, “Jori” H21, Secale cereale, “Chaupon” H22, T. tauschii, TA1644 H23, T. tauschii, TA1642 H24, T. tauschii, TA2452 H25, S. cereale, “Chaupon” H26, T. tauschii, TA2473 H27, T. turgidum, PI422297 a

Biotype GP

A

B

C

D

E

J

L

Sa R R ? R R R

S S R ? R R S

S S S ? R R S

S S R ? R S S

S S MR ? R S S

S S S ? S R S

S R S ? S S R

S S S ? S S S

R R R R — — — — R R — — — — — — — —

R R R R — — — — R R — — — — — — — —

R R R R R R R — R R — — — — — — — —

R MR R MR R R R — R R R — — — — — — —

R R R R R R R R R R — — R R R — R —

R — R — — R R — R — — — — — — — — —

— — S — — S — — — — ? — — — — — — —

R R S MR R R R R R R — R R R R R R R

R = resistant; MR = moderately resistant; S = susceptible.

Modified from Patterson, F.L. et al., Proposed system of nomenclature for biotypes of Hessian fly (Diptera: Cecidomyiidae) in North America, J. Econ. Entomol., 85:307-311, 1992; and Ratcliffe, R.H. and J.H. Hatchett, Biology and genetics of the Hessian fly and resistance in wheat, In: New Developments in Entomology, K. Bobdari, Ed., Research Signpost, Scientific Information Guild, Trivandrum, 1997, 47. (Refs. 100 and 110)

Purdue University in conjunction with USDA-ARS scientists. The primary focus of these efforts is resistance to biotypes E and L.112 Cephus cinctus Resistance. Since 1946, 11 spring wheat cultivars with resistance to C. cinctus have been developed by North American scientists.158 Many of the first cultivars had useful levels of sawfly resistance but lacked adequate yield levels and baking qualities. Currently grown spring wheats with sawfly resistance have medium to high levels of yield, good baking quality, and disease resistance. © 1999 by CRC Press LLC

Winter wheats have recently been identified with useful levels of C. cinctus resistance.14,85 In Syria, several bread and durum wheat cultivars express resistance to C. pygmeus and Trachelus spp.83 Reduced light intensity decreases the expression of the solid stem character in resistant cultivars.116 Under field growing conditions, reduced natural light intensity, resulting from prolonged periods of cloud cover, can inhibit the full expression of the stem solidness character in some resistant cultivars. This occurrence has slowed the acceptance of some resistant cultivars by producers. Oulema melanoplus Resistance. Numerous sources of resistance to O. melanoplus have been identified in wheat. The original two sources of resistance from Russia, CI8515 and CI9321,153,157 were used as parents to develop “Downy”155 which is resistant to O. melanoplus in Hungary.97 Several additional sources of resistance to O. melanoplus also have been identified in Hungary.66,67,97 Some wheat cultivars with dense growth of long, erect trichomes deter oviposition by O. melanoplus, much more than cultivars with sparse growth of short trichomes.59,150 Eggs deposited on the trichome “field” rising above the leaf surface suffer mortality due to desiccation and puncture by trichomes.160 Larvae also die from punctures of the alimentary canal sustained after ingestion of trichome fragments.159 Some pubescent wheats also exhibit antixenosis to adults and larvae of M. destructor and R. padi.117,118 Aceria tosichilla Resistance. Resistance to A. tosichilla in cereals was first demonstrated in Triticum x Agropyron hybrids.2 The gene from Agropyron is expressed as a dominant trait and was later designated as Cmc2.162 Mite resistance from a single gene (Cmc1) from T. tauschii, also inherited as a dominant trait, was incorporated into wheat.144 Several wheats with translocations from S. cereale also effectively reduce the incidence of wheat streak mosaic virus and cultivars with this resistance are commercially available.80,81 Research in Kansas45 (Table 4) determined that sources of resistance with the Cmc1 and Cmc2 genes are resistant to A. tosichilla populations from South Dakota or Alberta, Canada, but susceptible to A. tosichilla from Kansas and Texas. Wheats with resistance originating from S. cereale are resistant to A. tosichilla from all locations, but are susceptible to A. tosichilla from Kansas adapted to and reared on the normally resistant “TAM-107”. A common wheat with D. noxia resistance (PI222655) is resistant to all A. tosichilla populations except the one from Alberta, Canada (Table 4). Resistance to A. tosichilla has also been detected in the D. noxia-resistant wheats CI9355, PI221699, PI222651, PI222661, PI222679, PI222680, and PI222682,43 and in a chromosome addition hybrid (Add 6V-1) and a chromosome substitution hybrid (Sub 6V-1) arising from wheat and the annual grass Haynaldia villosa (L.) Schur.16 In contrast to the effects of pubescence on O. melanoplus and R. padi, pubescent wheats are susceptible to infestation by A. tosichilla.42

III.

THE VALUE OF WHEAT RESISTANCE TO ARTHROPODS

A. ECONOMIC VALUE The economic benefits resulting from the production of arthropod-resistant wheat cultivars are quite significant in production agriculture. Wheat resistance research

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TABLE 4 Interactions Between Genes in North American Wheat Germplasm and Virulence Genes in Aceria tosichilla Mite colony collection site Plant source Aegilops squarrosa Agropyron elongatum Secale cereale (PI475772) S. cereale (TAM-107) Triticum aestivum (PI222655)c T. aestivum (“Tomahawk”) a b c

Kansas

Kansasa

Texas

South Dakota

Alberta, Canada

Sb S R R R S

S S S S R S

S S R R R S

R R R R R S

R R R R S S

Kansas mite colony reared on TAM-107 extensively before bioassay. Susceptibility and resistance based on numbers of mites per plant. Resistant to D. noxia.

From Harvey, T.L. et al., Survival of five wheat curl mite, Aceria tosichilla Keifer (Acari: Erophyidae), Strains on mite resistant wheat, Exp. Appl. Acarol., 19:459-463, 1995. With permission.

provides a substantially greater return for each dollar invested in development, compared with research to develop chemical insecticides or miticides. In the late 1960s, several U.S. wheat cultivars developed with resistance to M. destructor returned approximately $600 on each U.S. dollar invested in cultivar research, compared with a $5 return on each dollar spent on the development of insecticides during the same period, a 120-fold greater return on investment.96 The following section documents the economic benefit of varietal resistance in wheat to several arthropod pests. S. avenae-M. dirhodum-R. padi Complex. The aphids R. padi, S. avenae, and M. dirhodum cause significant wheat yield reductions as they inject salivary enzymes into plants and remove nutrients during feeding. Sitobion avenae is considered the major aphid pest of small cereal grains in central and western Europe,12,55,121 whereas R. padi is considered the cereal aphid of greatest economic significance in Canada, southeastern Europe, and Scandinavia.69,98,146 In the U.S., wheat seedlings infested with R. padi and S. avenae suffer 57 and 13% reductions in kernel weight, respectively.63 Resistance to these aphids is yet to be incorporated into European wheat production on a large scale, and economic data on the value of the resistance identified is unavailable. Schizaphis graminum. Greenbug damage to wheat is well documented in Brazil, Chile, Pakistan, and the U.S.39,51,108,145 Numerous severe S. graminum outbreaks have occurred on wheat, barley Hordeum vulgare L., oats Avena sativa L., and sorghum in several U.S. production areas since 1949. The value of yield losses in U.S. wheat production related to S. graminum infestation and feeding from 1951 to 1960 was estimated at 1.4% of the total crop.70 Based on a 1993 wheat crop value of $7.6 billion © 1999 by CRC Press LLC

(U.S.),4 this loss would exceed $106 million. The incorporation of the multibiotype S. graminum-resistant germplasm GRS1201 into locally adapted cultivars should be worth over $20 million annually in the states of Kansas, Texas, and Oklahoma alone.154 Diuraphis noxia. Russian wheat aphid reduces wheat yield by direct feeding damage, but is an inconsequential vector of wheat diseases. Damaged wheat plants react to injection of D. noxia saliva by rolling the leaves longitudinally around the main leaf vein to form a tubular refuge for feeding aphids. In addition to removal of plant sap that weakens plants and lowers grain yield, prolonged infestations cause emerging panicles to be partially trapped in the leaf sheath, resulting in a “goose neck”-shaped panicle and substantially reduced grain yields.151 Diuraphis noxia is a primary pest of wheat in Mexico, South Africa, and Lesotho, and in the northwestern Great Plains of the U.S.35,86,151 With its rapid spread into several previously uninfested areas of the world, D. noxia has caused significant economic losses to wheat production. The cumulative losses to all U.S. small grain production for D. noxia control, yield losses, and lost community economic activity from 1986 to 1993 was valued at $850 million.71 Insecticide applications were reduced by approximately 50% when “Halt,” the first U.S. D. noxia-resistant wheat cultivar, was produced in Colorado during 1996 on approximately 1.13 million ha. This resulted in a savings to wheat producers of approximately $14 million.165 In the Free State province of South Africa, D. noxia-related yield losses of approximately 5% of the annual crop area (525,000 ha) are approximately $6.8 million (U.S.), and insecticidal control of the aphid ($13.30 per ha) costs producers an additional $7 million, for a total of $13.8 million for annual D. noxia control. In 1996, approximately 9.6% of the total crop was planted to resistant cultivars. Thus, the estimated value of D. noxia resistance in 1996 was 9.6% of the total $13.8 million control costs, or $1.33 million. Six resistant cultivars have been released for production thus far, but five of these have been marketed for only 1 to 2 years. The area planted to D. noxia-resistant cultivars is expected to increase dramatically during the next 5 years, and should exceed 50% of the total wheat area by the year 2000. Use of resistant cultivars should increase the annual value of D. noxia resistance to nearly $7 million.166 Mayetiola destructor. Hessian fly is distributed throughout North America, Europe, North Africa, and Russia.27,28 The economic importance of this insect is related to larval feeding damage between the plant leaf sheath and plant stem, which stunts wheat seedling growth and causes plant death or reduces the grain yield in surviving plants. Larval feeding affects approximately 50% of the U.S. crop every year, causing about $100 million in damage.70,113 Damage from M. destructor feeding on wheat in various states in the U.S. varies between years and geographic locations. In Georgia in 1989, damage was estimated to be $28 million,60 while yield losses in Texas in 1984 were estimated at over $5 million.57 As mentioned previously, extensive research in the U.S. on the gene for gene interaction between M. destructor and fly-resistant cereal germplasm has led to the development and release of many fly-resistant wheats since the mid 1960s. These cultivars were developed because of the pervasive distribution of M. destructor on wheat in the soft wheat region of the eastern U.S. Hessian fly damage claimed 1.5% © 1999 by CRC Press LLC

of the annual U.S. wheat crop from 1951 to 1960, a loss of approximately $114 million, based on the 1993 wheat crop value.4 The current value of M. destructor resistance in wheat compensates for much of the loss estimate, because of the positive effects that resistant cultivars have had during the past 25 years. Between 1950 and 1983, 60 different M. destructor-resistant cultivars were released.51 By 1984, over 10 million ha of fly-resistant wheats were planted in 30 different states in the U.S.124 The value of M. destructor resistance in the southeastern U.S. is beginning to be recognized. Survey and loss data developed by the Georgia Agricultural Experiment Station documented that the use of M. destructor-resistant cultivars from 1990 to 1994 decreased fly damage 69% compared with losses incurred from 1985 to 1989. Conversely, wheat yields increased by 24% from 1985 to 1994 as the percentage of resistant cultivars increased from 5 to 90%. The approximate value of M. destructor resistance in Georgia, caused by reduced yield losses and insecticide applications, is approximately $7 million.11 In South Carolina, annual M. destructor management costs have declined from $12 million to $200,000 since the establishment of an M. destructor management program which uses resistant cultivars.154 Of the cultivars grown in the north central and southeastern U.S. during the past decade, 60 to 80% have possessed some level of resistance to M. destructor. Had these resistance genes been highly durable, their value would have been worth more than $50 million annually. However, biotype L now dominates that area of U.S. wheat production, and is virulent to all of the M. destructor resistance genes in commercial wheats grown there. Until cultivars with biotype L resistance are grown on a large scale in north central and southeastern U.S., yield losses due to M. destructor feeding may reoccur, especially when fly populations reach economically damaging levels. Mayetiola destructor severely reduced wheat production in Morocco during the late 1980s.27 Since then, a 14-year Morocco–U.S. cooperative research effort has led to the development and release of 32 lines of bread wheat with M. destructor resistance. Of these, 27 are Moroccan bread wheats. The benefit to cost ratio of this research has been estimated at 9:1.7 Thus, every Moroccan dirham invested in this research yielded an average return of 9 dirhams for society. Cephus cinctus. Wheat stem sawfly occurs in the northern Great Plains, damaging wheat in the U.S. states of North Dakota and Montana and the Canadian provinces of Alberta, Manitoba, and Saskatchewan.158 Grain yield is reduced from the effects of larvae feeding inside the plant stem during development, which restricts nutrient flow to grain heads. Larval feeding also weakens the stems and causes them to lodge, which further reduces grain yield. Estimated C. cinctus-related yield losses to wheat in North Dakota and Montana exceed $10 million in some years.70 In 1989, a severe infestation in central Montana caused estimated wheat yield losses of 80%. Similar losses have been reported in the Canadian provinces affected by C. cinctus. The European wheat stem sawfly, C. pygmeus (L.), and two related sawflies Trachelus judaicus (Kônow) and T. libanensis (André), frequently damage cereal grains in North Africa and western Asia.82 Although no current estimates are available, the annual value of C. cinctus resistance in Montana in 1948 was estimated at $3.8 million.3 If conservative (3.5%) annual inflationary estimates are used to amplify this figure,

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the current annual value of C. cinctus resistance is greater than $15 million in Montana. Oulema melanoplus. Cereal leaf beetle damages wheat and other small grains by skeletoning leaves, which reduces the grain yield of infested plants.161 The range of this insect extends throughout Europe, into the Caucasian area of Russia and into western North Africa and the Middle East.152 This insect was first noted in the U.S. in Michigan wheat plantings in 1959, and has since extended its range into wheatproducing states of the eastern and central U.S. An O. melanoplus population has recently been detected much farther west, in Idaho.167 Populations of O. melanoplus in the U.S. are currently below economic damage levels and widespread population outbreaks have not occurred in several years.154 However, wheat grain losses resulting from past O. melanoplus infestations have approached 55% in spring wheat156 and 23% in winter wheat.34 In field studies comparing resistant and susceptible cultivars under O. melanoplus infestation, yield losses of susceptible cultivars ranged from 17 to 25%, while the yield of resistant cultivars was not reduced.155 Several parasitic insects have played a major role in reducing O. melanoplus populations since the resistant cultivars were released,68 and there have been no major efforts to demonstrate the economic value of O. melanoplus resistance. Aceria tosichilla. Aceria toschilla is a native pest of North American grasslands and causes major annual losses in wheat yields. Mites feed and lay eggs on the upper leaf surfaces of wheat plants. As the mite infestation increases, damaged leaves curl toward the midvein, often preventing new leaves or grain heads from emerging and elongating. This mite is important because it is the primary vector for wheat streak mosaic virus (WSMV),129 an important disease of wheat in Europe, North America, and Asia, which periodically causes severe yield losses.131 Estimates of yield losses from WSMV infection during severe virus outbreaks range from 37% in Canada to 13% in Kansas.5,44,79 The average estimated yield loss attributed to WSMV in Kansas from 1976 to 1996 was 2%.168 The development of genetic resistance to A. tosichilla in Canada and the U.S. has provided substantial monetary savings to agricultural producers in the form of increased wheat yields and decreased production costs for miticides. Mite resistance in commercial cultivars reduces the incidence of WSMV, preventing approximately three-fourths of the virus losses incurred in A. tosichilla-susceptible cultivars. Using 1993 crop values and production figures,4 this amounts to a current annual value of more than $150 million (U.S.) for A. tosichilla resistance in North American wheats. Combined Value of Arthropod Resistance in Wheat. The economic value of genetic resistance in wheat for all of the arthropod pests discussed above amounts to just over $250 million (U.S.) each year (Table 5). This includes $20 million for S. graminum resistance, $12.9 million for D. noxia resistance, $50 million for M. destructor resistance, $15.7 million for C. cinctus resistance, and $151.8 million for A. tosichilla resistance. All of these values are gross values, without the addition of any economic multiplier effects. Frequently, the use of multiplier effects will double or triple the value of a cost or a savings in the economics of agricultural productivity.

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TABLE 5 Global Economic Value of Arthropod-Resistant Wheat Arthropod pest

Approximate value ($U.S. million)

Schizaphis graminum Diuraphis noxia Mayetiola destructor Cephus cinctus Aceria tosichilla Total value

B. ECOLOGICAL

AND

20.0 12.9 50.0 15.7 151.8 250.4

HUMANITARIAN VALUES

In addition to the significant economic advantages mentioned previously, ecological and environmental benefits arise from the use of arthropod-resistant wheat cultivars. Populations of beneficial insects and microorganisms increase in the agroecosystem when pesticide use is reduced, as does species diversity. With reduced pesticide applications, wheat in the vegetative growth stage is safer for livestock grazing and for use as habitat by game birds. The environmental benefits of producing wheat using arthropod resistance instead of pesticides include cleaner soils and ground water supplies. A significant reduction in insecticide use on U.S. wheat is envisaged with greater use of resistant cultivars, as was estimated to have occurred with the use of arthropod-resistant cultivars of barley, corn, and sorghum.120 In at least one instance, the value of arthropod resistance in wheat has humanitarian value. In South Africa, D. noxia-resistant wheat cultivars have been given to farmers in Lesotho, where wheat production had been severely disrupted by D. noxia damage and civil strife. Farmers renamed the cultivar “Puseletso,” meaning “that which has been returned to us,” and wheat production in Lesotho has been reestablished.166

IV.

FUTURE DIRECTIONS FOR ARTHROPOD RESISTANCE RESEARCH

Molecular marker techniques are being used more frequently in the development of pest-resistant wheat. One technique, restriction fragment length polymorphism (RFLP) analysis, is based on inherent variations in the sites where restriction endonucleases cut DNA. RFLP analysis makes use of genetic markers derived from cloned DNA fragments to analyze different fragment lengths as alleles of particular genes. Once arthropod resistance genes are shown to be associated (linked) with an RFLP marker, selection for these genes can be based on the genotype of the RFLP marker, rather than the plant phenotype. The marker can then be used to assess plant populations segregating for resistance before the phenotypic trait for resistance is expressed. This process can greatly accelerate the rate of breeding for arthropod-

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resistant plants.142 There is a substantial need for improved techniques to access linked traits associated with resistance to arthropods and other pests. Current research is limited in the use of the tertiary germplasm pool, where genetic recombination is restricted for the non-homologous chromosomes transferred in T. aestivum.30 Using RFLP analysis, high-density genetic maps are being constructed by entomologists, geneticists, and plant breeders to locate genes of economic value, including arthropod and disease resistance genes, that flank DNA markers. An extensive library of DNA markers has been developed for wheat and related cereal crops in the Kansas State University Wheat Generic Resource Center.37 Several of these markers have already been used to map the RFLP linkage locations of genes for resistance to A. tosichilla, D. noxia, and M. destructor.16,24,25,36,75,88

V.

CONCLUSIONS

Pronounced economic and environmental accomplishments have resulted from the cooperative efforts of entomologists, geneticists, and plant breeders who have used the genetic diversity of wild Triticum and closely related species to develop arthropodresistant wheats during the past 40 years. From a strictly economic perspective, the current value of this resistance is more than $250 million (U.S.) each year. The ecological value of this resistance has not been rigorously assessed, but the cultivation of arthropod-resistant cultivars has greatly decreased pesticide usage in the U.S. This decline in pesticide use has contributed to a healthier environment for humans, livestock, and wildlife. Wheat producers have benefitted from arthropod-resistant cultivars in the form of reduced production costs, and consumers have benefitted because their food has been produced more economically and safely. Although many arthropod-resistant cultivars have been developed, research and development must continue to maintain the benefits of this resistance in global food production. Researchers and extension educators must continue to monitor the development of virulence genes in newly developing biotypes of wheat arthropod pests. Where possible, accurate, efficient, and economical techniques based on molecular genetic markers must be developed and implemented to monitor biotypes. The pools of resistance genes available to researchers to breed into wheat cultivars that succumb to these new biotypes must be increased. This will require sustained or enhanced funding for collection, curation, and evaluation of germplasm to identify these genes. Additional efforts should be made to identify resistance where it is lacking, or where it has been identified, to breed it into an existing cultivar. Acceptance and use of the resistance identified to R. padi and S. avenae should be undertaken by producers in areas, especially Europe, where these pests cause significant losses to wheat production. Significant levels of resistance to M. dirhodum are yet to be identified. This and other voids should challenge researchers and their supporting government or donor constituents to cooperatively develop this resistance and place it into production. Whether they are developing new resistant cultivars or improving existing ones, researchers must continue to identify wheat germplasm with arthropod resistance, from both conventional and transgenic sources. A large amount of germplasm

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remains to be evaluated. Only a modest percentage of the more than 300 species of Triticeae have been evaluated for arthropod resistance. Those evaluated include A. elongatum, S. cereale, T. speltoides, T. tauschii, and T. turgidium, some very rich sources of resistance. However, many opportunities await the interdisciplinary research efforts of entomologists, geneticists, molecular biologists, and plant breeders to identify additional resistance genes. Wheat is a food that is critical to the survival of the world population. With the human population predicted to double to over 10 billion people by 2040, it is essential that global wheat production be increased to meet this need. Arthropod-resistant wheat cultivars should be integral components of that production system because of their proven economic and environmental benefits. A continual development of arthropod-resistant cultivars depends heavily on the availability of diverse, wellmaintained collections of Triticeae adequately supported by local, regional, national, and international funding.

ACKNOWLEDGMENTS The authors thank Roger Ratcliffe and Edward Souza for their reviews and suggestions to improve the chapter.

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68. Lampert, E. P., D. L. Haynes, A. J. Saywer, D. P. Jokinen, S. G. Wellso, R. L. Gallun, and J. J. Roberts, Effects of regional releases of resistant wheats on the population dynamics of the cereal leaf beetle (Coleoptera: Chrysomelidae), Ann. Entomol. Soc. Am., 76:972-980, 1983. 69. Leather, S. R., K. F. A. Walters, and A. F. G. Dixon, Factors determining the pest status of the bird cherry-oat aphid, Rhopalosiphum padi (L.) (Hemiptera: Aphididae), in Europe: a study and review, Bull. Ent. Res., 79:345-360, 1989. 70. LeClerg, E. L., H. T. Cook, C. D. Van Hauweling, R. J. Anderson, A. M. Vance, K. Dorwood, and H. R. Thomas, Injurious Crop Insects, Losses in Agriculture, Agriculture Handbook No. 291, U.S. Department of Agriculture, Agricultural Research Service, Washington, D.C., 1965, 43. 71. Legg, D. and S. Amosson, Economic impact of the Russian wheat aphid in the western United States:1991-1992, Great Plains Agr. Council Publication 147, Colorado State University Cooperative Extension, Fort Collins, 1993. 72. Leszczynski, B., Winter wheat resistance to the grain aphid Sitobion avenae (Fabr.) (Homoptera, Aphididae), Insect Sci. Applic., 8:251-254, 1987. 73. Lilenfield, F. A. and H. Kihara, Genome-analysis in Triticum and Aegilops. Concluding Review, Cytologia, 16:101-123, 1951. 74. Lowe, H. J. B., Resistance and susceptibility to colour forms of the aphid Sitobion Avenae in spring and winter wheats (Triticum aestivum), Ann. Appl. Biol., 99:87-98, 1981. 75. Ma, Z.-Q., B. S. Gill, M. E. Sorrells, and S. D. Tanksley, RFLP markers linked to Hessian fly-resistance genes in wheat (Triticum aestivum L.) from Triticum tauschii (Coss.) Schmal., Theor. Appl. Genet., 85:750-754, 1993. 76. Maas, F. B. III, F. L. Patterson, J. E. Foster, and J. H. Hatchett, Expression and inheritance of resistance of 'Marquillo' wheat to Hessian fly biotype D, Crop Sci., 27:49-52, 1987. 77. Maas, F. B. III, L. Patterson, J. E. Foster, and H. W. Ohm, Expression and inheritance of resistance of ELS 6404-160 durum wheat to Hessian fly, Crop Sci., 29:23-28, 1989. 78. Marais, G. F. and F. du Toit, A monosomic analysis of Russian wheat aphid resistance in the common wheat PI294994, Plant Breed., 111:246-248, 1993. 79. Martin, T. J., T. L. Harvey, G. G. Bender, and D. L. Seifers, Control of wheat streak mosaic virus with vector resistance in wheat, Phytopathol., 74:963-964, 1984. 80. Martin, T. J., T. L. Harvey, G. C. Bender, D. L. Seifers, and J. H. Hatchett, Wheat curl mite resistant wheat germplasm, Crop Sci., 23:809, 1983. 81. Martin, T. J., T. L. Harvey, and R. W. Livers, Resistance to wheat streak mosaic virus and its vector, Aceria tulipae, Phytopathology, 66:346-349, 1976. 82. Miller, R. H., Insect pests of wheat and barley in West Asia and North Africa, ICARDA Tech. Manual No. 9, Rev.1, ICARDA, Aleppo, 1987. 83. Miller, R. H., S. El Masri, and K. Al Jundi, Plant density and wheat stem sawfly (Hymenoptera: Cephidae) resistance in Syrian wheats, Bull. Ent. Res., 83:95-102, 1993. 84. Monte, J. V., C. L. McIntyre, and J. P. Gustafson, Analysis of phytogenetic relationship in the Triticeae using RFLP’s, Theor. Appl. Genet. 86:649-655, 1993. 85. Morrill, W. L., J. W. Gabor, E. A. Hockett, and G. D. Kushnak, Wheat stem sawfly (Hymenoptera: Cephidae) resistance in winter wheat, J. Econ. Entomol., 85:2008-2011, 1992. 86. Morrison, P., Current distribution and economic impact, In: F. B. Peairs and S. D. Pilcher, Eds., Proc. 2nd Russian Wheat Aphid Workshop, Denver, 1988, 5.

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87. Niassy, A., J. D. Ryan, and D. C. Peters, Variations in feeding behavior, fecundity, and damage of biotypes B and E of Schizaphis graminum (Homoptera:Aphididae) on three wheat genotypes, Environ. Entomol., 16:1163-1168, 1987. 88. Nieto-Lopez, R. M. and T. K. Blake, Russian wheat aphid resistance in barley: inheritance and linked molecular markers, Crop Sci., 34:655-659, 1994. 89. Nkongolo, K. K., J. S. Quick, A. E. Limin, and D. B. Fowler, Sources and inheritance of resistance to Russian wheat aphid in Triticum species and Triticum tauschii, Can. J. Plant Sci.,71:703-708, 1991. 90. Nkongolo, K. K., J. S. Quick, W. L. Meyers, and F. B. Peairs, Russian wheat aphid resistance of wheat, rye, and triticale in greenhouse tests, Cereal Res. Comm., 17:227-232, 1989. 91. Obanni, M., H. W. Ohm, J. E. Foster, and F. L. Patterson, Genetics of resistance of PI 422297 durum wheat to the Hessian fly, Crop Sci., 29:249-252, 1989. 92. Obanni, M., F. L. Patterson, J. E. Foster, and H. W. Ohm, Genetic analyses of resistance of durum wheat PI 428435 to the Hessian fly, Crop Sci., 28:223-226, 1988. 93. Oellerman, C. M., F. L. Patterson, and R. L. Gallun, Inheritance of resistance in ‘Lusco’ wheat to Hessian fly, Crop Sci., 23:222-224, 1983. 94. Ohm, H. W., R. H. Ratcliffe, F. L. Patterson, and S. E. Cambron, Resistance to Hessian fly conditioned by genes H19 and proposed gene H27 of durum wheat line PI422297, Crop Sci., 37:113-115, 1997. 95. Painter, R. H., Biological strains of Hessian fly, J. Econ. Entomol., 23:322-326, 1930. 96. Painter, R. H., Crops that resist insects provide a way to increase world food supply, Kansas State Agric. Exp. Stn. Bull. 520, Kansas State University, Manhattan, 1968. 97. Papp, M., Resistance of winter wheat cultivars to the cereal leaf beetle (Oulema melanoplus L.) and bird cherry oat aphid (Rhopalosiphum padi L.), Novenytermeles, 39:11-22, 1990. 98. Papp, M. and A. Mesterhazy, Resistance to bird cherry-oat aphid (Rhopalosiphum padi L.) in winter wheat varieties, Euphytica, 67:49-57, 1993. 99. Patterson, F. L., J. E. Foster, and H. W. Ohm, Gene H16 in wheat for resistance to Hessian fly, Crop. Sci., 28:652-654, 1988. 100. Patterson, F. L., J. E. Foster, H. W. Ohm, J. E. Hatchett, and P. L. Taylor, Proposed system of nomenclature for biotypes of Hessian fly (Diptera: Cecidomyiidae) in North America, J. Econ. Entomol., 85:307-311, 1992. 101. Patterson, F. L. and R. L. Gallun, Inheritance of resistance of Seneca wheat to race E of Hessian fly, In: E. R. Sears and L. M. Sears, Eds., Proc., 4th Int. Wheat Genetics Symp., Missouri Agricultural Experiment Station, Columbia, 1973, 445. 102. Peters, D. C., E. A. Wood, Jr., and K. J. Starks, Insecticide resistance in selections of the greenbug, J. Econ. Entomol., 75:339-340, 1975. 103. Porter, D. R., J. A. Webster, and C. A. Baker, Detection of resistance to Russian wheat aphid in hexaploid wheat, Plant Breed., 110:157-160, 1993. 104. Porter, D. R., J. A. Webster, R. L. Burton, G. L. Puterka, and E. L. Smith, New sources of resistance to greenbug in wheat, Crop Sci., 31:1502-1504, 1991. 105. Porter, D. R., J. A. Webster, and B. Friebe, Inheritance of greenbug biotype G resistance in wheat, Crop Sci., 34:625-628, 1994. 106. Porter, K. B., G. L. Peterson, and O. Vise, A new greenbug biotype, Crop Sci., 22:847-850, 1982. 107. Puterka, G. J., D. C. Peters, D. L. Kerns, J. E. Slosser, L. Bush, D. W. Worrall, and R. W. McNew, Designation of two new greenbug (Homoptera: Aphididae) biotypes G and H, J. Econ. Entomol., 81:1754-1759, 1988.

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108. Ramirez, I. A., A review of barley yellow dwarf in the Southern cone countries of South America, In: P. A. Burnett, Ed., World Perspectives on Barley Yellow Dwarf, CIMMYT, El Battan, 1990, 362. 109. Ratanatham, S. and R. L. Gallun, Resistance to Hessian fly (Diptera: Cecidomyiidae) in wheat as affected by temperature and larval density, Environ. Entomol.,15:305-310, 1986. 110. Ratcliffe, R. H. and J. H. Hatchett, Biology and genetics of the Hessian fly and resistance in wheat, In: K. Bobdari, Ed., New Developments in Entomology, Research Signpost, Scientific Information Guild, Trivandrum, 1997, 47. 111. Ratcliffe, R. H., H. W. Ohm, F. L. Patterson, and S. E. Cambron, Biotype composition of Hessian fly (Diptera: Cecidomiidae) populations from Arkansas, Georgia, Illinois, North Carolina and Virginia, J. Entomol. Sci., 32:154-164, 1997. 112. Ratcliffe, R. H., H. W. Ohm, F. L. Patterson, S. E. Cambron, and G. G. Safranski, Response of resistance genes H9 -H19 in wheat to Hessian fly (Diptera: Cecidomiidae) laboratory biotypes and field populations from the eastern United States, J. Econ. Entomol., 89:1309-1317, 1996. 113. Ratcliffe, R. H., G. C. Safranski, F. L. Patterson, H. W. Ohm, and P. L. Taylor, Biotype status of Hessian fly (Diptera: Cecidomyiidae) populations from the eastern United States and their responses to 14 Hessian fly resistance genes, J. Econ Entomol., 87:1113-1121, 1994. 114. Raupp, W. J., A. Amri, J. H. Hatchett, B. S. Gill, D. L. Wilson, and T. S. Cox, Chromosomal location of Hessian fly-resistance genes H22, H23, and H24 derived from Triticum tauschii in the D-genome of wheat, J. Hered.,84:142-145, 1993. 115. Rejesus, R. M., The world wheat situation, CIMMYT World Wheat Facts and Trends Supplement, 1995, Ongoing Research at CIMMYT: Understanding Wheat Genetic Diversity and International Flows of Genetic Resources, CIMMYT, El Battan, 1995. 116. Roberts, D. W. A. and C. Tyrrell, Sawfly resistance in wheat. IV. Some effects of light intensity on resistance, Can. J. Plant Sci., 41:457-465, 1961. 117. Roberts, J. J. and J. E. Foster, Effect of leaf pubescence in wheat on the bird cherry oat aphid (Homoptera: Aphidae), J. Econ. Entomol., 76:1320-1322, 1983. 118. Roberts, J. J., R. L. Gallun, F. L. Patterson, and J. E. Foster, Effects of wheat leaf pubescence on the Hessian fly, J. Econ. Entomol., 72:211-214, 1979. 119. Saidi, A. and J. S. Quick, Inheritance and allelic relationships among Russian wheat aphid resistance genes in winter wheat, Crop Sci., 36:256-258, 1996. 120. Schalk, J. M. and R. H. Ratcliffe, Evaluation of ARS program on alternative methods of insect control: host plant resistance to insects, Bull. Entomol. Soc. Am., 22:1-7, 1976. 121. Schepers, A., Chemical control, In: A. K. Minks and P. Harrewijn, Eds., Aphids, Their Biology, Natural Enemies and Control, Elsevier, Amsterdam, 1989, 89. 122. Schroeder-Teeter, S., R. S. Zemetra, D. J. Schotzko, C. M. Smith, and M. Rafi, Monosomic analysis of Russian wheat aphid (Diuraphis noxia) resistance in Triticum aestivum line PI137739, Euphytica, 74:117-120, 1994. 123. Sebesta, E. E. and E. A. Wood, Jr., Transfer of greenbug resistance from rye to wheat with x-rays, Agron. Absts., 61-62, 1978. 124. Seigenthaler, V. L., J. E. Stepanich, and L. W. Briggle, Distribution of the Varieties and Classes of Wheat in the United States, 1984, Crops Branch, Estimates Division, Statistical Reporting Service, U.S. Department of Agriculture Statistics Bull. 739, U.S. Government Printing Office, Washington, D.C., 1986. 125. Shands, R. G. and W. B. Cartwright, A fifth gene conditioning Hessian fly response in common wheat, Agron. J., 45:302-307, 1953.

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126. Simmons, S. R., Growth, development and physiology, In G. E. Heyne, Ed., Wheat and Wheat Improvement, Agron. Monograph 13, ASA, CSSA, & SSSA, Madison, 1987, 77. 127. SINGER (System-wide Information Network on Genetic Resources), Consultative Group on International Agricultural Research, on the World Wide Web (http://nocl.cgiar.org/cgintrus.htm), 1996. 128. Singh, S. R. and E. A. Wood, Jr., Effect of temperature on fecundity of two strains of the greenbug, J. Econ. Entomol., 56:l09-110, 1963. 129. Slykhuis, J. T., Aceria tulipae Keifer (Acarina: Eriophyidae) in relation to the spread of wheat streak mosaic, Phytopathology, 45:116-128, 1955. 130. Smith, C. M., D. J. Schotzko, R. S. Zemetra, E. J. Souza, and S. SchroederTeeter, Identification of Russian wheat aphid (Homoptera: Aphididae) resistance in wheat, J. Econ. Entomol., 84:328-332, 1991. 131. Smith, K. M., A Textbook of Plant Virus Diseases, Academic Press, New York, 1972. 132. Sosa, O., Jr., Biotype L, ninth biotype of the Hessian fly, J. Econ. Entomol., 71:458-460, 1978. 133. Sosa, O., Jr., Hessian fly: resistance of wheat as affected by temperature and duration of exposure, Environ. Entomol., 8:280-281, 1979. 134. Sosa, O., Jr., Biotypes J and L of the Hessian fly discovered in an Indiana wheat field, J. Econ. Entomol., 74:180-182, 1981. 135. Sosa, O., Jr. and J. E. Foster, Temperature and the expression of resistance in wheat to the Hessian fly, Environ. Entomol., 5:333-336, 1976. 136. Sotherton, N. W. and H. F. van Emden, Laboratory assessments of resistance to the aphids Sitobion avenae and Metopolophium dirhodum in three Triticum species and two modern wheat cultivars, Ann. Appl. Biol., 101:99-107, 1982. 137. Souza, E., J. M. Windes, S. S. Quisenberry, D. J. Schotzko, P. F. Lamb, S. Halbert, R. S. Zemetra, and C. M. Smith, Registration of Idaho 471A and Idaho 471B wheat germplasms, Crop Sci., 37:1031, 1997. 138. Souza, E., J. M. Windes, S. S. Quisenberry, D. J. Schotzko, P. F. Lamb, S. Halbert, R. S. Zemetra, and C. M. Smith, Registration of Idaho 472 wheat germplasm, Crop Sci., 37:1032, 1997. 139. Stebbins, N. B., F. L. Patterson, and R. L. Gallun, Interrelationships among wheat genes for resistance to Hessian fly, Crop Sci., 2:177-180, 1980. 140. Stebbins, N. B., F. L. Patterson, and R. L. Gallun, Interrelationships among wheat genes H3, H6, H9, and H10 for Hessian fly resistance, Crop Sci., 22:1029-1032, 1982. 141. Stebbins, N. B., F. L. Patterson, and R. L. Gallun, Inheritance of resistance of PI 94587 wheat to biotypes B and D of Hessian fly, Crop Sci., 23:251-253, 1983. 142. Tanksley, S. D., N. D. Young, A. H. Paterson, and M. W. Bonierbale, RFLP mapping in plant breeding: new tools for an old science, Biotechnol., 7:257-263, 1989. 143. Teetes, G. L., C. A. Schaefer, J. R. Gipson, R. C. McIntyre, and E. E. Latham, Greenbug resistance to organophosphorous insecticides on the Texas high plains, J. Econ. Entomol., 68:214-216, 1975. 144. Thomas, J. B. and R. L. Conner, Resistance to colonization by the wheat curl mite in Aegilops squarrosa and its inheritance after transfer to common wheat, Crop Sci., 26:527-530, 1986. 145. Tonet, G. L. and R. F. P. Dasilva, Antibiosis of wheat genotypes to C-biotypes of Schizaphis graminum (Rondani, 1852) (Homoptera, Aphididae), Pesquisa Agr. Brasil, 29:1181-1186, 1994. 146. Tremblay, C., C. Cloutier, and A. Comeau, Resistance to the bird cherry-oat aphid, Rhopalosiphum padi L. (Homoptera: Aphididae) in perennial gramineae and wheat x perennial gramineae hybrids, Environ. Entomol.,18:921-932, 1989.

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147. Tyler, J. M. and J. H. Hatchett, Temperature influence on expression of resistance to Hessian fly (Diptera: Cecidomyiidae) in wheat derived from Triticum tauschii, J. Econ. Entomol.,76:323-326, 1983. 148. Tyler, J. M., J. A. Webster, and O. G. Merkle, Designations for genes in wheat germplasm conferring greenbug resistance, Crop Sci., 27:526-527, 1987. 149. Tyler, J. M., J. A. Webster, and E. L. Smith, Biotype E greenbug resistance in wheat streak mosaic virus-resistant wheat germplasm lines, Crop Sci., 25:686-688, 1985. 150. Wallace, L. E., F. H. McNeal, and M. A. B. Berg, Resistance to both Oulema melanoplus and Cephus cinctus in pubescent-leaved and solid stemmed wheat selections, J. Econ. Entomol., 67:105-110, 1974. 151. Walters, M. C., F. Penn, F. du Toit, T. C. Botha, K. Aalbersberg, P. H. Hewitt, and S. W. Broodryk, The Russian wheat aphid, farming in South Africa, Leaflet Series, Wheat C3, Government Printer, Pretoria, 6, 1980. 152. Webster, J. A., The cereal leaf beetle in North America: breeding for resistance in small grains, Ann. N. Y. Acad. Sci., 287:230-237, 1977. 153. Webster, J. A., S. H. Gage, and D. H. Smith, Jr., Suppression of the cereal leaf beetle with resistant wheat, Environ. Entomol., 2:1089-1091, 1973. 154. Webster, J. A. and P. Kenkel, Benefits of managing small grain pests with plant resistance, In: J. A. Webster and B. R. Wiseman, Eds., Economic, Environmental, and Social Benefits of Insect Resistance in Field Crops, Thomas Say Publications, Entomological Society of America, Lanham, in press. 155. Webster, J. A., D. H. Smith, Jr., and R. P. Hoxie, Effect of cereal leaf beetle on the yields of resistant and susceptible winter wheat, Crop Sci., 22:836-840, 1982. 156. Webster, J. A., D. H. Smith, Jr., and C. Lee, Reduction in yield of spring wheat caused by cereal leaf beetles, J. Econ. Entomol., 65:832-835, 1972. 157. Webster, J. A., D. H. Smith, Jr., E. Rathke, and C. E. Cress, Resistance to cereal leaf beetle in wheat: density and length of leaf-surface pubescence in four wheat lines, Crop Sci., 15:199-202, 1975. 158. Weiss, M. J. and W. L. Morrill, Wheat stem sawfly (Hymenoptera: Cephidae) revisited, Amer. Entomol., 38:241-245, 1992. 159. Wellso, S. G., Cereal leaf beetle: larval feeding, orientation, development, and survival on four small-grain cultivars in the laboratory, Ann. Entomol. Soc. Am., 66:1201-1208, 1973. 160. Wellso, S. G., Cereal leaf beetle: interaction with and ovipositional adaptation to a resistant wheat, Environ. Entomol.,8:454-457, 1979. 161. Wellso, S. G., Cereal leaf beetle (Coleoptera: Chrysomelidae) and winter wheat: host plant resistance relationships, Great Lakes Entomol., 19:193-197, 1986. 162. Whelan, E. D. and G. E. Hart, A spontaneous translocation that transfers wheat curl mite resistance from decaploid Agropyron elongatum to common wheat, Genome, 30:289-292, 1988. 163. Wood, E. A., Jr., Biological studies of a new greenbug biotype, J. Econ. Entomol., 54:1171-1173, 1961. 164. Zohary, D. and M. Hopf, Domestication of Plants in the Old World, Clarendon Press, Oxford, 1993. 165. Peairs, F.B., Personal communication, 1996. 166. du Toit, F., unpublished data. 167. Sandvol, L.E., Personal communication, 1997. 168. Bowden, R.L., Personal communication, 1997.

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3

Insect Resistance in Barley Germplasm David R. Porter, Dolores W. Mornhinweg, and James A. Webster

CONTENTS I. Introduction II. The Barley Crop A. History B. Importance C. Germplasm Resources III. Pests of Barley A. Pest Status B. Sources of Resistance 1. Diuraphis noxia 2. Schizaphis graminum 3. Rhopalosiphum padi 4. Oulema melanopus 5. Other Pests IV. Conclusions References

I.

INTRODUCTION

Barley, Hordeum vulgare L., germplasm collections have long played a crucial role in providing valuable genes for breeding pest-resistant, high-performance cultivars. Continuous improvement in yield and quality of barley cultivars is dependent on the availability and judicious use of the diverse genetic resources found in our national and international germplasm collections. Increasing genetic diversity within commercial cultivars could promote the expansion of production potential through enhanced adaptation and tolerance to biotic and abiotic stresses. However, by its very nature, the process of developing new barley cultivars has resulted in a narrowing of the genetic base that is now available to producers. The number of alleles of commonly studied loci encountered in samples of H. vulgare ssp. spontaneum K. Koch, the progenitor of cultivated barley, decreased dramatically in the progression from primitive materials to the most advanced modern cultivars.1 Repeated

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cycles of selection for specific quality traits under limited environments, within existing adapted germplasm, have produced narrow pools of elite barley genotypes.49 The challenges and potential dangers of limited genetic variability in cultivated crops is not a new concern. The northern corn leaf blight (caused by Helminthosporium turcicum Pass) that reduced U.S. corn production in the 1970s is a classic example of the consequences of over-reliance on a very narrow gene base. Lack of genetic diversity for pest resistance in commercial barley cultivars limits the potential for expansion of current production areas into regions where insect pests are problematic. In addition, present production areas are being threatened by new pests (e.g., Russian wheat aphid, Diuraphis noxia [Mordvilko]). Currently, almost half of the total U.S. malting barley production is limited to three midwestern states, namely Minnesota, North Dakota, and South Dakota.50 Two cultivars, “Stander” and “Robust,” account for approximately 75% of the acreage planted to malting barley in these states.2 This spatially and genetically concentrated production of malting barley could be vulnerable to new pests. It might be possible to expand barley’s production area beyond the regions where it is now grown if the crop were more pest-resistant. For example, D. noxia has severely impacted feed barley production in eastern Colorado. All commercially available barleys are susceptible to D. noxia, and the low profit margin of feed barley production restricts the use of insecticides. Because damaging populations of D. noxia are now common in eastern Colorado, feed barley production acreage has declined 90% in this area.59 The development and use of D. noxia-resistant barleys would reclaim this lost production area. This chapter will focus on the use of germplasm collections for the enhancement of pest resistance in barley. Specifically, we review barley history and importance, germplasm resources, pests of barley, and successes in identifying and developing pest-resistant germplasm lines and cultivars.

II.

THE BARLEY CROP

A. HISTORY The origin of barley, one of the oldest domesticated crops, is thought to be in the Fertile Crescent area of the Near East.20 Early Egyptians believed the goddess Isis gave them barley, and thus it was the first grain used for food.21 Based on the oldest evidence available, it was not until about 8000 B.P. that barley was used for brewing alcoholic beverages.19 Beer made from fermented barley figured prominently in the commerce of ancient Egypt, where it was sold for local consumption and even exported. Many ancient civilizations viewed fermented beverages as sources of supernatural powers, and drunkenness was considered a religious experience.19 The importance of barley as food continues to take a lesser role and today it is more often used in malt production and livestock feed.41 Barley under domestication is divided into two types: two-rowed forms (primitive condition) in which only the central spikelet at each node of the inflorescence rachis is fertile and six-rowed forms in which all three spikelets are fertile.43,58 Mature barley spikes can be erect or nodding; dense or lax; awnleted, awnless, or hooded;

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and hulled or hull-less (naked).43 Primitive hulled barleys were probably cultivated by 9000 B.P. in the Near East.58 Today, hulled two-rowed and hulled six-rowed barleys are widely grown.

B. IMPORTANCE Barley is the world’s fourth most important cereal after wheat Triticum aestivum L., maize Zea mays L., and rice Oryza sativa L.40 However, barley’s economic impact is much greater if its value to the brewing industry is considered. In 1995, Canada was the largest single producing country with 11.7 mmt.41 Europe and the former Soviet Union account for 68% of the total world production of 161 mmt. In the U.S., the 1995 value of barley production ($1.0 billion) made it the fifth most important cereal behind maize ($23.1 billion), wheat ($9.7 billion), rice ($1.5 billion), and sorghum, Sorghum bicolor (L.) Moench, ($1.4 billion). The 1994 U.S. per capita consumption of malt-equivalent barley food products was 772 g compared with 65 kg of wheat flour. Of the total U.S. supply of barley in 1994 (12.6 mmt), approximately 48% was used for animal feed; 38% for food, alcohol, and seed; and 14% for export.41

C. GERMPLASM RESOURCES Major Hordeum seed collections are maintained by the U.S. Department of Agriculture-Agricultural Research Service (USDA-ARS), National Small Grains Collection (NSGC), Aberdeen, Idaho; the International Center for Agricultural Research in the Dry Areas (ICARDA), Aleppo, Syria; and the International Maize and Wheat Improvement Center (CIMMYT), El Battan, Mexico. The U.S. collection has grown steadily from the initial introduction of four accessions in 189436 to the USDA collection at Aberdeen, Idaho, with over 30,000 plant introduction (PI) accessions from 20 Hordeum species (see Chapter 12 in this volume). This collection contains cultivars, advanced germplasm lines, landraces, and genetic stocks, with some dating back to 1900. Extensive collections from other countries are obtained through plant explorations and through exchanges with national and international germplasm repositories. The U.S. barley collection at Aberdeen, Idaho, is an active collection, compared with the long-term storage collection at the USDA-ARS National Seed Storage Laboratory, Fort Collins, Colorado. Other countries (e.g., England, Russia, Germany, Brazil, The Netherlands, Bulgaria, and Czech Republic) also store and maintain large collections of barley germplasm. ICARDA maintains a collection of over 22,000 accessions and CIMMYT has a collection of over 9,000 barleys.45

III.

PESTS OF BARLEY

A. PEST STATUS Over 100 species of insects attack barley,48 most of which are occasional pests that cause economic damage during outbreaks. For example, the army cutworm, Euxoa auxiliaris (Grote), is an occasional pest in the U.S., but it can devastate barley fields © 1999 by CRC Press LLC

in outbreak years.48 In contrast, the primary pest D. noxia causes serious economic losses every year throughout much of the barley-producing areas of the western U.S. The bird cherry-oat aphid, Rhopalosiphum padi L., is not a primary pest in the U.S., but is one of the most severe pests in spring barley in northern Europe.57 Aphids are the most important pests of barley. These insects puncture the leaves and feed in the phloem. The Russian wheat aphid, greenbug Schizaphis graminum (Rondani), and bird cherry-oat aphid are considered the most serious aphid pests in North America. In some European countries the corn leaf aphid, Rhopalosiphum maidis (Fitch), is considered the most serious pest, causing up to 35% annual yield loss.51 The greenbug was the most serious insect pest of barley in North America until the Russian wheat aphid appeared in the 1980s.48 Diuraphis noxia has caused over $1 billion (U.S.) in losses in the western U.S. since it was detected in Texas in 1986.39 Economic losses caused by S. graminum are not well documented, but this aphid and R. padi are more damaging on spring barley than are R. maidis and the English grain aphid, Sitobion avenae F.26 In addition to removing essential plant nutrients, some aphids are vectors of plant viruses. Rhopalosiphum padi is often considered to be problematic only through its ability to vector barley yellow dwarf virus (BYDV), but its feeding damage can cause up to 50% yield loss in the absence of BYDV.26

B. SOURCES

OF

RESISTANCE

Diuraphis noxia, S. graminum, and R. padi have received the most attention with regard to identifying sources of plant resistance and developing resistant germplasm. Each of these aphids will be discussed individually, along with research on cereal leaf beetle Oulema melanopus L., Hessian fly Mayetiola destructor Say, R. maidis, and grasshoppers Melanoplus spp. 1. Diuraphis noxia The USDA collection has been a valuable source of germplasm in the search for pest-resistant material, most recently for D. noxia.52 All U.S. barley cultivars in production were highly susceptible to D. noxia when this aphid was detected in the southern Great Plains in 1986. Early attempts to find resistance among greenbugresistant barleys were unsuccessful.55 Efforts to find resistant barley germplasm commenced when 524 USDA accessions, originating from regions where barley and D. noxia coexist, were evaluated in greenhouse tests.52 From this research and further analysis of the categories of resistance (antibiosis, antixenosis, and tolerance) of resistant plants, five resistant accessions from Afghanistan (PIs 366444, 366447, 366449, 366450, and 366453) and one from Iran (CI1412) were identified (Table 1).52 Progeny of resistant plants selected from screening tests were used as parents in a resistance breeding program. From this work, one highly resistant accession (PI366450, collected by J. D. Gray in 1965) was selected and used in an accelerated germplasm enhancement program. It was necessary to select for homogeneity of resistance within PI366450 because this accession was heterogeneous for

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TABLE 1 Evaluation of Barley Genetic Resources for Insect Resistance No. of entries Insect Diuraphis noxia Russian wheat aphid Schizaphis graminum Greenbug Rhopalosiphum padi Bird cherry-oat aphid Oulema melanopus Cereal leaf beetle Mayetiola destructor Hessian fly Rhopalosiphum maidis Corn leaf aphid Melanoplus spp. Grasshoppers

Screened

Resistant

Ref.

524 111 23,070 577 1,230 1,005 474 500 172 8,634 5,114

6 15 109 194 160 15 41 29 6 17 33

52 5 33 12 9 54 24,25 57 16 15 22

121

7

23

32

8

29

D. noxia resistance. This selection resulted in the release of STARS-9301B, the first D. noxia-resistant barley germplasm.31 Genetic analysis of the resistance in STARS9301B indicated control by two genes, Dnb1 and Dnb2.32 Specifically, D. noxia resistance is expressed through recessive epistasis of the dominant gene Dnb2 on the incompletely dominant gene Dnb1. Multiple gene control also has been reported for STARS-9577B, a D. noxia-resistant germplasm line selected from CIho 4165 (USDA accession from Afghanistan).33 Analysis of D. noxia resistance in PI366444 and PI366453 indicated that at least two resistance genes are shared or tightly linked in both lines.37 Molecular marker analysis located one of the genes in the short arm of chromosome 5.37 Barley germplasm accessions have been evaluated under field conditions. Out of 111 spring barleys artificially infested with D. noxia, 15 entries were classified as resistant (Table 1).5 Two of these resistant lines, ASE/2CM//B76BB and “Gloria/Come,” underwent further analysis to determine the genetic control of resistance.44 Although studies indicated that the same dominant gene controls resistance in ASE/2CM//B76BB and “Gloria/Come,”44 it is not known if this gene is the same as Dnb1 or Dnb2 in STARS-9301B or a different gene.34 Cultivars developed using these resistance genes are at potential risk from new introductions of D. noxia or genetic recombinations within existing U.S. populations that produce plant resistance-breaking biotypes. Thus, when Puterka et al.42 discovered that PI366450 was susceptible to a D. noxia population from Iachmen, Kyrgyzstan, a

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search for additional sources of resistance was started. After screening the entire USDA collection of available H. vulgare ssp. vulgare accessions (23,070), 109 accessions were rated resistant to moderately resistant to D. noxia (Table 1).33 In addition, evaluations of accessions of wild Hordeum species revealed substantial inter- and intraspecific variability for resistance to D. noxia.10,27 Further research showed that the presence of clavicipitaceous endophytic fungi (Neotyphodium) in some of these wild Hordeum accessions was responsible for high levels of resistance to D. noxia.11 Transfers of D. noxia resistance from slender wheatgrass, Elymus trachycaulus Link, into H. vulgare produced resistant hybrids that may be useful in backcrossing.4 Thus, genetic diversity for D. noxia resistance exists in conserved germplasm. 2. Schizaphis graminum In the U.S., a greenbug outbreak in 1942 caused significant yield losses to small grains (estimated at $38 million) and precipitated interest in the development of resistant cultivars.3 By chance, S. graminum colonized two barley nurseries in Denton, Texas, and Lawton, Oklahoma, and differential S. graminum damage among nursery cultivars from China and Korea indicated the potential for host plant resistance. Subsequently, screenings of 577 germplasm lines identified S. graminum resistance in 194 accessions, including the Korean landrace “Omugi” (PI87181) (Table 1).12 Genetic analysis indicated that a single dominant gene controls resistance in PI87181.17 The S. graminum-resistant barleys “Kerr” and “Will” were developed using two different sources of S. graminum resistance, one from PI87181 and one from “Kearney,” respectively.30 The cultivar “Kerr” was developed in 1952 by crossing PI87181 with “Rogers.”14 Greenbug resistance in “Will” is likely from “Kearney,” a Nebraska S. graminum-resistant winter barley selected from Composite Cross III (CCIII) in 1941 and released in 1951. The source of the S. graminum resistance gene in CCIII is unknown; this population was developed by intercrossing 13 winter and spring cultivars in all combinations and was released in 1931.35 Genetic analysis of PI87181, “Kearney,” and two other S. graminum-resistant barleys (“Dobaku” and CIho 5087) indicated that the same dominant gene controls resistance in all four lines.47 It is quite possible that the resistance gene (Rsg1a)30 in PI87181 was common among many of the landraces of southeast Asia, specifically east-central China and Korea, collected during 1929 to 1930. The presence of the same gene in “Kearney” resulted because the 13 cultivars (e.g., “Everest,” “Esaw,” “Nakano Wase,” and “Smooth Awn 86”) used in the development of CCIII were collected from Asia or had Asian cultivars in their pedigree.3 The resistance gene Rsg1a was made available in sets of S. graminum-resistant and -susceptible nearisolines in 1988.6 Today, these near-isolines are being used for molecular genetic analysis of the S. graminum resistance.60 The chain of events highlighted above demonstrates the value of conserved germplasm and its use in pest resistance efforts. A landrace of winter barley collected in Korea over 67 years ago provided a gene that has saved U.S. producers countless dollars in pesticide applications, and now is the focus of experiments to better

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understand the nature of plant resistance to insects. The entire collection of winter barleys, totaling 1,230 accessions, was screened in 1961 and 160 accessions with S. graminum resistance were identified, some more resistant than PI87181 (Table 1).9 The occurrence of new S. graminum biotypes, which can damage previously resistant germplasm, has prompted searches for new sources of resistance. For example, the advent of greenbug biotype E in the U.S. prompted a 1981 search of the USDA collection which identified 15 accessions with varying levels of resistance (Table 1).54 A different gene (Rsg2b) is responsible for the resistance in one line (PI426756).30 Also, resistance to S. graminum has been detected in H. chilense,8 and wheat/H. chilense disomic addition lines have been analyzed to locate the resistance genes on chromosomes.7 These lines may be useful in providing sources of S. graminum resistance for cultivated barley in the future. Efforts to produce S. graminum-resistant cultivars, such as “Tambar 402,”18 “Tambar 500,”28 and “Post,”13 have used PI87181 (“Omugi”) and “Will” as the sources of the aphid resistance. 3. Rhopalosiphum padi Barley resistance to R. padi protects plants from direct feeding damage and may reduce the incidence of BYDV by reducing feeding of viruliferous aphids. Screening of 474 barley cultivars from the Canadian Genetic Stock of Barley showed 41 selections with tolerance and antibiosis resistance to R. padi (Table 1).24,25 In addition, two of the aphid-resistant selections (“Rojo” and CI3906-1) exhibited BYDV resistance.24 However, none of the 474 cultivars in the collection showed very high levels of aphid resistance, and only 43 selections (9%) were considered sources of resistance for breeding purposes.25 Use of these resistant selections to develop R. padi-resistant barley cultivars has not been reported. Hordeum species other than cultivated barley (H. vulgare ssp. vulgare) have been evaluated as potential sources of resistance to R. padi. Testing of a Swedish collection of 27 accessions of Hordeum species and interspecific hybrids (crosses between H. vulgare ssp. vulgare and related Hordeum species) showed significant differences in the ability of aphids to reproduce when feeding on the different species.56 A diploid species, H. bogdani Wilenski, had very high levels of resistance, but genetic incompatibilities between this species and H. vulgare make it difficult to transfer the resistance to cultivated barley. In addition, a collection of 500 accessions of H. vulgare ssp. spontaneum K. Koch from Israel was evaluated under natural infestations of R. padi in Sweden, and 29 resistant selections were reevaluated in greenhouse tests (Table 1).57 Two of the resistant selections were crossed with cultivated barley and some of the progeny were more resistant than the resistant parents. Efforts to produce R. padi-resistant barley cultivars using these resistance sources are ongoing.57 4. Oulema melanopus The search for cereal leaf beetle resistance began in 1963 when 172 cultivars of barley were field screened for resistance to adult and larval feeding (Table 1).16 Over

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94% of the entries were moderately or severely damaged. In a subsequent screening of 8,634 accessions from the USDA collection, less than 1% were resistant to larval feeding damage.15 In contrast to yield losses of up to 38% for susceptible cultivars, two resistant accessions (CI6671 and CI6469) produced high yields in beetle-infested plots.53 These two resistant accessions were used as parents in a breeding program to produce two improved O. melanopus-resistant spring barley germplasms, CI15820 and CI15821.46 However, Oulema melanopus-resistant cultivars have not been made available to producers. 5. Other Pests Extensive screening of the USDA barley collection has identified resistance to other insect pests, such as M. destructor, R. maidis, and Melanoplus spp. The bulk of the USDA collection (5,114 accessions) was evaluated for M. destructor resistance from 1945 to 1950.22 Of the total accessions tested, 7 accessions were highly resistant and 26 moderately resistant to M. destructor (Table 1).22 Genetic analysis of three highly resistant cultivars, “Delta,” “Nile,” and “Abusir,” indicated that two dominant genes (Hf and Hf2) control resistance to M. destructor.38 Cultivars with resistance to M. destructor have not been released to the barley industry. A screening of 121 winter barley cultivars for R. maidis resistance detected high levels of resistance in seven entries (Table 1).23 Unfortunately, two of the resistant entries (“Davie” and “Rogers”) from this study are susceptible to S. graminum. Another line in this study (PI87181) was susceptible to corn leaf aphid, but is resistant to greenbug. These differential plant responses indicate that different genes control resistance to the two aphid species.23 Rhopalosiphum maidis-resistant barley cultivars are not available. Melanoplus spp.-resistant cultivars were discovered under natural infestations in Canadian field tests from 1944 to 1947 (Table 1).29 We are unaware of any followup research on barley resistance to Melanoplus spp.

IV.

CONCLUSIONS

Germplasm repositories have a rich history of providing new sources of barley resistance to insect pests. Seed from the unpretentious barley plant, gathered by intrepid collectors from isolated areas, is often a treasure-trove of genetic resources. All that is required to uncover these assets is a reliable bioassay for screening tests, a focused approach to systematically evaluating a large number of accessions, and perseverance from dedicated entomologists and plant breeders. History shows that resistance has been found in H. vulgare ssp. vulgare L. accessions for every pest studied to date. However, if levels of resistance identified in cultivated barley types are not high enough to be useful in a resistance breeding program, then searches of other Hordeum species should proceed. This chapter shows that the world’s Hordeum germplasm stocks are a source of strong resistance to barley pests.

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REFERENCES 1. Allard, R. W., Methods for germplasm identification, In: H.T. Stalker and J.P. Murphy, Eds., Plant Breeding in the 1990s, C.A.B. International, Wallingford, 1991, 122. 2. Anonymous, Barley Variety Survey, American Malting Barley Association, Inc., Milwaukee, 1996. 3. Atkins, I. M. and R. G. Dahms, Reaction of small-grain varieties to greenbug attack, USDA Tech. Bull. 901, 1945. 4. Aung, T., Intergeneric hybrids between Hordeum vulgare and Elymus trachycaulus resistant to Russian wheat aphid, Genome, 34:954-960, 1991. 5. Calhoun, D. S., P. A. Burnett, J. Robinson, and H. E. Vivar, Field resistance to Russian wheat aphid in barley: I. symptom expression, Crop Sci., 31:1464-1467, 1991. 6. Carver, B. F., G. H. Morgan, L. H. Edwards, and J. A. Webster, Registration of four pairs of greenbug-resistant vs. susceptible near-isolines of winter barley germplasms, Crop Sci., 28:1034-1035, 1988. 7. Castro, A. M., A. Martin, and L. M. Martin, Location of genes controlling resistance to greenbug (Schizaphis graminum Rond.) in Hordeum chilense, Plant Breed., 115:335-338, 1996. 8. Castro, A. M., L. M. Martin, A. Martin, H. O. Arriaga, N. Tobes, and L. B. Almaraz, Screening for greenbug resistance in Hordeum chilense Roem et Schult., Plant Breed., 112:151-159, 1994. 9. Chada, H. L., I. M. Atkins, J. H. Gardenhire, and D. E. Weibel, Greenbugresistance studies with small grains, Texas Agric. Exp. Stn. Bull. B-982, Texas A&M University, College Station, 1961. 10. Clement, S. L. and D. G. Lester, Screening wild Hordeum species for resistance to Russian wheat aphid, Cereal Res. Commun., 18:173-177, 1990. 11. Clement, S. L., A. D. Wilson, D. G. Lester, and C. M. Davitt, Fungal endophytes of wild barley and their effects on Diuraphis noxia population development, Entomol. Exp. Appl., 82:275-281, 1997. 12. Dahms, R. G., T. H. Johnston, A. M. Schlehuber, and E. A. Wood, Jr., Reaction of small-grain varieties and hybrids to greenbug attack, Okla. Agric. Exp. Stn. Tech. Bull. T-55, Oklahoma State University, Stillwater, 1955. 13. Edwards, L. H., E. L. Smith, H. Pass, and G. H. Morgan, Registration of Post barley, Crop Sci., 25:363, 1985. 14. Edwards, L. H., E. L. Smith, H. Pass, and E. A. Wood, Jr., Registration of Kerr barley, Crop Sci., 10:725, 1970. 15. Gallun, R. L., E. H. Everson, R. F. Ruppel, and J. C. Craddock, Cereal leaf beetle resistance studies (1963-1964), USDA Special Rep. W-200, 1964. 16. Gallun, R. L., R. Ruppel, and E. H. Everson, Resistance of small grains to the cereal leaf beetle, J. Econ. Entomol., 59:827-829, 1966. 17. Gardenhire, J. H. and H. L. Chada, Inheritance of greenbug resistance in barley, Crop Sci., 1:349-352, 1961. 18. Gardenhire, J. H., M. E. McDaniel, and N. A. Tuleen, Registration of Tambar 402 barley, Crop Sci., 22:1259, 1982. 19. Hardwick, W. A., History and antecedents of brewing, In: W. A. Hardwick, Ed., Handbook of Brewing, Marcel Dekker, Inc., New York, 1995, 37. 20. Harlan, J. R., On the origin of barley, In: Barley: Origin, Botany, Culture, WinterHardiness, Genetics, Utilization, Pests, USDA Agric. Handbook No. 338, U.S. Department of Agriculture-Agriculture Research Service, Washington, D.C., 1968, 9.

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21. Hedrick, U. P., Sturtevant’s Notes on Edible Plants, N.Y. Dept. of Agriculture, Vol. 2, Part 2, 1919. 22. Hill, C. C., W. B. Cartwright, and G. A. Wiebe, Barley varieties resistant to the Hessian fly, Agron. J., 44:4-5, 1952. 23. Hormchong, T. and E. A. Wood, Jr., Evaluation of barley varieties for resistance to the corn leaf aphid, J. Econ. Entomol., 56:113-114, 1963. 24. Hsu, S. J. and A. G. Robinson, Resistance of barley varieties to the aphid Rhopalosiphum padi (L.), Can. J. Plant Sci., 42:247-251, 1961. 25. Hsu, S. J. and A. G. Robinson, Further studies on resistance of barley varieties to the aphid Rhopalosiphum padi (L.), Can. J. Plant Sci., 43:343-348, 1962. 26. Kieckhefer, R. W. and B. H. Kantack, Yield loss in spring barley caused by cereal aphids (Homoptera: Aphididae) in South Dakota, J. Econ. Entomol., 79:749-752, 1986. 27. Kindler, S. D. and T. L. Springer, Resistance to Russian wheat aphid in wild Hordeum species, Crop Sci., 31:94-97, 1991. 28. Marshall, D. S., J. H. Gardenhire, B. A. Shafer, K. B. Porter, M. D. Lazar, M. E. McDaniel, L. R. Nelson, and W. D. Worrall, Registration of ‘Tambar 500’ barley, Crop Sci., 33:1104, 1993. 29. McBean, D. S. and A. W. Platt, Differential damage to barley varieties by grasshoppers, Sci. Agric., 31:162-175, 1951. 30. Merkle, O. G., J. A. Webster, and G. H. Morgan, Inheritance of a second source of greenbug resistance in barley, Crop Sci., 27:241-243, 1987. 31. Mornhinweg, D. W., D. R. Porter, and J. A. Webster, Registration of STARS9301B barley germplasm resistant to the Russian wheat aphid, Crop Sci., 35:602, 1995. 32. Mornhinweg, D. W., D. R. Porter, and J. A. Webster, Inheritance of Russian wheat aphid resistance in spring barley, Crop Sci., 35:1368-1371, 1995. 33. Mornhinweg, D. W., D. R. Porter, and J. A. Webster, Germplasm enhancement for RWA resistance, Barley Newsl., 39:92-94, 1995. 34. Mornhinweg, D. W., D. R. Porter, and J. A. Webster, Inheritance of Russian wheat aphid resistance in spring barley germplasm line STARS-9577B, Barley Genetics Newsl., 25:34-35, 1995. 35. Moseman, J. G., Use of introduced germplasm in the USDA-ARS national barley collection in barley cultivars, In: H. L. Shands and L. E. Wiesner, Eds., Use of Plant Introductions in Cultivar Development, Part 1, CSSA Special Publication no. 17, Crop Science Society of America, Madison, 1991, 49. 36. Moseman, J. G. and D. H. Smith, Jr., Germplasm resources, In: D. C. Rasmusson, Ed., Barley, ASA, CSSA, & SSSA, Madison, 1985, 57. 37. Nieto-Lopez, R. M. and T. K. Blake, Russian wheat aphid resistance in barley: inheritance and linked molecular markers, Crop Sci., 34:655-659, 1994. 38. Olembo, J. R., F. L. Patterson, and R. L. Gallun, Genetic analysis of the resistance to Mayetiola destructor (Say) in Hordeum vulgare L., Crop Sci., 6:563-566, 1966. 39. Patrick, C. D. and S. Amosson, Economic impact of the Russian wheat aphid in the United States: 1993-1994, Texas A&M Univ., College Station, in press. 40. Poehlman, J. M., Adaptation and distribution, In: D. C. Rasmusson, Ed., Barley, ASA, CSSA, & SSA, Madison, 1985, 1. 41. Pratt, B., Statistics of grain and feed, In: Agricultural Statistics, USDA, National Agricultural Statistics Service, U.S. Government Printing Office, Washington, D.C., 1996, 1.

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42. Puterka, G. J., J. D. Burd, and R. L. Burton, Biotypic variation in a worldwide collection of Russian wheat aphid (Homoptera: Aphididae), J. Econ. Entomol., 85:1497-1506, 1992. 43. Reid, D. A., Morphology and anatomy of the barley plant, In: D. C. Rasmusson, Ed., Barley, ASA, CSSA, & SSSA, Madison, 1985, 73. 44. Robinson, J., F. Delgado, H. E. Vivar, and P. A. Burnett, Inheritance of resistance to Russian wheat aphid in barley, Euphytica, 62:213-27, 1992. 45. SINGER (System-wide Information Network on Genetic Resources), Consultative Group on International Agricultural Research, on the World Wide Web (http://noc1.cgiar.org/cgintrus.htm), 1997. 46. Smith, Jr., D. H., J. A. Webster, and J. E. Grafius, Registration of two cereal leaf beetle resistant barley germplasms, Crop Sci., 24:828, 1984. 47. Smith, O. D., A. M. Schlehuber, and B. C. Curtis, Inheritance studies of greenbug (Toxoptera graminum Rond.) resistance in four varieties of winter barley, Crop Sci, 2:489-491, 1962. 48. Starks, K. J. and J. A. Webster, Insects and related pests, In: D. C. Rasmusson, Ed., Barley, ASA, CSSA, & SSSA, Madison, 1985, 335. 49. Ullrich, S. E., Barley Germplasm Status in the USA, Barley Crop Germplasm Committee Report, Pullman, 1996. 50. U.S. Department of Agriculture, Prospective Plantings Report, National Agricultural Statistics Service, U.S. Government Printing Office, Washington, D.C., 1996. 51. Verma, G. P., D. N. Mahto, and M. H. Haque, The loss in grain yield due to aphids and other pests on certain varieties of barley grown in North Bihap, Sci. Cult., 45:370-371, 1979. 52. Webster, J. A., C. A. Baker, and D. R. Porter, Detection and mechanisms of Russian wheat aphid (Homoptera: Aphididae) resistance in barley, J. Econ. Entomol., 84:669-673, 1991. 53. Webster, J. A. and D. H. Smith, Jr., Yield losses and host selection of cereal leaf beetles in resistant and susceptible barley, Crop Sci., 19:901-904, 1979. 54. Webster, J. A. and K. J. Starks, Sources of resistance in barley to two biotypes of the greenbug Schizaphis graminum (Rondani) (Homoptera: Aphididae) Prot. Ecol., 6:51-55, 1984. 55. Webster, J. A., K. J. Starks, and R. L. Burton, Plant resistance studies with Diuraphis noxia (Homoptera: Aphididae), a new United States wheat pest, J. Econ. Entomol., 80:944-949, 1987. 56. Weibell, J., Screening for resistance against Rhopalosiphum padi (L.). 2. Hordeum species and interspecific hybrids, Euphytica, 36:571-576, 1987. 57. Weibell, J., Resistance to Rhopalosiphum padi (Homoptera: Aphididae) in Hordeum vulgare subsp. spontaneum and in hybrids with H. vulgare subsp. vulgare, Euphytica, 78:97-101, 1994. 58. Zohary, D. and M. Hopf, Domestication of Plants in the Old World, Clarendon Press, Oxford, 1993. 59. Peairs, F.B., Personal communication, 1997. 60. Hays, D.B., Personal communication, 1997.

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4

Genetic Diversity of Sorghum: A Source of Insect-Resistant Germplasm George L. Teetes, Gary C. Peterson, Kanayo F. Nwanze, and Bonnie B. Pendleton

CONTENTS I. II. III. IV. V.

Introduction Importance and Use Basic Plant Characteristics Origin and Domestication Germplasm Resources A. Germplasm Collections B. Sorghum Conversion Program VI. Modern Genetic Selection in Sorghum VII. Resistance to Insects VIII. Role of Insects in Diversifying the Genetics of Sorghum IX. Co-evolutionary Relationship of Insects and Sorghum X. Case Studies XI. Economic Benefit of Insect-Resistant Sorghum XII. Molecular Genetic Techniques for Development of Insect-Resistant Sorghum XIII. Conclusions Acknowledgments References

I.

INTRODUCTION

Modern sorghum, Sorghum bicolor (L.) Moench, is a product of human ingenuity.3 This marvelous plant has been domesticated, selected, and changed to meet human needs. Sorghum yields well in areas too hot and too dry to produce other crops such as maize, Zea mays L., and thus has prevented starvation of millions of people who live in the environmentally harshest and most austere regions of the world.

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The crop grows well in small, subsistence plots under primitive cultivation methods or as vast monocultures utilizing the most modern production practices and equipment. Sorghum is known for its production reliability. But, sorghum is not without an array of about 75 species of insects that infest the crop, and several of these insects threaten yield certainty and food security. Early domestication of sorghum and subsequent movement to many regions of the world, especially semi-arid tropical areas, resulted in the evolution of a genetically diverse crop species. This genetic diversity has been exploited in crop improvement programs to increase yield and yield stability by incorporating beneficial genes for many traits including adaptation to a wide variety of habitats and uses, higher yield, and more desirable grain. More recently, a major effort has been made to increase genetic resistance for defense against abiotic and biotic stresses, including insect pests. In each sorghum agroecosystem one or two key insect pests such as sorghum midge Stenodiplosis sorghicola (Coquillett), greenbug Schizaphis graminum (Rondani), or shoot fly Atherigona soccata (Rondani), dominate control practices. Several species, such as Banks grass mite Oligonychus partensis (Banks) and corn earworm Helicoverpa zea (Boddie), are secondary or induced pests injurious as a result of changes in cultural practices or cultivars, or because of injudicious use of insecticides applied for a key pest. Most insects that infest sorghum are occasional pests that cause economic damage only in localized areas or sporadically in time. Occasional insect pests include species of wireworms, white grubs, cutworms, aphids, leaf- and panicle-feeding caterpillars, and leaf- and panicle-feeding bugs. The challenge for crop protectionists is to use the genetic diversity of sorghum to combat a complex of insect pests. Development and use of sorghums resistant to key insect pests provide the greatest immediate benefit to an integrated pest management approach. However, there is opportunity to identify, incorporate, and deploy sorghums resistant to most insect pests of the crop. Toward this goal, much has been accomplished but much remains to be done.

II.

IMPORTANCE AND USE

Sorghum usually is considered fifth in importance among the world’s cereal crops. Worldwide, about 50 million ha of sorghum are grown annually, with the greatest hectarage located in Africa and Asia (Table 1).24

TABLE 1 World Production of Sorghum in 1995 Region

Hectares

Africa Asia North America Central and South America Other Total

22,000,000 15,000,000 4,000,000 2,000,000 3,000,000 46,000,000

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Sorghum grain is used for human food and as feed for animals. Through the ages, sorghum has been a vital food source for billions of people, especially in the semiarid tropics of Africa and Asia. Sorghum grain as human food is prepared in many traditional ways. Unleavened bread prepared with flour ground from grain is a common food made from sorghum. Sometimes the dough is fermented before baking. Sorghum grain also is boiled into a porridge or gruel. Beer commonly is made from the grain. Sorghum stems and foliage are used for green chop, hay, silage, and pasture, and after grain harvest stalks may be used for fuel or construction. Sorghum as a feed grain for livestock and poultry is first coarsely ground, broken, or soaked by various procedures. These treatments expose a larger portion of the kernel to an animal’s digestive enzymes, and less passes undigested through the animal. Generally, sorghum grain has the nutritional qualities of maize, but the kernel size is smaller.

III.

BASIC PLANT CHARACTERISTICS

Sorghum is a large-stemmed tropical grass having the ability to grow to great heights. Sorghum is a perennial, grown as an annual, is a self-pollinating species, but is handled in advanced agricultural situations as a cross-pollinating crop. The rate of outcrossing of sorghum generally is considered to be about 5%. The diverse types in the species are all diploids with a 2N = 20 chromosome number, and they will intercross.62 Related grassy species are 2N = 40. Cytoplasmic-genetic male sterility is extremely important in hybridization of sorghum. Sorghum is classified into four groups: (1) grain sorghum, (2) forage or saccharine sorghum, (3) broomcorn, and (4) grass sorghum.3 Grain sorghum is known by a variety of names: guinea corn in West Africa, kafir corn in South Africa, durra in the Sudan, mtama in eastern Africa, jowar in northern India, kaoliang in China, and milo in America. Types with sweet and juicy stems from which syrup can be extracted are known as sorgos, while those with long stiff panicle branches from which whisk brooms are made are broomcorns. Sorghum is adapted to many environments, requiring 90 to 140 days to mature. Yields usually are higher from later rather than earlier maturing cultivars. Yields have exceeded 11,000 kg/ha, with above-average yields ranging from 7,000 to 9,000 kg/ha where moisture is not a limiting factor.3 In areas where sorghum commonly is grown, yields of 3,000 to 4,000 kg/ha are obtained under better conditions, dropping to 300 to 1,000 kg/ha as moisture becomes limiting. Sorghum hybrids are grown in developed countries where commercial seed industries exist. Typically these hybrids are short statured and machine harvested. In most developing countries, landrace varieties instead of hybrids are grown, and farmers select and save seed from their crops for planting. In subsistence agriculture, sorghum is harvested by hand and usually stored unthreshed until needed for food preparation, then threshed and decorticated.

IV.

ORIGIN AND DOMESTICATION

While there is general consensus of opinion that sorghum originated in the northeastern quadrant of Africa, there is less agreement on the region of Africa where it © 1999 by CRC Press LLC

was first domesticated.10,16,17,30,45,52 Harlan30 proposed that sorghum was a “noncentric crop,” that is, a crop whose current distribution of genetic variability revealed no evident center of origin or diversity. Based on comparative morphological studies and numerical taxonomy, three independent centers of sorghum domestication have been proposed — the Ethiopian area of eastern Africa, tropical western Africa, and southeastern Africa.10 After considering the distribution of races of sorghum in Africa, Harlan30 concluded that initial domestication of sorghum occurred in a long belt across central Africa. Doggett16 concluded that sorghum was first cultivated by the Cushites in the Ethiopian highlands of eastern Africa. Domestication of sorghum was believed to have begun about 5,000 B.P.45 Afterwards, various tribes in different parts of Africa domesticated different races of sorghum as knowledge of crop cultivation spread. The durra race was domesticated in the Ethiopian-Sudan region, and the guinea race was developed in western Africa, at least in part by Mande tribesmen. The Bantu people adopted sorghum as a staple food as they began to populate the dry savannas east of the forest belt. From Tanzania southward the kafir race of sorghum came to be associated with the Bantu, just as the caudatum race became associated with the Nilotic and Nil-Hamitic people of Uganda and western Kenya. The fact that sorghum is an extremely diverse species with much morphological variability resulted in various tribes using sorghum in different ways, from building materials for huts, furniture, and mats, to beer, sugar, and cereal production.10 Cultivation of the durra race of sorghum spread from Ethiopia and Sudan into the Near East and to India soon after 4,000 B.P. Sorghum probably was not cultivated in ancient Egypt. It is absent from the extensive archaeological excavations of early farming sites in the Near East. In the first century (60 to 70 A.D.), Pliny48 mentioned the introduction of sorghum into Italy by caravans from India. The spread of sorghum cultivation into southeastern Asia and China also occurred around 3,000 B.P., but there are no authentic records of sorghum in China before the 13th century.15,48 At approximately this time period, sorghum was in Southeast Asia where the crop first came in contact with wild sorghum diploids such as S. propinquum (Kunth) Hitchc. Genetic exchange between wild and cultivated forms may have given rise to the characteristic koaliang types of China, Manchuria, and Japan.16 Sorghum, called “guinea corn,” “indian millet,” and “coffee corn,” probably was introduced from western Africa into the Western Hemisphere.16,47,48,58 Sorghum probably first entered the U.S. along with slaves. Benjamin Franklin is credited with introducing broomcorn.48 Sorghum is believed to have been under cultivation in the U.S. before the end of the 18th century.47 There are records of the early presence of sorghum (“broomcorn,” “indian millet,” and “coffeecorn”) in the U.S. in the 1800s.20,58

V.

GERMPLASM RESOURCES

A. GERMPLASM COLLECTIONS Organized collecting of Sorghum began in the mid-1950s at the time of the hybrid production era.19 Collections exceeding 30,000 accessions have been assembled at

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the International Crops Research Institute for the Semi-Arid Tropics (ICRISAT) Asia Center, India and at the Plant Genetic Resources Conservation Laboratory, Griffin, Georgia. The National Seed Storage Laboratory, Fort Collins, Colorado, is a security backup to the active collection at Griffin, Georgia. About 80% of the total world sorghum collection was introduced from developing countries of the semiarid tropics. Only 1.2% of the world sorghum collection is composed of wild and weedy relatives. Accessions in these world seed collections were obtained from tropical and temperate zones, from low and high elevations, and from sorghum grown during different seasons of the year. Sorghums from these varied environments ensured germplasm diversity. If resistance to any sorghum insect pest exists in the world, it probably can be found in these collections, although much of the existing germplasm has not yet been collected or evaluated for resistance to insects.

B. SORGHUM CONVERSION PROGRAM Many sorghums in the world collection are from Africa and Asia. Most of these tropical varieties, because they are photoperiod-sensitive and require long nights to induce flowering, cannot be evaluated in temperate zones of the world. As a result, only a small fraction of the total genetic diversity within the species could be evaluated in its original form and used to produce improved lines and hybrids in temperate areas. Sorghum improvement specialists have long recognized the restricted germplasm base of sorghum adapted for temperate areas. Early research that culminated in development of hybrid grain sorghum used some of the few temperately adapted sorghum cultivars that were naturally short and early maturing.59 Discovery and utilization of cytoplasmic-genetic male sterility to produce hybrid sorghum revolutionized sorghum production around the world. However, there was little germplasm available for temperate areas to make additional significant improvements in grain yield, adaptation, and resistance to abiotic and biotic stresses. Initiated in 1963, the cooperative Texas Agricultural Experiment Station, Texas A&M University, U.S. Department of Agriculture (USDA), Agricultural Research Service (ARS), Sorghum Conversion Program70 has made available a portion of the genetic diversity within the genus Sorghum. This program uses the genetics of sorghum height and maturity, and crossing and selection to convert tall, photoperiodsensitive sorghums into short, early types suitable for use in temperate areas. Other than the discovery of the sterility system that allowed for commercial production of hybrid sorghum, the sorghum conversion program has been the largest contributor to the improvement of sorghum in the developed world. Sorghum is a short-day species, but strains exhibit differential sensitivity to photoperiod. Four genes influence the photoperiod reaction that governs the time of floral initiation (duration of growth). Lateness of maturity and tallness are dominant characteristics, and each is controlled by genes at four independently inherited loci.58 In the sorghum conversion program, crossing and backcrossing are done during the winter at the USDA Experiment Station, Mayaguez, Puerto Rico. Short, early genotypes are selected from segregating populations during the long days of summer

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in Texas. Tropical sorghums are used as male parents in the crosses and backcrosses except the last cross, when the tropical line is used as the female to restore the converted line to its original cytoplasm. The original female (nonrecurrent parent) is an early, four-dwarf, B-line (male fertile) in normal cytoplasm. Each backcross is made to an F3 progeny grown from a short, early maturing F2 plant selected in Texas. Four cycles of backcrossing and selection are required for conversion; however, partially converted lines (lines with less than four backcrosses) sometimes are released as breeding stocks.

VI.

MODERN GENETIC SELECTION IN SORGHUM

It is readily apparent that much of the genetic diversity within sorghum is a result of its early domestication, movement to many regions of the world, and selection for many different habitats and uses. Plant genetic improvement efforts today generally attempt to maximize yield, which result in a recombination of some genetic components and, inadvertently, in the loss of others.34 Newly developed crop cultivars are grown over large areas and used for as many years as possible. Crop cultivars or hybrids generally are genetically homogeneous from one plant to another within a field. Thus, common dogma is that modern cultivated plants such as sorghum have a narrower genetic base than those of even a few decades ago. This genetic and phenotypic uniformity tends to increase the vulnerability of crop plants to insect pest attack, and the practice of monoculture further exacerbates the problem.34 However, the genetic base of any given crop can be broadened to include insectresistant characteristics that intrinsically protect plants from insect pests. Incorporating resistance to insects is one way to broaden a crop’s genetic base, obtain general benefits from increased genetic diversity, and gain specific benefits from resistance. Deliberate attempts to use plant resistance in any crop span primarily the last 100 years, with the greatest emphasis on resistance made during the last five decades. Even less time has been devoted to developing insect-resistant sorghums. Considering that the history of plant domestication is several thousand years old, the longneglected task of deliberately incorporating insect resistance into plants, including sorghum, has just begun. Hopefully, greater knowledge of the subject and better techniques will allow fast, effective progress.

VII.

RESISTANCE TO INSECTS

Resistance has been reported for many of the common insect and mite species infesting sorghum (Table 2). This table indicates the insects for which sorghum germplasm has been evaluated and provides estimates of the germplasm lines evaluated and the number of resistance sources identified. Thousands of sorghum germplasm lines, most from the world collection, have been evaluated for resistance to several aphid species, especially greenbug, shoot fly, stem borers, sorghum midge, and panicle-infesting bugs. These efforts are evidence of the commitment of sorghum improvement specialists to identify resistance, incorporate resistance into improved plant types, and deploy insect-resistant sorghum. Although the success rate in identifying resistant sources has been relatively low, resistance has been identified for © 1999 by CRC Press LLC

TABLE 2 Sorghum Germplasm Evaluated for Resistance to Insects Insect Schizaphis graminuma Melanaphis sacchari Rhopalosiphum maidis Sipha flava Atherigona soccata Spodoptera frugiperda Busseola fusca Chilo partellus Diatraea saccharalis Peregrinus maidisa Blissus leucopterus leucopterus Oligonychus pratensis Stenodiplosis sorghicola Calocoris angustatus Eurystylus oldi (marginatus)a

No. of germplasm lines evaluated

No. of resistant lines identified

Ref.

29,000 13,337 650 8,809 31,000 8,990 16,000 30,000 200 104 1,000 553 18,491 18,000 400

28 108 73 3 134 34 18 182 4 3 53 2 70 58 9

— 4, 5, 28, 46, 51, 55 38, 42, 49 9, 68, 77 53, 63, 65 12-14, 27, 78 26 53, 63, 65 27 — 8, 19, 50, 69 25 1, 41, 53, 63, 65, 79 53, 63-65 —

a

Information provided by H. C. Sharma, especially stem borers and panicle-feeding bugs, and Y. O. Doumbia for Eurystylus oldi.

almost all of the important insect pests. However, far fewer insect-resistant sorghums actually have become a component of integrated pest management in agricultural production. Most success has been with resistance to greenbug in the U.S., resistance to sorghum midge in Australia and India, and resistance to stem borers and shoot fly in Africa and India. Because most insects infesting sorghum, especially those insects originating in the Western Hemisphere, do not have a long history of association with sorghum, it is surprising that resistance would occur in sorghum to so many of the insects that infest the crop. This is a testament to sorghum genetic diversity.

VIII.

ROLE OF INSECTS IN DIVERSIFYING THE GENETICS OF SORGHUM

Evolutionary biologists have been preoccupied for almost a century with assessing the role of different abiotic and biotic stresses, including insects, in generating plant genetic diversity. Although caution is important when applying theories of evolutionary biology to agricultural systems, evolutionary biology is of interest to crop protectionists whose goal is to locate and incorporate into plants genes for defense against insect pests. A common, but not fully agreed upon, hypothesis is that much of the genetic diversity of plants has arisen as a product of co-evolution between plants and their natural enemies (i.e., insect pests).22 For this reason, centers of origin of domesticated plants are believed to be important sources of insect-resistant genes

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for incorporation into improved varieties. Two important issues are related to this hypothesis: (1) “selection imposed by natural enemies causes plant populations to diverge in defensive characteristics” and (2) “selection imposed by plant defensive characteristics causes the plant’s natural enemy populations to diverge in characteristics associated with exploitation of the host plant.” Issue 1 implies that plants within a population that are weakened or unable to survive the attack of insects are replaced through natural selection by better-adapted plants. Plants that survive will have genes for defense against the insect. The controversy regarding this issue is that selection by natural enemies of plants is normally too weak to account for evolution of defense traits such as secondary compounds, morphological characteristics, and other features.40 Also, resistance to an insect commonly is found in plants that have had no long-term association with the insect pest. The consequence of the second issue is sometimes an unfortunate, real-world catastrophe, formation of virulent biotypes, after deployment of resistant genotypes into an agroecosystem. This issue further illustrates the importance of plant genetic diversity, but also stresses the importance of considering the genetic diversity of the insect.

IX.

CO-EVOLUTIONARY RELATIONSHIP OF INSECTS AND SORGHUM

The co-evolutionary relationships between insect pests and sorghum have not been thoroughly assessed. Cursory examination reveals that most insects infesting sorghum are widely distributed species, generalist feeders, that did not co-evolve with sorghum, but resistance has been identified for many of these insects (Table 3). The insects infesting and having the longest and most host-specific association with sorghum are sorghum midge and sorghum shoot fly. Most insects infesting sorghum have many hosts and because of their proposed geographic origins could not have evolved with sorghum. Greenbug is a good example of a key insect pest of sorghum that did not evolve with sorghum, but resistance has been identified and used in agriculture. Resistance genes were selected for in the absence of the insects. Characters that cause resistance to these insects have physiological or ecological functions other than or in addition to protection of the sorghum plant. Seemingly, resistance traits developed in a plant population free of an insect pest are fortuitously derived from pleiotropic effects of genes maintained by natural selection pressures unrelated to the insect. Harris34 referred to this kind of resistance as allopatric resistance, which he defined as “heritable qualities possessed by a plant that influence the ultimate degree of damage done by an insect having no prior continuous co-evolutionary history with that insect.” In contrast, he defined sympatric resistance as “resistance that comes from insects having a prior, long-term history with a plant species.”

X.

CASE STUDIES

Two key insect pests that represent sympatric and allopatric relationships with sorghum are sorghum midge and greenbug, respectively. Sorghum midge is a dipterous, multivoltine species with a short life-cycle (16 days). Only species of the genus

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TABLE 3 Geographical and Ecological Relationships of Sorghum with Insects Infesting Sorghum Insect

Pest status

Schizaphis graminum Melanaphis sacchari

Key Occ

Rhopalosiphum maidis Sipha flava Atherigona soccata Mythimna separata Spodoptera exempta Spodoptera frugiperda Busseola fusca Chilo partellus Eldana saccharina Sesamia calamistis Sesamia inferens

Possible origin

Current distribution

Hosts

Co-evolved w/sorghum

Resistance reported

E & W Hemisph. E & W Hemisph.

Grasses Grasses

No No

Yes Yes

Occ Occ Key Occ Occ Occ Occ Key Occ Occ Occ

Mediterranean Asia, Africa, Tropical America Americas Americas Africa, Asia Asia Africa Americas S Africa India Africa Africa Asia

E & W Hemisph. W Hemisph. E Hemisph. Asia, Pacific E & W Africa W Hemisph. Africa India, E Africa, Far East Africa Africa Asia, Pacific

No No Yes No No No No No No No No

Yes Yes Yes Yes No Yes Yes Yes No No No

Diatraea saccharalis

Occ

C America

C & S America

No

Yes

Peregrinus maidis

Occ

C America

No

No

Poophilus costalis Locris rubens Blissus leucopterus leucopterus Oligonychus indicus Oligonychus pratensis

Occ Occ Occ

Africa W Africa Americas

Worldwide but more tropical, especially S Africa & C America Asia & Africa W Africa W Hemisph.

Grasses, Solanaceae Grasses Sorghum Grasses, many dicots Grasses, many dicots Grasses, many dicots Sorghum, maize, millet, grasses Sorghum, maize, sugarcane, grasses Sorghum, sugarcane, maize Grasses Sorghum, sugarcane, maize, rice, wheat, millet Sorghum, sugarcane, maize, aquatic grassy weeds Sorghum, maize, sugarcane

Sorghum, grasses Sorghum, grasses Grasses

No No No

No No Yes

Sec Sec

India Americas

E Hemisph. W Hemisph.

Sorghum, sugarcane, grasses Grasses

No No

No Yes

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TABLE 3 (continued) Geographical and Ecological Relationships of Sorghum with Insects Infesting Sorghum Insect

Pest status

Possible origin

Current distribution

Hosts

Co-evolved w/sorghum

Resistance reported

Tetranychus urticae Stenodiplosis sorghicola Cryptoblades adoceta Dichocrocis punctiferalis Heliothis armigera

Sec Key

Americas NE Africa

W Hemisph. E & W Hemisph.

Grasses Sorghum spp.

No Yes

No Yes

Occ Occ

Asia Asia

Oceania, Asia Asia, Oceania

Sorghum, maize, grasses Sorghum, grasses, fruit

No No

No No

Occ

E Hemisph.

Yes

Occ Sec

Asia W Hemisph.

No No

No Yes

Nola sorghiella Calocoris angustatus Dysdercus superstitiosus Eurystylus oldi (marginatus) Nysius raphanus Chlorochroa ligata Leptoglossus phyllopus

Occ Key Occ

Americas Asia Africa

S USA, C America E Hemisph. Africa

Sorghum, maize, cotton, tomato, many others Sorghum, grasses Sorghum, maize, cotton, tomato, legumes, many others Sorghum spp. Sorghum, grasses Sorghum, millet, grasses

No

Stenachroia elongella Helicoverpa zea

S Europe, Africa, SE Asia, Australia, New Zealand Asia Americas

No No No

No Yes No

Key

Africa

Africa

Grasses

No

Yes

Occ Occ Occ

Africa Americas Americas

Africa W Hemisph. W Hemisph.

No No No

No No No

Nezara viridula

Occ

Americas

Worldwide in warm areas

No

No

Oebalus pugnax

Occ

Americas

W Hemisph.

Sorghum, many others Sorghum, legumes Sorghum, legumes, cucurbits, pecan, persimmon 100+ hosts include sorghum, vegetables, legumes Grasses

No

No

Occ = occasional, Sec = secondary, Key = key pest.

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Sorghum are hosts of the insect. Adult females lay about 50 eggs (usually one per spikelet) between the glumes when sorghum spikelets are flowering. Larvae feeding cryptically on developing ovaries prevent normal kernel formation and cause direct grain loss. In contrast, greenbug is a holocyclic aphid that feeds on at least 60 grasses, including cultivated wheat Triticum aestivum L., oats Avena sativa L., barley Hordeum vulgare L., and rye Secale cereale L., during winter months, and sorghum during the summer. Parthenogenic females are mostly wingless. A greenbug produces about 80 young during a 3-week period. Ten to twelve generations can develop during the growing period of sorghum in the U.S. southern Great Plains, but usually after five or six generations greenbug abundance greatly declines because of parasitism. Greenbugs, feeding in colonies, extract copious amounts of sap from plant leaves and stems and inject a toxin as they feed that causes necrosis of leaf tissue. The aphid excretes honeydew and transmits important diseases such as maize dwarf mosaic virus. Biotypes of the species have been designated. While there are few prehistoric and historic facts about the origin of sorghum midge, there is certainly strong evidence to suggest that sorghum midge originated with sorghum in northeastern Africa.2,31-33,71 However, literature review of the dates of first discovery of sorghum midge in various sorghum-growing countries appears to suggest that after its discovery in Australia in 189472 and in the U.S. in 1895,7 it spread to Hawaii, the West Indies, Indonesia, and Sudan, and from Sudan gradually spread throughout Africa. Support for the contention that sorghum midge originated in Africa is as follows: 1. Examination in the Kew Herbarium and the herbarium of the British Museum of Natural History of botanical material of the genus Sorghum from Sudan revealed the presence of sorghum midge larvae and pupae in 45 samples which proved that sorghum midge was present in Sudan in 1869, 25 years before discovery in Australia or the U.S. 2. Only grasses of the genus Sorghum are hosts of the sorghum midge, and sorghums are indigenous to Africa. 3. Species diversity of the genus Stenodiplosis (formerly Contarinia) and perhaps associated parasitoids are apparently greater in Africa than in Australia or the U.S. 4. Many of the sources of plants resistant to sorghum midge are “zera zera” sorghums from Ethiopia and Sudan. 5. Sorghum midge is a much more severe pest in Australia and the U.S. than in northeastern Africa. There seems little doubt the sorghum midge is indigenous to Africa and an alien Australian, American, and Texan. The origin of the greenbug, like that of many insects that have become cosmopolitan in distribution, is unknown.67 General consensus is the insect is of Mediterranean (e.g., western or southwestern Eurasian) origin. The earliest available record of the insect is 1847 in Italy, where 5 years later the species was described.60 Soon after, in 1863, Passerini in Italy noted that sorghum was infested by greenbug.76 By © 1999 by CRC Press LLC

TABLE 4 History of Greenbug Biotypes and Some Ecological Characteristics Biotype

Ecological characteristics

A

Original population avirulent to “Dickinson Selection 28A” wheat; designated in 1961; cool season insect; rarely infested sorghum; currently, rarely naturally occurring Virulent to “Dickinson Selection 28A” wheat; designated in 1961; predominant biotype until late 1960s; cool season insect; “Piper” sudan grass was resistant; rarely infested sorghum; more common on grasses; currently, rarely naturally occurring Sorghum became important host; designated in 1968; predominant biotype until 1980s; became a warm and cool season insect; “Piper” sudan grass susceptible; resistant wheat and sorghum developed; naturally occurring Insecticide resistant, especially to organophosphates; designated in 1975; because this biotype is not based on virulence to plants, it often is omitted from a list of greenbug biotypes Virulent to most biotype C-resistant sorghum and wheat; designated in 1980; currently predominant biotype; resistant sorghum developed; still commonly occurring Damages Canada bluegrass; designated in 1986; somewhat similar to biotype A; not considered important to sorghum or wheat; naturally occurring Virulent to all resistant wheats; avirulent to sorghum; designated in 1988; not proven agriculturally important; naturally occurring Same host plant relationships as biotype E on wheat; avirulent to sorghum resistant to other biotypes; designated in 1988; virulent to “Post” barley; not proven agriculturally important Virulent to most biotype E-resistant sorghum in 1990, but not E-resistant wheat; designated in 1991; replacing biotype E; resistance identified in sorghum Avirulent to sorghum and some wheat and barley resistant to other biotypes; designated in 1994 from Idaho; damages “Post” barley; natural occurrence unknown Virulent to some sorghum sources resistant to biotype I; designated in 1996; some sources resistant to I retained resistance to K but at lower levels; ability to replace E and I unknown

B

C

D E F G H I J K

1907, the insect had been reported to occur in Europe, Asia, Africa, and North America.75 Greenbug now has spread into South America, but is not in Australia. In the U.S., the insect has infested small grains since about 1882.76 Infestations of johnsongrass, S. halepense (L.) Pers., in the U.S. were reported as early as 1909.76 Serious damage to sorghum occurred in Kansas in 1916.37 Sorghum became a primary cultivated summer host of greenbug in 1968 when outbreaks in sorghum occurred throughout most of the Great Plains region.36 The insect has remained a key insect pest of sorghum since 1968. Ten greenbug biotypes have been designated for their ability to damage resistant sorghum or wheat cultivars, seven of which have been identified within the last 16 years (Table 4).56 In contrast to the long evolutionary history shared by sorghum midge and sorghum, greenbug and sorghum have a relatively short history. Despite these different relationship histories, resistance in sorghum has been identified to both insects. Deployment of resistant sorghums has been more successful for greenbug than for sorghum midge. One reason is that genes for resistance to greenbug commonly have been simply inherited and mostly dominant, while genes for resistance to sorghum midge have been quantitatively inherited, mostly recessive, and complexly inherited.

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It is difficult to compare resistance mechanisms of sorghum midge and greenbug because of the differences in damage these insects cause to sorghum. Also, numerous greenbug biotypes complicate generalizations about resistance inheritance and mechanisms. The resistance to sorghum midge appears to be a non-preference response probably because of glume structure, time of day glumes close after flowering, and the tightness with which the glumes close.11,29,61,73,74 Greenbug resistance in sorghum probably is more related to the ability of the plant to tolerate enzymes such as pectinase secreted by the aphids as they suck juices from the plant.18 Resistance to these two insects is inherited differently and the mechanisms of resistance differ. However, it is difficult to relate these differences to evolutionary relationships of sorghum with sorghum midge or greenbug. One theory proposes that an insect with a long-term association with a host plant species has the genetic plasticity to overcome defense strategies of the host plant.35 That is, resistance in plants that evolved with an insect could be susceptible to being overcome by genetic changes in the insect pest. The theory that selection by insects causes plant populations to acquire resistance traits, and that selection imposed by these resistance traits provides the insect the ability to exploit these plants applies to one sorghum insect in these case studies. Sorghum midge is believed to have evolved with sorghum while greenbug did not. Popular theory would indicate that sorghum midge would constantly be changing to overcome resistance. However, no biotypes of sorghum midge have been reported, but many greenbug biotypes have been designated. Sorghum resistance to greenbug did not come from sources obtained through genetic feedback, but greenbug has the genetic plasticity to overcome resistance. Sorghum resistance to greenbug is qualitative, controlled by only one or a few genes that are mostly dominant. Commonly, resistant sorghums express all three resistance mechanisms, but tolerance resistance is the major mechanism and would have limited effect on greenbug biology. Resistance to sorghum midge that co-evolved with sorghum is believed to be quantitative, controlled by many genes that are mostly recessive and confer resistance to evasion by spikelets flowering when sorghum midges are not present, physical obstruction to sorghum midge oviposition in the spikelets, or antixenosis.11 Many other factors are associated with the ability of an insect to overcome plant resistance. Most effort to increase the durability of resistance has been on attempting to understand the genetics of the resistant plant. However, the genetics of the resistant plant is but one side of the issue. The genetics of the insect is the missing piece of the puzzle regarding the relationship of an insect with a resistant plant.

XI.

ECONOMIC BENEFIT OF INSECT-RESISTANT SORGHUM

The use of insect-resistant sorghum hybrids as a tactic in an integrated pest management approach functions to reduce insect pest abundance through antibiosis and antixenosis, and increases the ability of plants to withstand damage through tolerance. Intrinsic plant defense mechanisms have economical and ecological benefits. Often, only slight decreases in susceptibility of a crop cultivar to an insect pest enhance the effectiveness of other cultural control tactics such as crop rotation, host

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FIGURE 1

Effect of greenbug-resistant sorghum hybrids on insecticide usage in Texas.

destruction, and planting date adjustment; biological control, especially conservation of natural enemies; and insecticide control. Assessing the ecological benefits of insect pest-resistant cultivars by synergizing a multi-tactic management approach is difficult, and few such assessments have been made. Even determining the yield advantage from use of insect-resistant cultivars is difficult. The most commonly measured benefit to the use of insect-resistant crop cultivars is reduction in insecticide use. Although important, this benefit falls far short of the real economical and environmental benefits to the use of insect-resistant cultivars. Environmental benefits, such as cleaner water and safer food resulting from reduced insecticide use, must be tremendous, but largely are not measurable. Even the more direct benefits, such as reduced incidence of secondary pest outbreaks, are difficult to measure. Economic benefits have been assessed for sorghum resistant to some insect pests. In the U.S., economic assessments have been directed toward greenbug- and sorghum midge-resistant sorghum hybrids. Again, these evaluations are largely based on reduction in insecticide use because of use of insect-resistant sorghum hybrids. The effect of greenbug-resistant sorghum hybrids on insecticide use in Texas sorghum indicates how little insecticide was used before 1968 when greenbug became a serious pest of sorghum (Figure 1). In the mid-1960s, less than 10% of sorghum hectarage was treated with insecticide, but by the mid-1970s nearly 60% of the hectarage was treated. That trend in insecticide use continued until about 1977 when the first greenbug-resistant sorghum hybrids became commercially available. By 1980, hectarage of treated sorghum dropped from about 60 to 30% and has remained at about that level since. More than $9 million (U.S.) would be saved each year if 4 million acres in Texas were planted with resistant sorghum and insecticides costing $7.50 per acre were not needed to control greenbug. Eddleman et al.21 using a multicommodity agricultural sector model quantified the economic impart of greenbug-resistant sorghum hybrids. They reported development of greenbug-resistant sorghum hybrids resulted in $116 million in net gain © 1999 by CRC Press LLC

to U.S. consumer-taxpayers and a $273 million increase in benefit to producers for a total of $389 million net welfare to the U.S. under the existing farm program provisions. The annual rate of return on the research investment ($8.54 million) was 48.2%. Without farm commodity programs, the net social benefit to the U.S. would be $113 million, yielding an annual rate of return of 33.4% on the research investment. A similar, but less detailed cost/benefit analysis was conducted for sorghum midge-resistant sorghum hybrids.24 Ervin et al.23 calculated that for each dollar invested in research, the value of benefits to increased crop yields ranged from $24.20 to $2.70 depending on the discount rate. The average value of benefits was $9.90 for each dollar spent to develop the technology. This assessment would set the economic value of sorghum midge-resistant sorghum hybrids at $27 million.

XII.

MOLECULAR GENETIC TECHNIQUES FOR DEVELOPMENT OF INSECT-RESISTANT SORGHUM

Evolving molecular genetic techniques are providing exciting opportunities to better understand the genetic relationship between an insect and a resistant plant. This kind of information is needed especially for insects such as greenbug that have overcome several resistance genes in sorghum. DNA marker technology currently is used to map and identify gene structure conferring resistance traits in plants. Understanding the genetics of the resistance in plants will provide the knowledge to improve resistance deployment strategies. However, it is becoming clear that, with regard to plant resistance to insects, the genetics of the insect attacking the plant is equally, if not more, important to understand. Molecular genetic techniques are beginning to provide the opportunity to better understand the relationship of plant genetics to insect genetics. A detailed genetic map of the sorghum genome has been assembled.6 This map has been the basis to assess the magnitude of gene conservation among grass species54 and to study sorghum genetics of height and maturity,44 resistance to greenbug,43 and other important traits. Molecular genetic techniques also have been used to study the genetic structure of insect populations. Use of these techniques provides important clues to geographical variation, evolution of mating systems, heterozygosity, relatedness of individuals, and other factors that determine how insect populations respond to environmental conditions or human-imposed stresses, and set the framework for how insects evolve. Hoy39 reviewed selected examples of questions in insect population biology and ecology evaluated by the use of molecular techniques such as randomly amplified polymorphic DNA-polymerase chain reaction (RAPD-PCR), restriction fragment length polymorphism (RFLP) (ribosomal DNA, mitochondrial DNA, nuclear DNA), DNA fingerprinting, DNA sequencing, and dot blot hybridization. Powers et al.57 compared mtDNA restriction patterns to assess relationships among greenbug biotypes B, C, E, and F, and estimated a long period of time since these biotypes shared a common ancestral mitochondrial genome. Shufran et al.66 used DNA fingerprinting and found that almost every individual greenbug collected in any particular field on any one date had a different intergenic spacer (IGS) length; 13 major IGS size classes

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were present. Their data suggested that field populations of greenbug were comprised of many unique clones rather than a proliferation of one or a few. Better understanding the molecular bases of interactions of sorghum insect pests with resistant sorghum genotypes hopefully will point to gene deployment strategies to reduce the selective advantage favoring evolution of resistance-breaking biotypes. Many evolutionary concepts are based on the theory that substantial genetic variability exists in virulence of insect populations, i.e., greenbug, to resistant plants. The growing amount of evidence that grass species are genetically similar supports the possibility that genes for resistance to an insect pest could be conserved during the evolutionary divergence of host species. This issue has important implications to insect resistance in sorghum and raises the possibility that resistance genes to greenbug in sorghum may have been conserved from wheat, other small grains, or their progenitors. Traditional methods of assessing the genetics of insects, plants, and their interactions have not been adequate to explain evolutionary relationships, especially formation of biotypes virulent to resistant plant genotypes. Phylogenic analysis of data from comparative DNA sequence studies offers a methodology to reconstruct evolutionary history and relationships at all taxonomic levels. Molecular methods are providing reliable markers to distinguish between biotypes, races, and cryptic species of economically important insects39 — problems long resistant to solution, but central to the management of insect pests of sorghum.

XIII.

CONCLUSIONS

The genetic diversity of sorghum provides a wonderful opportunity to improve many agricultural properties of the cultivated crop, especially resistance to insects. The success in identifying sources of resistance to many of the insects infesting sorghum is a tribute to the genetic diversity of sorghum and to the efforts made by sorghum improvement specialists. However, identification of resistance sources is but the beginning of a long and difficult venture to deploy insect-resistant genotypes into agricultural production systems. The almost unimaginable ability of insects to overcome human-imposed defenses such as resistance will require a much better understanding of the genetic relationships between insects and resistant plants. Traditional entomological and plant breeding methods will continue to be important in this process, but molecular genetics techniques will greatly enhance traditional methods. Sorghum is genetically diverse, but so are the insects that infest the crop.

ACKNOWLEDGMENTS The authors are grateful to H. C. Sharma for information on sorghum germplasm evaluated for resistance to insect pests, especially stem borers and panicle-feeding bugs, and to Y. O. Doumbia for information on sorghum lines evaluated for resistance to Eurystylus oldi (marginatus). The U.S. Agency for International Development (Grant LAG-1254-G-00-6009-00), Pioneer Hi-bred International, and the National Grain Sorghum Producers Association are acknowledged for research support.

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REFERENCES 1. Ali, A. E. B., Identification of Sorghum Midge (Diptera: Cecidomyiidae) Resistant Lines Among Converted Exotic Sorghum, M.S. Thesis, Texas A&M University, College Station, 1986. 2. Barnes, H. F., Gall midges injurious to the sorghums, In: H. R. Barnes, Ed., Gall Midges of Economic Importance, Volume 7, Crosby Lockwood, London, 1956, 151. 3. Bennett, W. F., B. R. Tucker, and A. B. Maunder, Modern Grain Sorghum Production, Iowa State University Press, Ames, 1990. 4. Chang, C-P. and M-N. Fang, Studies on the resistance of sorghum variety to sorghum aphid, Melanaphis sacchari (Zehntner), Chinese J. Entomol., 4:97-105, 1984. 5. Chang, S-C., Sources of resistance in sorghum to sugarcane aphid Melanaphis sacchari (Zehntner), Rep. Corn Res. Center, Tainan DAIS, 15:11-14, 1981. 6. Chittenden, L. M., K. F. Schertz, Y-R. Lin, R. A. Wing, and A. H. Paterson, A detailed RFLP map of Sorghum bicolor x S. propinquum, suitable for high-density mapping, suggests ancestral duplication of sorghum chromosomes or chromosomal segments, Theor. Appl. Genet., 87:925-933, 1994. 7. Coquillett, D. W., A cecidomyiid injurious to seeds of sorghum, USDA Div. Entomol. Bull., 18:81-82, 1898. 8. Dahms, R. G. and J. B. Sieglinger, Reaction of sorghum varieties to the chinch bug, J. Econ. Entomol., 47:536-537, 1954. 9. Davis, F. H., G. L. Teetes, and J. W. Johnson, Yellow sugarcane aphid in sorghum, Sorghum Newsl., 21:112-113, 1978. 10. de Wet, J. M. and J. P. Huckabay, The origin of Sorghum bicolor, II, distribution and domestication, Evolution, 21:787-802, 1967. 11. Diarisso, N. Y., Spikelet Flowering Time and Morphology as Causes of Sorghum Resistance to Sorghum Midge, Ph.D. Dissertation, Texas A&M University, College Station, 1997. 12. Diawara, M. M., B. R. Wiseman, and D. J. Isenhour, Spodoptera frugiperda (Lepidoptera: Noctuidae) resistance in converted sorghum accessions: a research summary, Sorghum Newsl., 32:46-49, 1991. 13. Diawara, M. M., B. R. Wiseman, and D. J. Isenhour, Sorghum resistance to whorl feeding by larvae of the fall armyworm (Lep.: Noct.), J. Agric. Entomol., 9:41-53, 1992. 14. Diawara, M. M., B. R. Wiseman, D. J. Isenhour, and G. R. Lovell, Resistance to fall armyworm in converted sorghums, Fla. Entomol., 73:111-117, 1990. 15. Doggett, H., Sorghum, Longmans, Green and Co. Ltd., London, 1970. 16. Doggett, H., Sorghum, In: N. W. Simmonds, Ed., Evolution of Crop Plants, Longman, New York, 1976, 112. 17. Doggett, H., Sorghum, Longman Scientific and Technical, Harlow, Essex, 1988. 18. Dreyer, D. L. and B. C. Campbell, Association of the degree of methylation of intercellular pectin with plant resistance to aphids and with induction of aphid biotypes, Experimentia, 40:224-226, 1984. 19. Duncan, R. R., J. Dahlberg, and M. Spinks, International activities in sorghum germplasm acquisition during the past thirty-five years, In: International Germplasm Transfer: Past and Present, Special Publication 23, Crop Science Society of America, Madison, 1995, 117. 20. Eaton, A., Manual of Botany for the Northern and Middle States of America, Containing Generic and Specific Descriptions of Indigenous Plants and Common Cultivated Exotics, Growing North of Virginia, Websters and Skinners, Albany, 1922.

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21. Eddleman, B. R., C. C. Chang, and B. A. McCarl, Economic Benefits from INTSORMIL Grain Sorghum Variety Improvements in the United States, Tex. Agric. Exp. Stn. Report TAMRF 9045, Texas A&M University, College Station, 1991. 22. Ehrlich, P. R. and P. H. Raven, Butterflies and plants, Evolution, 18:586-608, 1964. 23. Ervin, R. T., T. M. Khalema, G. C. Peterson, and G. L. Teetes, Cost/benefit analysis of a sorghum hybrid resistant to sorghum midge (Diptera: Cecidomyiidae), Southwest. Entomol., 21:105-115, 1996. 24. FAO, FAO Production Yearbook, FAO Statistics Series, Food and Agriculture Organization of the United Nations, Rome, 1995. 25. Foster, D. G., Sorghums Resistant to the Banks Grass Mite, Oligonychus pratensis (Banks), M.S. Thesis, Texas A&M University, College Station, 1976. 26. Gebrekidan, B., Breeding sorghum for resistance to insects in Eastern Africa, Insect Sci. Appl., 6:351-357, 1985. 27. Guiragossian, V. and J. A. Mihm, Improving host-plant resistance to fall armyworm and sugarcane borer in sorghum, In: Proc. Int. Sorghum Entomology Workshop, 15-21 July 1984, Texas A&M University, College Station, ICRISAT, 1985, 201. 28. Hagio, T., M. Umehara, and S. Ono, Varietal reaction of sorghum to sugarcane aphid (Melanaphis sacchari Zehntner) in the seedling stage, Sorghum Newsl., 28:51-52, 1985. 29. Hallman, G. J., G. L. Teetes, and J. W. Johnson, Relationship of sorghum midge (Diptera: Cecidomyiidae) density to damage to resistant and susceptible sorghum hybrids, J. Econ. Entomol., 77:83-87, 1984. 30. Harlan, J. R., Crops and Man, American Society of Agronomy, Madison, 1975. 31. Harris, K. M., The sorghum midge, Contarinia sorghicola (Coq.), in Nigeria, Bull. Entomol. Res., 52:129-146, 1961. 32. Harris, K. M., The sorghum midge complex (Diptera: Cecidomyiidae): taxonomy, biology and assessments of field populations, Bull. Entomol. Res., 63:305-325, 1963. 33. Harris, K. M., The sorghum midge: a review of published information, 1895-1983, Proc. Int. Sorghum Entomology Workshop, 15-21 July 1984, Texas A&M University, College Station, ICRISAT, 1985, 229. 34. Harris, M. K., Allopatric resistance: searching for sources of insect resistance for use in agriculture, Environ. Entomol., 4:661-669, 1975. 35. Harris, M. K., Arthropod-plant interactions related to agriculture, emphasizing host plant resistance, In: M. K. Harris, Ed., Biology and Breeding for Resistance to Arthropods and Pathogens in Agricultural Plants, Tex. Agric. Exp. Stn., MP-1451, Texas A&M University, College Station, 1980, 23. 36. Harvey, T. L. and H. L. Hackerott, Recognition of a greenbug biotype injurious to sorghum, J. Econ. Entomol., 62:776-779, 1969. 37. Hayes, W. P., Observations on insects attacking sorghums, J. Econ. Entomol., 15:349-356, 1922. 38. Howitt, A. J. and R. H. Painter, Field and Greenhouse Studies Regarding the Sources and Nature of Resistance of Sorghums, Sorghum vulgare Pers., to the Corn Leaf Aphid, Rhopalosiphum maidis (Fitch), Agric. Exp. Stn., Tech. Bull. 82, Kansas State College of Agriculture and Applied Science, Manhattan, 1956. 39. Hoy, M. A., Insect Molecular Genetics: An Introduction to Principles and Applications, Academic Press, San Diego, 1994. 40. Jermy, T., Evolution of insect/host plant relationships, Am. Nat., 124:609-630, 1984. 41. Johnson, J. W., D. T. Rosenow, and G. L. Teetes, Resistance to the sorghum midge in converted exotic sorghum cultivars, Crop Sci., 13:754-755, 1973.

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42. Johnson, J. W., G. L. Teetes, and D. T. Rosenow, Evaluation of sorghum hybrids for resistance to the corn leaf aphid, Sorghum Newsl., 18:91, 1975. 43. Katsar, C. S., A. H. Paterson, G. C. Peterson, and G. L. Teetes, Use of molecular genetics to improve greenbug resistance in sorghum, In: Proc. 20th Biennial Grain Sorghum Res. and Utilization Conf., New Orleans, 1997, 71. 44. Lin, Y-R., K. F. Schertz, and A. H. Paterson, Comparative analysis of QTLs affecting plant height and maturity across the Poaceae, in reference to an interspecific sorghum population, Genetics, 141:391-411, 1995. 45. Mann, J. A., C. T. Kimber, and F. R. Miller, The Origin and Early Cultivation of Sorghums in Africa, Tex. Agric. Exp. Stn. Bull., Texas A&M University, College Station, 1454, 1983. 46. Manthe, C. S., Sorghum Resistance to the Sugarcane Aphid (Homoptera: Aphididae), Ph.D. Dissertation, Texas A&M University, College Station, 1992. 47. Martin, J. H., Sorghum Improvement, USDA Agricultural Yearbook, U.S. Government Printing Office, Washington, D.C., 1936. 48. Martin, J. H., History and classification of sorghum, Sorghum bicolor (Linn.) Moench, In: J. S. Wall and W. M. Ross, Eds., Sorghum Production and Utilization, Avi Publishing, Westport, 1970, 1. 49. McColloch, J. W., The corn leaf aphid (Aphis maidis Fitch) in Kansas, J. Econ. Entomol., 14:89-94, 1921. 50. Mize, T. W. and G. Wilde, New resistant germplasm to the chinch bug (Heteroptera: Lygaeidae) in grain sorghum: contribution of tolerance and antixenosis as resistance mechanisms, J. Econ. Entomol., 79:42-45, 1986. 51. Mote, U. N., M. D. Shinde, and D. R. Bapat, Screening of sorghum collections for resistance to aphids and oily malady of winter sorghum, Sorghum Newsl., 28:13, 1985. 52. Murdock, G. P., Africa: Its People and Their Culture History, McGraw-Hill, New York, 1959. 53. Nwanze, K. F., N. Seetharama, H. C. Sharma, and J. W. Stenhouse, Biotechnology in pest management: improving resistance in sorghum to insect pests, African Crop Sci. J., 3:209-215, 1995. 54. Paterson, A. H., Y-R. Lin, Z. Li, K. F. Schertz, J. F. Doebley, S. R. M. Pinson, S-C. Liu, J. W. Stansel, and J. E. Irvine, Convergent domestication of cereal crops by independent mutations at corresponding genetic loci, Science, 269:1714-1718, 1995. 55. Pi, C-P. and J-S. Hsieh, Preliminary studies on aphid resistance in sorghum, Natl. Sci. Counc. Monthly, 10:153-160, 1982. 56. Porter, D. R., J. D. Burd, K. A. Shufran, J. A. Webster, and G. L. Teetes, Greenbug (Homoptera: Aphididae) biotypes: selected by resistant cultivars or preadapted opportunists?, J. Econ. Entomol., 90:1055-1065, 1997. 57. Powers, T. O., S. G. Jensen, S. D. Kindler, C. J. Stryker, and L. J. Sandall, Mitochondrial DNA divergence among greenbug (Homoptera: Aphididae) biotypes, Ann. Entomol. Soc. Am., 82:298-302, 1989. 58. Quinby, J. R., Sorghum Improvement and the Genetics of Growth, Texas A&M University Press, College Station, 1974. 59. Quinby, J. R., N. W. Kramer, J. C. Stephens, K. A. Lahr, and R. E. Karper, Grain Sorghum Production in Texas, Tex. Agric. Exp. Stn. B-912, Texas A&M University, College Station, 1958. 60. Rondani, C., Aphis graminum n. sp., Nuov. Ann. Sci. Nat. Bologna, 6:9-11, 1852.

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61. Rossetto, C. J., Tipos de Resistencia de Sorgo, Sorghum bicolor (L.) Moench, a Contarinia sorghicola (Coquillett, 1898), Tese de Livre Docente, Universidade Estadual Paulista “Julio de Mesquita Filho,” Jaboticabal, 1977. 62. Schertz, K. F., Biology of sorghum, In: M. K. Harris, Ed., Biology and Breeding for Resistance to Arthropods and Pathogens in Agricultural Plants, Tex. Agric. Exp. Stn. MP-1451, Texas A&M University, College Station, 1980, 124. 63. Sharma, H. C., Host plant resistance to insects in sorghum and its role in integrated pest management, Crop Prot., 12:11-34, 1993. 64. Sharma, H. C. and V. F. Lopez, Genotypic resistance in sorghum to head bug, Calocoris angustatus Lethiery, Euphytica, 58:193-200, 1992. 65. Sharma, H. C., S. L. Taneja, K. Leuschner, and K. F. Nwanze, Techniques to Screen Sorghum for Resistance to Insect Pests, ICRISAT Information Bull. 32, Andhra Pradesh, 1992. 66. Shufran, K. A., W. C. Black IV, and D. C. Margolies, DNA fingerprinting to study spatial and temporal distributions of an aphid, Schizaphis graminum (Homoptera: Aphididae), Bull. Entomol. Res., 81:303-313, 1991. 67. Starks, K. J. and Z. B. Mayo, Jr., Biology and control of the greenbug attacking sorghum, In: Proc. Int. Sorghum Entomology Workshop, 15-21 July 1984, Texas A&M University, College Station, ICRISAT, Andhra Pradesh, 1985, 149. 68. Starks, K. J. and K. A. Mirkes, Yellow sugarcane aphid: plant resistance in cereal crops, J. Econ. Entomol., 72:486-488, 1979. 69. Starks, K. J. and D. E. Weibel, Sorghum cultivars rated for resistance to chinch bug, Sorghum Newsl., 25:82-83, 1982. 70. Stephens, J. C., F. R. Miller, and D. T. Rosenow, Conversion of alien sorghums to early combine genotypes, Crop Sci., 7:396, 1967. 71. Teetes, G. L., Entomology of johnsongrass/sorghum/sorghum midge and agriculture, In: M. K. Harris and C. E. Rogers, Eds., The Entomology of Indigenous and Naturalized Systems in Agriculture, Westview Press, Boulder, 1988, 125. 72. Tryon, H., The insect enemies of cereals belonging to the genus Cecidomyia, Trans. Nat. Hist. Soc. Queensland, 1:80-83, 1895. 73. Waquil, J. M., G. L. Teetes, and G. C. Peterson, Adult sorghum midge (Diptera: Cecidomyiidae) nonpreference for resistant hybrid sorghum, J. Econ. Entomol., 79:455-458, 1986. 74. Waquil, J. M., G. L. Teetes, and G. C. Peterson, Sorghum midge (Diptera: Cecidomyiidae) adult ovipositional behavior on resistant and susceptible sorghum hybrids, J. Econ. Entomol., 79:530-532, 1986. 75. Webster, F. M., Investigations of Toxoptera graminum and its parasites, Ann. Entomol. Soc. Am., 2:67-87, 1909. 76. Webster, F. M. and W. J. Phillips, The spring grain-aphis or 'green bug', USDA Bur. Entomol. Bull., 110:1-153, 1912. 77. Webster, J. A., Yellow sugarcane aphid (Homoptera: Aphididae): detection and mechanisms of resistance among Ethiopian sorghum lines, J. Econ. Entomol., 83:1053-1057, 1990. 78. Wiseman, B. R. and G. R. Lovell, Resistance to the fall armyworm in sorghum seedlings from Ethiopia and Yemen, J. Agric. Entomol., 5:17-20, 1988. 79. Wuensche, A. L., An Assessment of Plant Resistance to the Sorghum Midge, Contarinia sorghicola, in Selected Lines of Sorghum bicolor, Ph.D. Dissertation, Texas A&M University, College Station, 1980.

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Section II Legume Crops

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5

Bean Germplasm Resources for Insect Resistance Cesar Cardona and Julia Kornegay

CONTENTS I. Introduction II. Genetic Resources III. Host Plant Resistance for Major Pests A. Leafhoppers B. Bean Stem Maggot C. Bean Pod Weevil D. Mexican Bean Weevil and Bean Weevil E. Mexican Bean Beetle F. Other Arthropods IV. Conclusions References

I. INTRODUCTION The common bean, Phaseolus vulgaris L., is the world’s most important edible foodlegume (Table 1). Approximately 18 mmt of dry beans and 3 mmt of snap beans are produced annually. Nearly 80% of dry bean production occurs in developing countries on small-scale farms. Latin America, the center of Phaseolus domestication, produces nearly half of the world’s supply of dry beans. The principle producing countries or regions are Brazil, Mexico, and Central America. Africa, with nearly 4 million ha of annual bean production, is a secondary center for bean genetic diversity. The major producers are Kenya, Zaire, Tanzania, Uganda, Burundi, Rwanda, and Ethiopia. A significant amount of dry beans are also grown in Eastern Europe and North America. More than 50,000 ha of snap beans are grown annually in China.60 Beans play an increasingly important role in human nutrition in Africa where they are the second most important source of dietary protein and the third most important source of calories.60 Beans are ranked fourth as a protein source in Latin America.

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TABLE 1 The Global Importance of Common Beans as Compared to Other Edible Food Legumes Food legume

Production (mt)

Percent (%)

Common bean Chickpea Faba bean Lentil Cowpea Total

17,973,300 8,036,670 3,773,040 2,772,860 2,532,270 35,088,140

51.2 22.9 10.8 7.9 7.2 100.0

From FAOSTAT (Food Agriculture Organization of the United Nations, Production Statistics), on the World Wide Web (http://apps.fao.org), 1997. (Ref. 25)

TABLE 2 Status of the Phaseolus Bean Collection Available at CIAT Species Domesticated P. vulgaris P. lunatus P. coccineus P. polyanthus P. acutifolius Other wild speciesa Total a

Wild

Cultivated

Total accessions

856 149 69 2 145

24,532 1,527 529 290 127

25,388 1,676 598 292 272 167 28,393

Includes 21 species.

From CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1996. (Ref. 21)

II.

GENETIC RESOURCES

The largest Phaseolus germplasm collection is housed at the Centro Internacional de Agricultura Tropical (CIAT), Cali, Colombia. Out of 28,000 available accessions, approximately 25,000 are cultivated and wild P. vulgaris; the remaining 3,000 accessions are domesticated and wild species of Phaseolus (Table 2). More than 60 countries have donated Phaseolus germplasm to the CIAT collection since its formation in 1969, with the most important collections coming from Mexico, Honduras, Costa Rica, U.S., Colombia, Peru, Ecuador, and England.34 Another major Phaseolus collection (13,467 accessions) is maintained by the U.S. Department of Agriculture, Agricultural Research Service, Western Regional Plant Introduction © 1999 by CRC Press LLC

Station (WRPIS), Pullman, Washington.82 Other repositories at the N. I. Vavilov Institute, Russia, the Instituto Nacional de Investigaciones Agrícolas (INIA), Mexico, and the University of Cambridge, U.K., store and maintain sizeable collections.34

III.

HOST PLANT RESISTANCE FOR MAJOR PESTS

More than 100 insects and other invertebrate pests have been recorded on beans. Of these, about 14 are major pests of the crop.13 Whiteflies Bemisia tabaci (Gennadius) and Trialeurodes vaporariorum (Westwood), leafhopper Empoasca kraemeri Ross & Moore, chrysomelids Diabrotica balteata LeConte and Cerotoma facialis Erickson, Mexican bean weevil Zabrotes subfasciatus (Boheman), and bean weevil Acanthoscelides obtectus (Say) are widely distributed and important in Latin America. Bean pod weevil Apion godmani Wagner, Mexican bean beetle Epilachna varivestis Mulsant, and slug Vaginulus plebeius (Fisher) are important in parts of Mexico and Central America. In the U.S., Mexican bean beetle, seedcorn maggot Delia platura Meigen, and mites Tetranychus spp. are the main pests of beans.67 Bean stem maggot or bean fly, Ophiomyia spp., is the key pest in Africa, followed by chrysomelid Ootheca bennigsenni Weise, pod borer Maruca testulalis (Geyer), aphids Aphis craccivora Koch and A. fabae Scopoli, and bruchids.1 The economic importance of the invertebrate species affecting beans varies widely between and within regions. In spite of their impact on yield and crop quality, this crop has been neglected and only in recent times has there been a concerted effort to breed for resistance to insects. Knowledge on mechanisms of resistance, inheritance, and biochemical basis of resistance is limited to a few major pests and breeding efforts are limited to five species. It is unfortunate that no useful sources of resistance have been reported for several economically important polyphagous pests such as whiteflies, chrysomelids, leafminers, thrips, and pod borers (Table 3). Bean cropping systems and insect control tactics vary widely, from total reliance on natural control in most of Africa to excessive use of insecticides in the hillsides of the Andes.1,20 For most bean farmers who cannot afford costly pesticides, or do not know how to use them, host plant resistance is the best method of insect control. Plant resistance in beans and other smallholder crops should receive more attention. Pesticide use in the developing world is increasing and there is growing awareness of human health hazards and environmental degradation.5,78 It was initially thought that increases in Latin America were mainly caused by the growth of plantation crops, an important source of export revenue.5 However, the developing world’s share of the world agrochemical usage, currently valued at $10.6 billion (U.S.) is forecast to rise from 19% to an estimated 35% by the year 2000.78 Much of this growth stems from a wider and more intensive use of chemical protection by smallholder farmers. Extensive use of pesticides by many bean farmers in Central and South America began after the mid-1970s.70 Recent surveys by CIAT and national agricultural research organizations in this region show that many bean growers have adopted the risky and environmentally degrading practice of conducting weekly, and often unnecessary, insecticidal sprays with up to 22 applications in 90 days.19 Given this situation, and because biological control and cultural practices have limited scope in © 1999 by CRC Press LLC

TABLE 3 Major Arthropod Pests of Beans and their Present Status in Terms of Development of Host Plant Resistance Resistance Insecta Bean pod weevil Bean stem maggot Bean weevil Chrysomelids Helicoverpa Leafhopper Leafminers Lygus bug Mexican bean beetle Mexican bean weevil Red mite Seed corn maggot Thrips Whiteflies a b

Sources

Mechanism

Inheritance

Breeding methodsb

Yes Yes Yes No No Yes No Yes Yes Yes Yes Yes No Yes?

Antibiosis, antixenosis Tolerance, antixenosis Antibiosis — — Tolerance, antixenosis — Antibiosis, antixenosis Tolerance, antixenosis Antibiosis Tolerance Unknown — —

Two genes Polygenic (?) Two genes — — Polygenic — Unknown Unknown Monogenic Unknown Unknown — —

P P BC — — RS — None None BC None None — —

Scientific names in text. BC, backcross; P, pedigree; RS, recurrent selection.

beans,70 the need for a more sustained effort in developing insect-resistant cultivars is evident. In this chapter, we summarize and discuss present knowledge on host plant resistance to major insect pests of beans.

A. LEAFHOPPERS The leafhopper E. kraemeri is the most important insect pest of common beans in Latin America and southern Florida.70,73 In North America, E. fabae Harris is the principal species attacking beans.63,69 Under early infestations, yield losses to E. kraemeri feeding can be as high as 95%.70 Thousands of bean germplasm accessions from the CIAT, WRPIS, and other repositories have been screened for resistance to leafhoppers. Low levels of resistance to E. fabae were detected after screening over a thousand bean varieties, breeding lines, and WRPIS accessions in the late 1950s and early 1960s in the U.S.80 Additional screening for E. fabae resistance was conducted in North America.46,69 When several thousand accessions from the CIAT Phaseolus collection were evaluated for resistance to E. kraemeri, only 4% of 18,000 accessions were resistant.43 High levels of resistance to E. kraemeri also have been identified in P. acutifolius A. Gary20,21 and P. lunatus L.49 accessions. Tolerance to feeding damage is the most common mechanism of resistance to leafhoppers in common beans.42,80 Electronic feeding studies showed that leafhopper probing and feeding behavior on a tolerant cultivar (EMP 84) was less damaging to

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the plant than that exhibited on a susceptible cultivar.8 This tolerance may be related to a unique behavior adjustment of the leafhopper, which leads to less feedinginduced damage. In addition, free and no-choice feeding and ovipositional experiments identified low to moderate levels of nonpreference (antixenosis) to leafhopper oviposition in some cultivars.42,43,79 Although Pillemer and Tingey63 found that hooked trichomes reduced E. fabae nymphal populations on two bean cultivars, this resistance mechanism does not affect E. kraemeri. Resistance to E. kraemeri is quantitatively inherited.27,44 Additive and dominance gene effects were significant for plant yield in three resistant lines under leafhopper attack, but the magnitude of the dominance gene effects was greater than the additive effect.44 For nymphs per leaf, only additive gene effects were significant in two lines, EMP 89 and EMP 94, having moderate levels of antixenosis to leafhopper oviposition. Transgressive segregation for higher yield under leafhopper attack was found in F2 populations combining antixenosis and tolerance resistance mechanisms. More than 300 E. kraemeri-resistant breeding lines have been developed as a result of breeding for bean resistance to this pest at CIAT. Based on the results of the inheritance and combining ability studies, a modified recurrent selection program was developed to increase bean yields under leafhopper attack.39 The advantage of this breeding strategy is that it increases the frequencies of genes with additive effects which may be at a low level initially in the population. Selection can be delayed within populations until later generations when the genotypes are relatively fixed. Genotype by environment interactions are also minimized by the repeated evaluation and yield testing of breeding populations over time. The modified recurrent selection breeding program has been successful in increasing overall tolerance to leafhopper feeding damage and in incorporating resistance into a wide variety of bean market types (Table 4).17 By the sixteenth cycle of recurrent selection, there was significant progress in increasing resistance within small, red-seeded lines, a group that had been difficult to improve in earlier cycles. Breeding lines that were developed using a pedigree breeding strategy had non-protected yields 10 to 16% below that of “ICA Pijao,” the standard commercial check cultivar, while lines developed using the modified recurrent selection breeding strategy had yields ranging 9 to 17% above “ICA Pijao.”39 The most resistant lines have leafhopper action threshold levels equivalent to seven nymphs per leaf compared with the susceptible check cultivar with three nymphs per leaf.11,18 Some of E. kraemeri-resistant lines developed at CIAT also show high levels of tolerance to E. fabae in Canada.69 Of these, several small, white-seeded EMP lines show the greatest promise as they combine resistance to leafhoppers, high yields, and good canning qualities required by farmers and consumers.

B. BEAN STEM MAGGOT The bean stem maggot or bean fly is one of the most important pests of common bean in Africa, Asia, and Australia, causing complete loss of bean seedlings and more mature plants in certain areas.22,31,47,55,77 It has not been reported in the Americas. Three species of bean stem maggot attack beans (1) O. phaseoli (Tryon), (2) O. spencerella Greathead, and (3) O. centrosematis de Meijere. In Africa,

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TABLE 4 Best Carioca, Cream, and Red-Seeded Bean Lines from the XVII Cycle of Recurrent Selection for Leafhopper Resistance Yield (kg/ha) Line Carioca EMP 413 EMP 412 EMP 407 EMP 332b Cream EMP 392 EMP 400 EMP 394 EMP 267c “ICA Pijao” d BAT 41 e Red EMP 385 EMP 375 EMP 378 EMP 335f “ICA Pijao” d BAT 41e

Protected

Unprotected

Susceptibility index

% Gain in tolerance a

2750 2388 2171 2099

1924 1750 1597 1459

0.77 0.79 0.84 0.88

33.6 31.9 27.6 24.1

2487 2180 2398 1985 2315 1849

1865 1708 1631 1261 867 276

0.67 0.69 0.76 0.89 1.15 1.51

41.2 39.4 33.3 21.9 — —

2890 2196 2342 2380 2326 1993

2442 2014 2024 1543 1077 269

0.67 0.70 0.73 1.01 1.33 1.92

49.6 47.3 45.1 24.2 — —

Adapted from CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1992. (Ref. 17) a With respect to the tolerant check (“ICA Pijao”). b Improved Carioca check. c Improved Cream check. d Tolerant check. e Susceptible check. f Improved Red check.

O. phaseoli is the most important species. However, O. phaseoli and O. spencerella were reported as equally important pests at two sites in Kenya, but their prevalence depended on environmental conditions.55 The major damage to a bean plant is caused by third instar larvae, which destroy the medullar tissue of the stem at ground level. The concentration of puparia in the stems causes swelling, splitting, and rotting of the base of affected plants. Infested bean seedlings suffer premature leaf fall and are either killed or severely stunted.1 Evaluations of bean cultivars for resistance to bean stem maggot have been done in Ethiopia, Malawi, Tanzania, Uganda, and Taiwan.16,21,31,37 Msangi and Karel54 reported that cultivars with high numbers of leaf trichomes had lower oviposition punctures in Tanzania. Low O. phaseoli larval and pupal counts were also correlated with narrow stem diameters. Nevertheless, few of the reportedly resistant cultivars

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have been consistently resistant to bean stem maggot. In addition, the CIAT core collection of P. vulgaris was field evaluated in Tanzania to identify new sources of resistance to this insect.21 Several germplasm accessions from Africa, Mexico, Guatemala, and Peru exhibited high levels of resistance in the initial screenings. Evaluation of the core collection is continuing in Tanzania and Ethiopia. Tolerance to stem damage caused by pupating larvae is reported as a mechanism of resistance to O. phaseoli. Low egg counts were associated with high leaf pubescence, thin stems, and long internodes.37,65 Other studies indicate that nonpreference for oviposition, differential larval survival and development, and tolerance may be involved in resistance to bean stem maggot.21 Because of difficulties in identifying good sources of resistance to bean stem maggot, only limited breeding work has been undertaken for this pest. When crosses were made between “Ikinimba” and “ZPV 292” and local Tanzanian susceptible lines, the progeny from “Ikinimba” as a parent had generally less plant mortality in both generations than progeny from “ZPV 298.” However, both sets of crosses were less affected by the insect than the susceptible local cultivar. Presently, bean populations among the eight best resistant sources produced via random mating are under field evaluations in Tanzania.21

C. BEAN POD WEEVIL The bean pod weevil, A. godmani, is a very important pest of beans in Mexico and parts of Central America.50-51 Oviposition takes place within newly formed pods during daytime and neonate larvae penetrate the pod wall. Second and third instars feed on developing seeds and pupation takes place within the mature pod. As a result of larval feeding, seed damage can be as high as 90% in some areas of Mexico and Central America.4,10 Sources of resistance to the pod weevil were identified in Mexico by Guevara32 and in El Salvador by Mancía.50 Other highly resistant germplasm accessions were reported in cultigens4,29,30 and in wild P. vulgaris.2 Most resistant germplasm are landraces from the highlands of Mexico and therefore are not well adapted to the tropical conditions of Central America.4 Ovipositional antixenosis and antibiosis have been suggested as possible mechanisms of resistance to pod weevil, with the latter more important and common in resistant accessions. In hypersensitivity, the resistant accessions respond to oviposition by forming a callus that impedes larval penetration of the pod wall. As a result, neonate larvae cannot reach the developing seed and starve to death.28 The biochemical basis of resistance to this insect is unknown. Studies on the inheritance of antibiosis resistance suggest that two genes segregating independently are responsible for the resistance. One gene, Agm, has no effect when present alone, whereas the other gene, Agr, alone is capable of conferring intermediate resistance. Resistance is higher when both genes are present.29 Resistant cultivars were developed in Mexico32 and probably in El Salvador50 during the 1960s. Pod weevil-resistant cultivars developed by CIAT in collaboration with national programs in the 1970s were poorly adapted to conditions in Central America and had unacceptable seed characteristics. Not until new and better adapted

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resistance sources were incorporated into a pedigree program4 was progress made in developing cultivars with combined resistance to bean common mosaic virus and bean pod weevil. These lines yielded as well or better than local cultivars in the absence of the insect, and better than local cultivars when the pod weevil was present in high numbers. However, they were susceptible to bean golden mosaic virus when epidemics of this virus occurred in Central America in 1983. The present breeding scheme for pod weevil resistance includes the simultaneous screening for resistance to bean common and bean golden mosaic viruses, anthracnose, and pod weevil.

D. MEXICAN BEAN WEEVIL

AND

BEAN WEEVIL

The Mexican bean weevil, Z. subfasciatus, and the bean weevil, A. obtectus, are two important bruchid species affecting stored beans throughout much of the world. The larvae bore into the seeds and render them unfit for human consumption. Losses caused by bruchids in beans have been estimated at 13%.70 After screening more than 8,000 germplasm accessions and finding no usable resistance sources,71 Schoonhoven et al.72 detected high levels of resistance to both weevil species in 10 accessions (G12862B, G12866, G12891, G12929, G12931, G12942, G12949, G12952, G12953, PI325690) from a small collection of wild P. vulgaris from the states of Nayarit, Jalisco, Colima, Michoacan, Mexico, and Guerrero in Mexico. To date, resistance to either bruchid has not been detected in wild P. vulgaris accessions from other countries or regions. Antibiosis is responsible for resistance in wild bean accessions to both Z. subfasciatus and A. obtectus.15,72 Resistance has a significant impact on adult emergence, days to adult emergence, survival of late first and early second instar larvae, and adult size. Resistance to both bruchids has also been found in P. acutifolius and P. lunatus.20,21,23,74 Arcelin, a major storage seed protein, and present only in resistant wild Mexican accessions, is responsible for resistance to Z. subfasciatus.41,56 The protein is inherited as a single dominant gene, Arc,66 and there are six allelic variants48,57,68 which are codominant. Purified arcelin inhibits the development of Z. subfasciatus larvae in artificial seeds.14,56 The mode of action of arcelin is unknown but the high dose indicates that it is probably not digested well by the insect.52 The protein does not confer resistance to A. obtectus.40,56 Recent work by Fory et al.26 confirmed the importance of arcelin in resistance to Z. subfasciatus and demonstrated that α-amylase inhibitors (α-AIs) would, at best, play a minor role as resistance factors. The biochemical basis of resistance to A. obtectus is unknown. A successful backcross breeding method14,56 has incorporated resistance to Z. subfasciatus in bean cultivars by a combination of biochemical tests for the presence of arcelin and insect feeding bioassays. More than 100 arcelin-1 and arcelin2 containing lines have been delivered to national programs in Africa and Latin America.20,21 In breeding for resistance to Z. subfasciatus, it is interesting to note that arcelin sources (variants) were ranked for resistance as 5>4>1>2>6>314,21 and backcross lines obtained with variants 1-5 were ranked as 1>2>5>3>4 (Table 5). The unique nature of arcelin-1, which contains both dimer and tetramer proteins,58 may be responsible for the higher levels of resistance obtained by using this variant in breeding. Although the value of arcelin-based resistance to Z. subfasciatus has

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TABLE 5 Levels of Resistance to Zabrotes subfasciatus in Arcelin Donor Parents and Selected Resistant Bean Lines Derived from Crosses % emergence Arcelin variant

Donor parent

Breeding line

Donor parent

Breeding line

9.3 20.5 79.5 8.4 4.0 52.0 97.3

1.8 27.0 72.1 92.1 32.1 —a —

51.0 48.8 42.0 70.0 64.6 45.6 31.3

53.0 42.9 35.6 40.9 62.9 — —

Arcelin 1 Arcelin 2 Arcelin 3 Arcelin 4 Arcelin 5 Arcelin 6 “Ica Pijao” b a b

Days to adult emergence

No lines containing arcelin-6 have been developed. Commercial cultivar used as standard susceptible check.

Modified from Cardona, C., et al., Comparative value of four arcelin varients in the development of dry bean lines resistant to the Mexican bean weevil, Entomol. Exp. Appl., 56:197-206, 1990; and CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1996. (Refs. 14 and 21)

been demonstrated in laboratory and field tests12,61 and several national programs have tested resistant lines, this resistance has not reached farmers in Latin America in the form of weevil-resistant cultivars. This is unfortunate because savings in insecticide costs to small farmers could reach hundreds of millions of U.S. dollars annually. The inheritance of resistance to A. obtectus is more complicated.40 Delay in adult emergence was inherited as two recessive genes, whereas reduction of emergence was lost in the progeny. The pattern of resistance could reflect the action of several segregating genes. Additional work on the inheritance of resistance to this species has not been published. And little progress has been made in breeding for resistance to A. obtectus, largely because only a very few of the F2 segregants from resistant parents show acceptable resistance levels and there is a tendency for resistance to decrease as generations are advanced.40

E. MEXICAN BEAN BEETLE The Mexican bean beetle, E. varivestis, primarily a pest of soybeans, attacks beans in the U.S., Mexico, and Central America.10 Damage is caused by both larvae and adults feeding on the leaves and occasionally on stems and pods. Damage is more serious in the highlands of Mexico where yield losses can be as high as 35%.59 Research has identified few sources of resistance to Mexican bean beetle. For example, Benepal6 detected resistance in less than 5% of 7,800 P. vulgaris accessions from the WRPIS collection. Early studies by Wolfenbarger and Sleesman,81 Campbell and Brett9, and Lapidus et al.45 suggested antixenosis and possibly antibiosis as

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mechanisms of resistance. In Mexico, “Guanajuato 18” and “Zacatecas 48” (P. vulgaris) and “Puebla 86” (P. coccineus) exhibited low levels of antibiosis or antixenosis resistance.53 Also, Pacheco and Serrano59 detected low levels of resistance in 123 snap bean cultivars in Mexico.

F. OTHER ARTHROPODS There are a number of other insect and mite pests of beans that are economically important in specific countries and regions. Searches of germplasm for resistance to these insects have been undertaken, but no organized breeding for plant resistance has been done. Evaluations for resistance to the seed corn maggot Delia platura (Meigen),33,76 the chrysomelid Ootheca bennigsennis Weise,38 and tetranychid mites,35,36 have not shown marked differences in varietal and accession susceptibility. Other insects studied include the pod borer, Etiella zinckenella (Treitschke) in Brazil64 and Lygus hesperus Knight in the U.S.3,24 There are reports from Puerto Rico on resistance to Bemisia tabaci (Westwood), a major pest of the crop in Latin America. According to Blair and Beaver7 and Peña et al.,62 antixenotic resistance (reduced oviposition rates) may exist in a few bean lines bred for resistance to bean golden mosaic virus. If confirmed, and if the levels of white fly resistance are high enough, this could be an important finding.

IV.

CONCLUSIONS

We have seen herein that genetic variability exists in beans for resistance to insects. The importance of large germplasm collections is exemplified by the finding of high levels of resistance to bruchids in wild P. vulgaris accessions72 and the detection of valuable sources of resistance to bean pod weevil in landraces from Mexico.4,29,51 The search for new sources of resistance to insects in beans must continue, with emphasis on leafhoppers, whiteflies, and beanflies. Knowledge of plant resistance mechanisms is limited to a few major insect pests.15,28,42,43,79 A better understanding of the mechanisms of resistance to beanfly and Mexican bean weevil is needed. There is also need for more information on the biochemical nature of resistance and inheritance of resistance to several major insect pests. These must be understood in great detail before we can develop adequate breeding strategies. The importance of this kind of research is exemplified by the use of arcelin as a biochemical marker for breeding for resistance to the Mexican bean weevil.14 Current efforts in trying to develop molecular markers for bean pod weevil resistance83 and bean weevil26 must be encouraged. Sources of resistance to leafhoppers and bean weevil in P. acutifolius are being used in inter-specific crosses with embryo rescue techniques.84 Overcoming barriers to inter-specific crosses will also facilitate the exploitation of high levels of resistance to leafhopper49 and bean weevil20,21,74 detected in P. lunatus. Good progress has been made in developing cultivars that are resistant to a single pest.4,14,39 If insect resistance breeding is to contribute toward stability of bean production, and serve as the foundation of integrated pest management systems, current efforts in breeding for multiple insect and disease resistance21,75 need to be encouraged.

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REFERENCES 1. Abate, T. and J. K. O. Ampofo, Insect pests of common beans in Africa, Ann. Rev. Entomol., 41:45-73, 1996. 2. Acosta, J., R. Rosales, and R. Garza, Resistance to the pod weevil in wild Phaseolus vulgaris, Annu. Rep. Bean Improv. Coop., 35:103-104, 1992. 3. Alvarado-Rodríguez B., T. F. Leigh, K. W. Foster, and S. S. Duffey, Resistance in common bean (Phaseolus vulgaris) to Lygus hesperus (Heteroptera: Miridae), J. Econ. Entomol., 79:484-489, 1986. 4. Beebe, S., C. Cardona, O. Díaz, F. Rodríguez, E. Mancía, and S. Ajquejay, Development of common bean (Phaseolus vulgaris L.) lines resistant to the pod weevil, Apion godmani Wagner, in Central America, Euphytica, 69:83-88, 1993. 5. Bellotti, A. C., C. Cardona, and S. L. Lapointe, Trends in pesticide use in Colombia and Brazil, J. Agric. Entomol., 7:191-201, 1990. 6. Benepal, P. S., Screening beans (Phaseolus vulgaris L.) for resistance to Mexican bean beetle (Epilachna varivestis Mulsant), Ann. Rep. Bean Improv. Coop., 19:17-18, 1976. 7. Blair, M. W. and J. S. Beaver, Resistance to the sweetpotato whitefly (Bemisia tabaci), the vector of bean golden mosaic virus in dry beans (Phaseolus vulgaris), Ann. Rep. Bean Improv. Coop., 35:154-155, 1992. 8. Calderón, J. D. and E. A. Backus, Comparison of the probing behaviors of Empoasca fabae and E. kraemeri (Homoptera: Cicadellidae) on resistant and susceptible cultivars of common beans, J. Econ. Entomol., 85:88-89, 1992. 9. Campbell, W. V. and C. H. Brett, Varietal resistance of beans to the Mexican bean beetle, J. Econ. Entomol., 59:899-902, 1966. 10. Cardona, C., Insects and other invertebrate bean pests in Latin America, In: H. F. Schwartz and M. Pastor-Corrales, Eds., Bean Production Problems in the Tropics, Centro Internacional de Agricultura Tropical, CIAT, Cali, 1989, 505. 11. Cardona, C. and M. L. Cortés, Evaluación económica de la tolerancia de variedades de frijol al lorito verde, Empoasca kraemeri Ross & Moore (Homoptera: Cicadellidae), Rev. Col. Entomol., 17:19-23, 1991. 12. Cardona, C., K. Dick, C. E. Posso, K. Ampofo, and S. M. Nahdy, Resistance of a common bean (Phaseolus vulgaris L.) cultivar to post-harvest infestation by Zabrotes subfasciatus (Boheman) (Coleoptera: Bruchidae), II. Storage tests, Trop. Pest Manag., 38, 173-175, 1992. 13. Cardona, C. and K. K. Karel, Key insect and other invertebrate pests of beans, In: S. R. Singh, Ed., Insect Pests of Legumes, John Wiley, Chichester, 1990, 157. 14. Cardona, C., J. L. Kornegay, C. E. Posso, F. Morales, and H. Ramírez, Comparative value of four arcelin variants in the development of dry bean lines resistant to the Mexican bean weevil, Entomol. Exp. Appl., 56:197-206, 1990. 15. Cardona, C., C. E. Posso, J. L. Kornegay, J. Valor, and M. Serrano, Antibiosis effects of wild dry bean accessions on the Mexican bean weevil and the bean weevil (Coleoptera: Bruchidae), J. Econ. Entomol., 82:310-315, 1989. 16. CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1983. 17. CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1992. 18. CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1993. 19. CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1994.

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20. CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1995. 21. CIAT (Centro Internacional de Agricultura Tropical), Bean Program Ann. Rep., Cali, 1996. 22. Davis, J. J., Bean fly and its control, Queensl. Agric. J., 95:101-106, 1969. 23. Dobie, P., J. Dendy, C. Sherman, J. Padgham, J. A. Wood, and A. M. R. Gatehouse, New sources of resistance to Acanthoscelides obtectus (Say) and Zabrotes subfasciatus Boheman (Coleoptera: Bruchidae) in mature seeds of five species of Phaseolus, J. Stored Prod. Res., 26:177-186, 1990. 24. Dutton, A. C., S. R. Temple, C. G. Summers, D. Helms, A. Hall, and J. Small, Evaluation of germplasm accessions and advanced progenies for response to Lygus hesperus (Knight), Univer. Calif. Dry Bean Res., 1990 Progress Rep., University of California, Davis, 1990. 25. FAOSTAT (Food Agriculture Organization of the United Nations, Production Statistics), on the World Wide Web (http://apps.fao.org/), 1997. 26. Fory, L. F., F. Finardi-Filho, C. M. Quintero, T. C. Osborn, C. Cardona, M. J. Chrispeels, and J. E. Mayer, α-Amylase inhibitors in resistance of common beans to the Mexican bean weevil and the bean weevil (Coleoptera: Bruchidae), J. Econ. Entomol., 89:204-210, 1996. 27. Galwey, N. W. and Al. M. Evans, The inheritance of resistance to Empoasca kraemeri Ross and Moore in the common bean, Phaseolus vulgaris L., Euphytica, 31:933-952, 1982. 28. Garza, R., Identificación de los componentes de la resistencia al ataque del picudo del ejote, Apion godmani, que poseen los materiales seleccionados, In: Informe de Investigación 1991 del Proyecto Apion en México. SARH-INIAP-CEVAMEX, Chapingo, 1992, 13. 29. Garza, R., C. Cardona, and S. P. Singh, Inheritance of resistance to the bean-pod weevil (Apion godmani Wagner) in common beans from Mexico, Theor. Appl. Genet., 92:357-362, 1996. 30. Garza, R. and J. S. Muruaga, Resistencia al ataque del picudo del ejote Apion spp en frijol Phaseolus spp, Agron. Mesoam., 4:77-80, 1993. 31. Greathead, D. J., A study in East Africa of beanflies (Diptera: Agromyzidae) affecting Phaseolus vulgaris and their natural enemies, with the description of a new species of Melanagromyza Hend, Bull. Entomol. Res., 59:541-561, 1968. 32. Guevara, J., El combate del picudo del ejote mediante la combinación de variedades resistentes e insecticidas, Agric. Tec. Mex., 1:17-19, 1961. 33. Hagel, G. T., D. W. Burke, and K. J. Silbernagel, Response of dry bean selections to field infestations of seedcorn maggot in central Washington, J. Econ. Entomol., 74:441-443, 1981. 34. Hidalgo, R., CIAT's world Phaseolus collection, In: A. van Schoonhoven and O. Voysest, Eds., Common Beans: Research for Crop Improvement, CAB International, Wallingford, 1991, 163. 35. Impe, G. van and T. Hance, 1993, Une technique d'evaluation de la sensibilité varietale au tetranyque tisserand, Tetranuchus urticae Koch (Acarina: Tetranychidae), Application au haricot, au concombre, a la tomate et au fraisier, Agronomie, 13:739-749, 1993. 36. Jara, B., C. Acosta, and C. Cardona, Efecto de cinco variedades de frijol sobre la biología y la fecundidad de la arañita roja, Tetranychus desertorum Banks (Acari, Tetranychidae), Rev. Col. Entomol., 7:33-39, 1981.

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37. Karel, A. K. and A. P. Maerere, Evaluation of common bean cultivars for resistance to bean fly (Ophiomyia phaseoli Tryon), Ann. Rep. Bean Improv. Coop., 28:1153-1158, 1985. 38. Karel, A. K. and C. L. Rweyemamu, Resistance to foliar beetle, Ootheca bennigseni (Coleoptera: Chrysomelidae) in common beans, Environ. Entomol., 14:662-664, 1985. 39. Kornegay, J. L. and C. Cardona, Development of an appropriate breeding scheme for tolerance to Empoasca kraemeri in common bean, Euphytica, 47:223-231, 1990. 40. Kornegay, J. L. and C. Cardona, Inheritance of resistance to Acanthoscelides obtectus in a wild common bean accession crossed to commercial bean cultivars, Euphytica, 52:103-111, 1991. 41. Kornegay, J. L., C. Cardona, and C. E. Posso, Inheritance of resistance to Mexican bean weevil in common bean, determined by bioassay and biochemical tests, Crop Sci., 33:589-594, 1993. 42. Kornegay, J. L., C. Cardona, and A. van Schoonhoven, The mechanisms of resistance in common beans to the leafhopper Empoasca kraemeri, Entomol. Exp. Appl., 40:272-279, 1986. 43. Kornegay, J. L., C. Cardona, J. van Esch, and M. Alvarado, Identification of common bean lines with ovipositional resistance to Empoasca kraemeri (Homoptera: Cicadellidae), J. Econ. Entomol., 82:649-654, 1989. 44. Kornegay, J. L. and S. R. Temple, Inheritance and combining ability of leafhopper defense mechanisms in common bean, Crop Sci., 26:1153-1158, 1986. 45. Lapidus, J. B., R. W. Cleary, R. H. Davidson, F. W. Fisk, and M. G. Augustine, Chemical factors influencing host selection by the Mexican bean beetle, Epilachna varivestis Muls., J. Agric. Food Chem., 1:462-462, 1963. 46. Lindgren, D. T. and D. P. Coyne, Injury and yield of leafhopper-infested dry beans, J. Am. Soc. Hort. Sci., 120:839-842, 1995. 47. Lin, C. S. and W. C. Mitchell, Host selection of the bean fly [Ophiomyia phaseoli (Tryon)], A. J. Taiwan Mus., 34:233-236, 1981. 48. Lioi, L. and R. Bollini, Identification of a new arcelin variant in wild bean seeds, Ann. Rep. Bean Improv. Coop., 32:28, 1989. 49. Lyman, J. M. and C. Cardona, Resistance in lima beans to a leafhopper, Empoasca kraemeri, J. Econ. Entomol., 75:281-286, 1982. 50. Mancía, J. E., Evaluación de variedades de frijol tolerantes al picudo de la vaina Apion godmani (Wagn.), SIADES (El Salvador), 2:15-20, 1973. 51. McKelvey, J. J., J. Guevara, and A. Cortés, Apion pod weevil: a pest of beans in Mexico, J. Econ. Entomol., 40:476-479, 1947. 52. Minney, B. H. P., A. M. R. Gatehouse, P. Dobie, J. Dendy, C. Cardona, and J. A. Gatehouse, Biochemical bases of seed resistance to Zabrotes subfasciatus (bean weevil) in Phaseolus vulgaris (common bean); a mechanism for arcelin toxicity, J. Insect Physiol., 36:757-767, 1990. 53. Montalvo, C. G. and C. Sosa, Evaluación de la resistencia de frijol hacia la conchuela Epilachna varivestis Muls. (Coleoptera: Coccinellidae), Agrociencia, 10:3-13, 1973. 54. Msangi, R. B. and A. K. Karel, Host plant resistance in common beans to bean fly (Ophiomyia phaseoli Tryon) In: N. Menjas and M. P. Selema, Eds., Proc. 4th Workshop on Bean Res. in Tanzania, Sokoine Univ. of Agriculture, Morogoro, 1986, 60. 55. Nderitu, J. H., H. Y. Kayumbo, and J. M. Mueke, Beanfly infestation on common beans (Phaseolus vulgaris L.) in Kenya, Insect Sci. Applic., 11:35-41, 1990. 56. Osborn, T. C., D. C. Alexander, M. S. Sun, C. Cardona, and F. A. Bliss, Insecticidal activity and lectin homology of arcelin seed protein, Science, 240:207-210, 1988.

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57. Osborn, T. C., T. Blake, P. Gepts, and F. A. Bliss, Bean arcelin 2. Genetic variation, inheritance and linkage relationships of a novel seed protein of Phaseolus vulgaris L., Theor. Appl. Genet., 71:847-855, 1986. 58. Osborn, T. C., M. D. Burow, and F. A. Bliss, Purification and characterization of arcelin seed protein from common bean, Plant Physiol., 86:399-405, 1988. 59. Pacheco, P. J. and L. M. Serrano, Selección de genotipos de frijol (Phaseolus vulgaris L.) para resistencia horizontal a Epilachna varivestis Mulsant (Coleoptera: Coccinellidae), Rev. Chapingo, 16:18-21, 1992. 60. Pachico, D., The demand for bean technology, In: G. Henry, Ed., Trends in CIAT Commodities, CIAT Working Doc. No. 128, Centro Internacional de Agricultura Tropical, Cali, 1993, 60. 61. Padgham, J., V. Pike, K. Dick, and C. Cardona, Resistance of a common bean (Phaseolus vulgaris L.) cultivar to post-harvest infestation by Zabrotes subfasciatus (Boheman) (Coleoptera: Bruchidae). I. Laboratory tests, Trop. Pest Manag., 38:167-172, 1992. 62. Peña, E. A., A. Pantoja, J. Beaver, and A. Armstrong, Oviposición de Bemisia tabaci Genn. (Homoptera: Aleyrodidae) en cuatro genotipos de Phaseolus vulgaris L. (Leguminosae) con diferentes grados de pubescencia, Folia Entomol. Mex., 87:1-12, 1993. 63. Pillemer, E. A. and W. M. Tingey, Hooked trichomes: a physical barrier to a major agricultural pest, Science, 193:482-484, 1976. 64. Ramalho, F. S., M. M. de Albuquerque, and R. C. Machado, Comportamento de linhagens e variedades de feijao (P. vulgaris L.) em relacao a Etiella zinckenella Treitschke, Rev. Agric. Recife, 53:171-178, 1978. 65. Rogers, D. J., Host plant resistance to Ophiomyia phaseoli (Tryon) (Diptera: Agromyzidae) in Phaseolus vulgaris, J. Austr. Entomol. Soc., 18:245-250, 1979. 66. Romero-Andreas, J., B. S. Yandell, and F. A. Bliss, Bean arcelin. 1. Inheritance of a novel seed protein of Phaseolus vulgaris L. and its effect on seed composition, Theor. Appl. Genet., 72:123-128, 1986. 67. Ruppel, R. F., Insects and bean pests: an annotated listing, Mich. Dry Bean Dig., 4:28-29, 1980. 68. Santino, A., B. Valsasina, L. Lioi, A. Vitale, and R. Bollini, Bean (Phaseolus vulgaris L.) seed lectins: novel electrophoretic variant of arcelin, Plant Physiol. (Life Sci. Adv.), 10:7-11, 1991. 69. Schaafsma, A. W. and T. Michaels, Resistance of common bean, Phaseolus vulgaris, to potato leafhoppers, Empoasca fabae (Harris), Ann. Rep. Bean Improv. Coop., 39:84-85, 1996. 70. Schoonhoven, A. van and C. Cardona, Insects and other bean pests in Latin America, In: H. F. Schwartz and G. Galvez, Eds., Bean Production Problems: Disease, Insect, Soil and Climatic Constraints of Phaseolus vulgaris, Centro Internacional de Agricultura, CIAT, Cali, 1980, 363. 71. Schoonhoven, A. van and C. Cardona, Low levels of resistance to the Mexican bean weevil in dry beans, J. Econ. Entomol., 75:567-569, 1982. 72. Schoonhoven, A. van, C. Cardona, and J. Valor, Resistance to the bean weevil and the Mexican bean weevil (Coleoptera: Bruchidae) in noncultivated common bean accessions, J. Econ. Entomol., 76:1255-1259, 1983. 73. Schoonhoven, A. van, G. J. Hallman, and S. R. Temple, Breeding for resistance to Empoasca kraemeri Ross and Moore in Phaseolus vulgaris L., In: L. R. Nault and J. G. Rodriguez, Eds., Leafhoppers and Planthoppers, Wiley, New York, 1985, 405.

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74. Shade, R. E., R. C. Pratt, and M. A. Pomeroy, Development and mortality of the bean weevil, Acanthoscelides obtectus (Coleoptera: Bruchidae), on mature seeds of tepary beans, Phaseolus acutifolius, and common beans, Phaseolus vulgaris, Environ. Entomol., 16:1067-1070, 1987. 75. Singh, S. P., C. Cardona, F. Morales, M. Pastor-Corrales, and O. Voysest, Gamete selection for upright carioca beans with resistance to leafhopper and four diseases, Ann. Rep. Bean Improv. Coop., 39:86, 1996. 76. Vea, E. V. and C. J. Eckenrode, Seedcorn maggot injury, New York's Food and Life Sci. Bull., 55:3, 1976. 77. Wallace, G. B., French bean diseases and bean fly in East Africa, East Afr. For. J., 7:170-175, 1939. 78. Whitaker, M. J., The challenge of pesticide education and training for tropical smallholders, Int. J. Pest Manag. 39:117-125, 1993. 79. Wilde, G. and A. van Schoonhoven, Mechanisms of resistance to Empoasca kraemeri in Phaseolus vulgaris, Environ. Entomol., 5:251-255, 1976. 80. Wolfenbarger, D. and J. P. Sleesman, Resistance in common bean lines to the potato leafhopper, J. Econ. Entomol., 54:846-849, 1961. 81. Wolfenbarger, D. and J. P. Sleesman, Resistance to the Mexican bean beetle in several bean genera and species, J. Econ. Entomol., 54:1018-1022, 1961. 82. Clement, S.L., Personal communication, 1997. 83. CIAT, unpublished. 84. Cardona, C., unpublished.

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6

Assessment of Soybean Germplasm for Multiple Insect Resistance David J. Boethel

CONTENTS I. Introduction A. Background and Importance B. Germplasm Resources C. Plant Resistance for Soybean Insect Management II. Insect-Resistant Germplasm A. Resistance Related to Pubescence B. Multiple Insect-Resistant Germplasm C. Mechanisms of Resistance D. Inheritance of Resistance E. Genotype by Environment Interactions F. Compatibility with Other Management Tactics G. Released Insect-Resistant Cultivars H. Biotechnological Advances III. Conclusions References

I.

INTRODUCTION

The role of plant resistance in the management of soybean insect pests can be characterized by two events. The first event was the fortuitous discovery over 60 years ago that plant pubescence conferred resistance to the potato leafhopper, Empoasca fabae (Harris).50,53 Incorporation of this trait into most commercial soybean cultivars resulted in population suppression that has remained stable over the years and virtually relegated the insect to non-pest status on the crop. The second event occurred in the late 1960s when soybean germplasm was identified with resistance to foliar feeding by the Mexican bean beetle, Epilachna varivestis Mulsant.144,145 Subsequent studies with three plant introductions (PIs), PI229358 (“Sodendaizu”), PI171451 (“Kosamame”), and PI227687 (“Miyako White”), revealed that these genotypes demonstrated resistance to multiple insect pests of

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soybean. These PIs became the models for research on inheritance of insect resistance, mechanisms of resistance, compatibility of plant resistance with other management tactics, and genotype by environment interactions. They were frequently used as donor parents in breeding programs in the U.S. and other countries to develop insect-resistant germplasm and cultivars.

A. BACKGROUND

AND IMPORTANCE

The soybean is a member of the family Leguminosae and is thought to be native to Eastern Asia, probably originating in north and central China.104 There are 18 species in the genus Glycine Willd., which contains two subgenera. The wild perennial soybeans are placed in the subgenus Glycine while the cultivated soybean G. max (L.) Merr., and its wild ancestor G. soja (Sieb. and Zucc.) belong to the subgenus Soja. Glycine max and G. soja have similar genomes, are cross-compatible, and usually produce vigorous fertile F1 hybrids, whereas the 16 species of wild perennial soybeans have widely diverse genomes.100 Although soybean was introduced into North America in 1765, it remained a relatively minor crop (810,000 ha in 1925), and until 1940 more soybeans were grown for forage than for seed.75 In the U.S., soybean hectarage grew rapidly during the 1960s and 1970s to a peak of 28.9 million ha in 1979. Although hectarage declined to a low of 23.4 million ha in 1990, soybean was grown on 25.3 million ha on 380,000 farms in 1995. Hectarage has increased each of the last 6 years with estimates for 26 million ha in 1997.2 Soybeans are an important source of protein for humans and livestock, and soy oil dominates the edible oil market. The U.S. is the world leader in soybean production with 58.6 mmt accounting for 46% of the world’s supply in 1995.2 Brazil (19%), China (11%), and Argentina (10%) also are major producers; however, the U.S. clearly leads in world trade (72%). The crop has become an important component of the nation’s export program.127 In 1992, soybean export sales were approximately $4.1 billion (U.S.), which according to U.S. Department of Agriculture (USDA) estimates, translates into 123,000 jobs in the U.S. Therefore, soybean is an important source of on-farm income and a major contributor to the general economy through the export market.

B. GERMPLASM RESOURCES Cultivar development has played an important role in establishing the soybean as a major crop in the U.S.42 and other countries.100 Yield has been the primary focus of breeding efforts, but genetic improvement for yield has been accompanied by a loss in genetic diversity. In North America, 10 PIs have contributed more than 80% to the northern gene pool with the same percentage associated with only seven PIs in the southern gene pool.26 The breeding strategy has been to use germplasm to obtain genes for quantitative traits.100 Then, recombinant phenotypes are identified from segregating progenies that possess the most favorable genes contributed by the elite parent, plus a few favorable genes contributed by the PI. The use of PIs, directly or as parents in controlled hybridization, has been a major factor in the development of commercial soybean cultivars.51 © 1999 by CRC Press LLC

The PIs used in cultivar development can be found in germplasm collections worldwide, and because the only survey of collections is a decade old, the exact number of accessions is not known. It is estimated that G. max accessions exceed 100,000, G. soja 10,000, and perennial Glycine species 3,500.100 Soybean germplasm can be categorized based on response to photoperiod which controls flowering and maturity. Photoperiod is controlled by latitude, and germplasm is placed in one of 13 maturity groups (MG) (000 through X, north to south in the northern hemisphere) depending on its adaption to a narrow band of latitude.134 The USDA Soybean Germplasm Collection is located at the University of Illinois, Urbana-Champaign. The collection contains 1,104 accessions of G. soja with most accessions from South Korea, Japan, Russia, and China. The accessions from South Korea primarily fall into MG V, Japan MGs IV through VII, Russia MGs 000 through II, and China MGs 000 through V. The same countries lead in the number of accessions of G. max with approximately 87% of the 16,581 accessions originating in China, Korea, Japan, and Russia. Of the G. max accessions, 81% are in MGs I through VII with the remaining germplasm adapted for more extreme northern (000-0) and southern (VIII-X) latitudes. Approximately 828 accessions of the wild perennial Glycine species are in the USDA Collection, while 2,500 accessions are in the Australian collection at Canberra. Although these wild soybeans have not been used extensively in soybean breeding programs because of difficulty of crossing, wild perennial germplasm has been identified with resistance to several pathogens and with tolerance to salt and certain herbicides.100 Chapter 12 in this volume has additional information on soybean germplasm holdings.

C. PLANT RESISTANCE

FOR

SOYBEAN INSECT MANAGEMENT

Paralleling the dramatic increase in soybean production in the Western Hemisphere in the early 1960s was the rapid development and implementation of integrated pest management (IPM) programs to deal with the complex of native and migratory insect fauna that had the potential to attack the crop. The expansion of soybean was dramatic in the southern U.S., an area which, because of its subtropical climate, was at a comparatively high hazard from pests.96 The scientists developing these IPM systems realized that profitable soybean production would not be possible if the crop required as much use of insecticides as that routinely used on other southern crops such as cotton Gossypium hirsutum L. Also, in the southern U.S. where insect problems were accentuated, soybeans were grown on land recently cleared of forests, permanent pastures, and temporary grazing land. These areas were near streams, lakes, ponds, and woodlands, which were sensitive to adverse effects of heavy insecticide use. These situations along with others, such as low value per hectare, made the prospects of insect-resistant soybean cultivars an attractive management tactic. In many early reviews describing the status and progress of insect management on soybean,64,68,96,97,138,140 plant resistance was touted as having the potential to be the most important management tactic. Efforts to develop advanced soybean breeding lines with multiple insect resistance and good agronomic qualities became commonplace in the U.S., with resistance breeding programs underway in 10 states in 1987.141 The primary emphasis

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was directed at foliage feeding pests130 with less focus on stink bugs, while the opposite occurred in breeding programs in Brazil.65 In Asia, agromyzid bean flies and pod borers were the targets of plant resistance studies.24,36 Periodically, progress in soybean plant resistance research has been reviewed in papers at the World Soybean Research Conference.65,117,130,142 From these reports, it is apparent that excellent sources of resistance (especially to major defoliators) exists, that germplasm with resistance to multiple insect pests is available, and that advanced breeding lines incorporating the resistant germplasm exhibit good agronomic traits, except the most important facet, yield. Although the advanced breeding lines had performed well in yield trials at different geographic locations, it was suggested that yield should be equal to or better than existing cultivars before release. Several reports indicated that private breeding programs included insect resistance as an objective and expressed optimism that multiple pest-resistant cultivars, to include resistance to insects, would become available as a component in pest management programs. Kogan65 acknowledged that genotypes resistant to several major pests had been identified and that breeding programs in various parts of the world were incorporating genes from these genotypes into advanced agronomic germplasm. He urged that progress must be made in the identification of chemical resistance mechanisms and their genetic control to take advantage of classical breeding and genetic engineering to produce high yielding pest-resistant cultivars. A recent review by Lambert and Tyler82 acknowledged that progress had been made toward development of acceptable insect-resistant cultivars, but that success had not been achieved. The lower yield potential of the insect-resistant cultivars that have been released, coupled with the late maturity of most of these cultivars, have limited the adoption by soybean producers. Lambert and Tyler82 suggested that the release of inferior cultivars and breeding lines should cease and a concerted effort initiated to develop high-yielding, highly resistant genotypes to overcome this. It was stressed that this new approach was imperative because of the economic and environmental benefits associated with the adoption of resistant cultivars. Earlier, Todd et al.137 had indicated that insect-resistant cultivars were not a major management tactic in soybean IPM programs but offered great potential for increasing profitability. Listed among the probable economic benefits were fewer insect outbreaks and higher economic thresholds resulting in reduced insecticide use and costs, reduced environmental contamination by insecticides, and increased insecticide efficacy and reduced insecticide rates on resistant cultivars. It is widely recognized that plant resistance offers excellent opportunities for reduced economic input, a situation especially attractive on a low value per hectare crop such as soybean. Undoubtedly, this factor has been instrumental in continuation of the plant resistance effort on soybean despite the limited success seen over the past 25 years in the release of insect-resistant cultivars.

II.

INSECT-RESISTANT GERMPLASM

Some level of insect resistance probably was selected “inadvertently” when agronomically superior cultivars were selected from PIs introduced from Asia and released to producers.65 Those early cultivars continue to be used as parents in current © 1999 by CRC Press LLC

breeding programs, and most commercial cultivars grown in the U.S. have normal pubescence, which imparts resistance to potato leafhopper. Indeed, pubescent germplasm and commercial cultivars have demonstrated resistance to potato leafhopper for over half a century. Other sources of insect resistance were found only after extensive screening of available germplasm.

A. RESISTANCE RELATED

TO

PUBESCENCE

Although Poos and Smith102 found that oviposition by potato leafhopper was greater, nymphal survival higher, and nymphal development time reduced on the non-pubescent soybean PI55069 compared with cultivars “Dixie” and “Herman” bearing hairyrough pubescence, they were not convinced the pubescence was the determining factor in resistance. Their skepticism resulted from field observations over three years, wherein potato leafhoppers were equally abundant on hairy and less hairy cultivars. Johnson and Hollowell53 were more convinced that the mechanical protection of dense rough-hairy pubescence was responsible for the resistance. They examined the progeny of crosses of rough-hairy pubescent and dominant glabrous soybean. Both the homozygous and heterozygous glabrous progeny were heavily infested with potato leafhoppers, severely stunted in growth, and had curled leaves with yellow necrotic margins (“hopperburn”151). The progeny with rough-hairy pubescence showed none of the symptoms, which lead to the conclusion that resistance was due to pubescence or a character conditioned by the same genes as pubescence. Wolfenbarger and Sleesman150 evaluated 16 soybean genotypes for resistance to potato leafhopper and found that only the cultivars and lines with normal, dense, or semi-appressed pubescence were resistant to nymphs. Because a genotype with sparse pubescence exhibited resistance to hopperburn, the authors felt that factors other than leaf pubescence were partly responsible for resistance. Wolfenbarger and Sleesman150 used strains with aberrant pubescence derived primarily from Asian germplasm to study different pubescence types. Bernard and Singh11 used the strains in crosses with cultivars (“Clark” and “Harosoy”) with normal pubescence to determine the inheritance of pubescence type in soybean. Each of the four types that originated in Asia differed from normal pubescence by one gene. The development of near isogenic lines differing in pubescence has helped researchers understand the role of pubescence in insect resistance. In Illinois, the cultivars “Clark” and “Harosoy” and their isolines were examined for resistance to potato leafhopper.119 In both the “Clark” and “Harosoy” genetic backgrounds, the curly pubescent and especially the glabrous plants were heavily infested and demonstrated severe stunting and hopperburn. Level of infestation was inversely proportional to height and level of pubescence, and this relationship was apparent across both cultivars. It was not determined if the differences in height in the lightly infested lines were due to potato leafhoppers, the action of genes closely linked with the pubescence genes, or the result of pleiotrophic effects of the pubescence genes themselves. In two studies conducted in South Carolina to examine isolines of the “Lee” cultivar and two PIs for resistance to the potato leafhopper, the glabrous and deciduous

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isolines were the most susceptible. Among pubescent isolines and PIs, resistance was related to orientation and size of the trichomes. The lowest numbers of potato leafhoppers were found on PI229358, a genotype having long, erect trichomes, slightly lower in density than the “Lee” isoline with normal pubescence.138,139 Broersma et al.18 also reported that glabrous PIs were susceptible to potato leafhopper. In the pubescent genotypes, the number of trichomes did not appear to be a major factor, but orientation, size, and other characteristics were significant in determining resistance. Isolines also have been used to study the influence of pubescence on several of the major lepidopterous pests of soybean. Results of the research indicate that resistance to larvae and adult oviposition preference are not positively correlated. In studies with the corn earworm, Helicoverpa zea (Boddie), and lines isogenic for dense, normal, and no pubescence developed from the cultivar “Tracy M,” Lambert and Kilen81 found that the average oviposition was 60.8, 29.9, and 9.3% on the dense, normal, and glabrous isolines, respectively. Although more eggs were laid on the dense and normal isolines, the fruiting structures were more preferred as oviposition sites on the glabrous isoline. In research of a similar nature, Lambert et al.76 used “Tracy M” lines isogenic for dense, normal, and glabrous pubescence to examine the effects on larval development of the corn earworm, velvetbean caterpillar, Anticarsia gemmatalis Hübner, and soybean looper, Pseudoplusia includens (Walker), and the oviposition preference of the latter two species. In concurrent studies with the soybean looper and isolines of the same pubescence type developed from the cultivar “Davis,” the presence of pubescence resulted in reduced larval growth, extended larval development time, and increased oviposition by adults. The authors indicated that the increased larval damage on the glabrous plants was more than offset by the reduction in oviposition on those plants. The conclusions drawn were that pubescence of soybean functions as a resistance mechanism to the larvae of lepidopterous defoliators but enhances adult oviposition.76 Beach and Todd6 came to a similar conclusion after studying the oviposition preferences of soybean looper on four soybean genotypes that differed in larval resistance. The beet armyworm, Spodoptera exigua (Hübner), response to pubescence on “Davis” isolines (see Reference 76) was opposite that seen for other lepidopterans. In the field, more beet armyworm larvae were found on glabrous isolines than those having dense or normal pubescence.152 Because beet armyworm females oviposit their eggs in clusters, it was suggested that they may have difficulty laying eggs on leaves with trichomes. In Asia, the soybean pod borer, Leguminivora glycinivorella Matsumura, and the limabean pod borer, Etiella zinchenella (Trictsche), are pests of soybean pods and pubescence influences their oviposition and larval feeding behavior. The soybean pod borer preferred to oviposit on soybean pods containing pubescence in China.36 On glabrous lines, oviposition occurred on plant parts other than the pods, and larvae suffered mortality while searching for or attempting to bore into pods. Of 3,000 germplasm accessions screened for resistance to soybean pod borer, 123 were highly resistant. Damage by limabean pod borer increased with increasing trichome density on pods suggesting that glabrous cultivars could be highly resistant in Taiwan.133

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After screening nearly 7,000 germplasm accessions in Taiwan, Chiang and Talekar24 identified four G. soja accessions as highly resistant to three species of agromyzid beanflies. One mechanism of resistance, trichome length and density on the under surface of leaves of vegetative stage soybean, was found to deter oviposition by the beanflies, especially Melanagromyza soja (Zehntner) and Ophiomyia centrosematis (de Maijere).21,22 Gai et al.32 confirmed these findings with M. soja. The accessions with semi-erect pubescence were more resistant, indicating that density of pubescence played a role in resistance to agromyzid beanflies. Whiteflies also are influenced by soybean pubescence. Arioglu et al.3 used nine isolines of the cultivar “Clark” that varied in pubescence density and type to study resistance to sweetpotato whitefly, Bemisia tabaci (Germadius), a major pest of soybean in Turkey. The glabrous isoline, L62-1385, was highly resistant while the isolines with dense pubescence were very susceptible. Because some of the isolines and cultivars that exhibit semi-dense pubescence also had small infestations of sweetpotato whitefly, it was suggested that pubescence length may be as important as density. Lambert et al.73 reported that soybean genotypes with increased trichome density tended to have higher populations of silverleaf whitefly, Bemisia argentifolii Bellows & Perring, and bandedwing whitefly, Trialeurodes abutilonea (Haldeman). However, genotypes that had the trichomes lying flat on the leaf surface appeared to be the most resistant. Two commercial cultivars, “Perrin” and “Cook,” were among the most resistant entries, allowing for immediate incorporation in existing soybean IPM programs. McAuslane et al.88 observed that greater populations of immature silverleaf whitefly occurred in pubescent and hirsute soybean than in glabrous soybean. Other studies with isogenic lines developed from a normal pubescent genotype with multiple insect resistance (D75-1016960) revealed that the partial resistance of glabrous soybean seen in the field resulted from reduced oviposition preference related to lack of foliar pubescence.87

B. MULTIPLE INSECT-RESISTANT GERMPLASM The multitude of insect pests that often simultaneously attack soybean throughout the world has prompted efforts to develop germplasm with multiple insect resistance. Van Duyn et al.144 screened most of the world germplasm collection of soybean MGs VII and VIII for resistance to Mexican bean beetle and found three PIs (171451, 227687, and 229358) to be highly resistant. One or more of these PIs was included in most germplasm screenings, frequently as resistant standards, for the past two decades. The three genotypes exhibit resistance to virtually all the major soybean insect pests (Table 1). Because foliar-feeding lepidopterans are major pests in the southern U.S., they have been the subjects of large screening trials on soybean from the maturity groups adapted to the region. Gary et al.33 examined 126 soybean PIs from MGs VI, VII, VIII, and IX for resistance to velvetbean caterpillar, soybean looper, corn earworm, beet armyworm, and tobacco budworm, Heliothis virescens (Fabricius). Although genotypes with no higher levels of resistance than the resistant standards, PIs 171451,

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TABLE 1 Soybean Genotypes Exhibiting Resistance to Multiple Insect Pests Insect pests Lepidoptera Accession PI163453 PI171444 PI171451

Coleoptera

Helicoverpa Pseudoplusia Anticarsia Spodoptera zea includens gemmatalis exigua X

Heteroptera

Trichoplusia Epilachna Cerotoma Diabrotica Epicauta Bemisia Stink ni varivestis trifurcata balteata vittata tabaci Bugs

X

X

X

X

X

X X X

X

PI209837 PI227219 PI227687

X

X X X

PI229358

X

X

X

X

PI245331 PI336119 PI381660 PI407073 PI407132 PI407300 PI407301 PI417061 PI417136 PI417310 PI423911 PI464935-1

X X X

X X X

X X

X

X X X

X X X

X

X

X X

X

X

X

X

X

X

X

X

X

X

X

X

X

X X

X

X X X X X X X

X X

X

X X

X

X X

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Heliothis virescens

X

X X

X

X

X

X

Ref. 90 34,35,57,83 25,33,54,62, 80, 83-85,114, 132,144,145 33 33 25,47,54,62,80, 83-85,114,124, 132,144,145 25,47,54,62,80, 83-85,114,124, 132,145,146 90 90 33 90 90 90 90 69,70,83 33,70 69,72 33 90

227687, and 229358, were identified, several demonstrated resistance to more than one of the insects. PI209837 exhibited the greatest resistance to most species and was suggested for use in the development of insect-resistant soybean cultivars. McKenna et al.90 examined the entire G. soja germplasm collection (473 accessions in 1985) to find genetically controlled resistance to transfer to soybean cultivars. No genotypes appeared more resistant than PI 229358 in field tests. In laboratory bioassays, PI245331 and PI407132 appeared to be good sources of resistance to velvetbean caterpillar and corn earworm and were more resistant to velvetbean caterpillar than the resistant standards, PI229358 and PI171451. Beet armyworms were best controlled by PI407300, and PIs 163453, 245331, and 407073 also demonstrated resistance to the beet armyworm and corn earworm. Several soybean PIs have demonstrated resistance to coleopterous pests. Although the effort is dominated by studies on Mexican bean beetle,62,69-71,114,144,145 research on beetles, primarily those that are pestiferous in the southern U.S., has shown that some of the genotypes exhibiting resistance to lepidopterous defoliators also impact them. Shortly after PIs 171451, 227687, and 229358 were found to be highly resistant to Mexican bean beetle,144 Clark et al.25 found similar results with bean leaf beetle, Cerotoma trifurcata (Forster), and striped blister beetle, Epicauta vittata (Fabricius). Layton et al.83 confirmed the findings on bean leaf beetle, but reported that these three PIs did not exhibit resistance greater than the cultivar “Davis.” Jensen52 also found “Davis” to have the greatest degree of resistance to bean leaf beetles among 15 commercial cultivars. This was surprising because “Davis” is often included as a susceptible standard in plant resistance studies involving lepidopterous pests.121 “Davis” proved to be highly susceptible to banded cucumber beetle, Diabrotica balteata Le Conte, with PIs 171444, 171451, 227687, 229358, and 417061 demonstrating resistance.83 After screening all MG V through VIII germplasm introduced before 1974 and all commercial soybean cultivars adapted to Louisiana, Gilman et al.34,35 found PI171444 to be the most resistant to the stink bug complex comprised of southern green stink bug Nezara viridula (L.) (predominant species), green stink bug Acrosternum hilare (Say), brown stink bug Euschistus servus (Say), and dusky stink bug Euschistus tristigmus (Say). Jones and Sullivan54 reported that PIs 171451, 227687, and 229358 exhibited resistance to southern green stink bug in two of three forcedfeeding tests with PI229358 appearing to be the most resistant. Of these three, only PI227687 had adequate resistance to reduce seed damage in field studies, and several other PIs were identified with higher levels of southern green stink bug resistance, especially PI171444.34 Layton et al.83 also found that PI171444 expressed resistance to bean leaf beetle and banded cucumber beetle. Thus, PI171444 possesses multiple pest resistance but it does not surface among the resistant genotypes for lepidopterous defoliators (Table 1). Included among the insects that are affected by multiple insect-resistant genotypes are occasional pests in the U.S. and those that are major pests in other countries. Although PI227687 was the most resistant PI to cabbage looper, Trichoplusia ni (Hübner), PIs 171451 and 229358 also reduced feeding by the insect.85 When these genotypes were examined for resistance to four defoliators in Taiwan, PI227687 showed the greatest resistance to beet armyworm, PI171451 to the lepidopterans © 1999 by CRC Press LLC

Portesia taiwana (Shiraki) and Orgyia sp., and PI229358 to the coleopteran Anomala cupripes (Hope).132 In Brazil, PI274453 and PI274454 demonstrated resistance to a complex of stink bugs (N. viridula, Piezodorous guildini [West], and Euschistus heros [Fabricius]) and, along with PI227687, resistance to Hedilepta indicata (Fabricius), a minor lepidopteran pest.84 Research with sweetpotato whitefly revealed that PIs 171451 and 229358 were not as preferred as commercial cultivars or PI227687 for oviposition, but PI227687 was more resistant than the other two PIs to the chrysomelid pests Diabrotica speciosa (Germ.) and Colaspis sp. (near occidentalis [L.]).84 Undoubtedly, if the accessions in Table 1 had been screened against all the pests listed, many would have demonstrated resistance to a larger group of pest species. The apparent broad spectrum of resistance for PIs 171451, 227687, and 229358 reflects the resistance screening effort devoted to these genotypes. Indeed, because these accessions are resistant to many soybean pests, they have been used as parents in insect resistance breeding programs worldwide. Earlier, it was noted that a narrow genetic base exists for soybean cultivars grown in the U.S.26 Even less genetic diversity exists for insect-resistant germplasm and cultivars.90 For example, the 23 insect-resistant soybean breeding lines released by soybean breeders and entomologists were derived from two donor parents.75 Of the resistant lines, 20 had PI229358 as the resistant parent, while the remaining 3 had PI171451 as the resistance source. Most of the resistant germplasm that has been released exhibit resistance to multiple insect pests (Table 2). The plant introduction PI229358 is the resistant donor for 29 of the lines, with 3 lines (D89-9121, MBB80-133, and L86K-73) derived from PI171451. The cultivar “Tracy M” is the source of resistance for D88-5328 and D88-5272. These results are not surprising considering that in 1985 all 10 of the public soybean insect resistance breeding programs in the U.S. were using PI229358 as the source of resistance, 8 were using PI171451, and 3 were using PI227687.130 Lambert and Tyler82 indicated that PI229358 has been used most often as the donor parent because it has the most agronomically desirable characters. A majority of soybean germplasm (MGs IV through VIII) are suited for the growing regions in the U.S. south where insect problems are severe (Table 2). The germplasm developed in Ohio, designated as HC83, has excellent resistance to Mexican bean beetle and Japanese beetle, Popillia japonica (Newman);38 however, resistance to lepidopterous pests has not been determined. The MG III germplasm, MBB80-133, L86K-73, and L86K-96, were developed as a joint effort by scientists in Maryland, Illinois, and Indiana.27 Although Mexican bean beetle was the primary target of this project, the lines have been screened against corn earworm, soybean looper, velvetbean caterpillar, and bean leaf beetle. The remaining germplasm emerged from breeding programs in North Carolina, South Carolina, Georgia, and the USDA-ARS program at Stoneville, Mississippi (Table 2). Foliar-feeding lepidopterans were the focus of these programs, and the releases demonstrate resistance to soybean looper, corn earworm, and velvetbean caterpillar. The Georgia line GATIR81-296 has been the subject of research to understand the ovipositional preference of soybean looper;6 to investigate the feeding behavior, growth, development

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TABLE 2 Soybean Germplasm Released by Soybean Breeders and Entomologists in the U.S. that Exhibits Resistance to Multiple Insect Pests Insect pests Germplasm N90-2282 N80-50232 N80-53201 D86-3429 D75-10169a D89-9121 D88-5328 D88-5272 D90-9216 D90-9220 D74-7058 D74-9397 D74-10301a D74-10467a D74-10479 HC83-46-1 HC83-46-2 HC83-50-1 HC83-123-9b MBB80-133c L86K-73c L86K-96c GAT-81-117 GAT-81-121 GAT-81-124 GAT-81-237 GAT-81-281 GAT-81-296d GAT-81-306 GAT-81-327 GAT-81-332 GAT-81-426 GAT-81-461 a b c d e

Helicoverpa zea

Pseudoplusia includens

X X X

X X

X X X X X X X X X X X

X X X X X X X X X X X X

X

X

X X X X X X X

X X X X X X X X X X X X X X X

X X X X X X X

Anticasia gemmatalis

Popillia japonica X X X

X X X X X X X

X

X X X

X X X X X X X X X X X X X X

Also resistant to Heliothis virescens. Also resistant to Spodoptera exigua. Also resistant to Cerotoma trifurcata. Also resistant to three Spodoptera sp., see Reference 4. Only one reference is provided for each germplasm line.

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Epilachna varivestis

X

X

X X X X X

X X X X X

X X X

X

Ref.e 19 19 19 43 46 61 60 60 60 60 48 48 48 48 48 38 38 38 113 27 27 27 82 82 82 82 82 82 82 82 82 82 82

time, and mortality of soybean looper and velvetbean caterpillar;5 and to determine its resistance to three Spodoptera spp.4 Although other soybean germplasm are resistant to beet armyworm, none have been screened against fall armyworm S. frugiperda (J. E. Smith), yellowstriped armyworm S. ornithogalli Guenée, and southern armyworm S. eridania (Cramer), except GATIR81-296 which was found to be resistant. Beach and Todd5 reported that GATIR81-296 possessed the best combination of good agronomic qualities and resistance to insect defoliators of any line tested in Georgia. This breeding line appeared to have resistance equal to PI229358, the resistant parent, a rare occurrence for insect-resistant germplasm that has been released and registered in the U.S. Virtually none of the resistant germplasm is equal in resistance levels to their donor parents.82 All the released breeding lines have greatly improved agronomic traits relative to their resistant parents but none yield as well as presently grown commercial cultivars. Thus, Lambert and Tyler82 believe all released lines, including the insect-resistant cultivars that have been developed, are useful only as advanced breeding lines. The main effort of the breeding program for resistance in Brazil, especially at the Instituto Agronomica de Campinas (IAC) at San Paulo, is directed against the stink bug complex of N. viridula, P. guildinii, and E. heros. The PIs 227687 and 274454 have been used as the sources of resistance for stink bugs, and although leaf feeders are of secondary interest, selection for resistance to them has occurred. Five PIs (171444, 171451, 227687, 229358, and 417061) and three germplasm lines (ED73-371, IAC78-2318, and IAC100) carry some degree of resistance to stink bugs and a leaf feeder.111 The IAC germplasm has demonstrated multiple insect resistance. IAC78-2318 has shown resistance to sweetpotato whitefly, a tortricid stem borer, Epinotia aporema (Wals.), and stink bugs.84 “Centenaria IAC100” was released in 1988 and it appears to have a high degree of tolerance to stink bug attack and some resistance to leaf feeders (species not specified)111 and corn earworm.72 D72-9601-1 selected for resistance to soybean looper in the U.S. was reported to exhibit resistance to velvetbean caterpillar.84 In 1989, Lourencao et al.84 reported that 18 improved cultivars had been released by the IAC. Although it was not specified as to whether insect-resistance was incorporated into each of the cultivars, it appears that multiple insect-resistant cultivars are a goal of the breeding program. Certainly, more effort against stink bug pests has been made in South America than in the U.S. Another area that has received scant attention in the U.S. is pod feeding resistance to species (i.e., Heliothine larvae) that feed on foliage and pods.40,47 This has not been the case in other parts of the world where pod-feeders are major pests. Pods of PI227687 were resistant to larvae of the limabean pod borer,133 a major pest of soybean in Asia and Oceania, especially Indonesia.131 Resistance screening for the soybean pod borer also receives high priority in China.36 The pest status of pod feeding insects in other areas of the world relative to their status in the U.S. may explain the differential effort. However, the corn earworm was ranked first among the eight major soybean pests in the U.S. in terms of economic impact,63 and it is more severe as a pod feeder than a defoliator. The availability of alternative strategies to manage pod feeders in the U.S. relative to other areas may explain why studies on pod feeding pests, other than stink bugs, number far less than those on foliage feeders. © 1999 by CRC Press LLC

Germplasm is available with multiple insect resistance, but the resistance is not equally effective against all insect pests (Tables 1 and 2).1,47,48,80,83,122-124 An evaluation of five breeding lines with PI229358 parentage for resistance to five leaf feeding species indicated that identification of breeding lines with resistance equal to this PI should be based on evaluations against each species.48 Using PI229358 as the resistant parent, Smith and Brim122 demonstrated resistance in the Mexican bean beetle-resistant F3 lines which did not correlate with leaf-feeding resistance to corn earworm. With PI227687 as the donor parent, Smith and Brim123 found low incidence of progeny with corn earworm resistance among backcross populations with high levels of Mexican bean beetle resistance. Their findings suggest that indirect selection for resistance to corn earworm via Mexican bean beetle resistance is ineffective, and that future breeding efforts should focus on direct selection for resistance to corn earworm. Lambert and Kilen79 also tested five leaf-feeding insects48 against F1 intercrosses of the PIs 171451, 227687, 229358 and found different levels of resistance among the parental material and the progeny. Subsequently, Lambert and Kilen80 suggested that the development of cultivars from PI229358 with resistance to several insect species may be possible because of the time consuming process of screening each insect species at each selection stage in a breeding program. However, they suggested that at the final stage of selection, germplasm should be screened for all potential pests in the area of adaption of the potential cultivar, because of the genetic diversity of the insects and the germplasm. Thus, the causal factors for the different degrees of resistance to multiple insects appear multi-fold. The genetic basis of resistance and the mechanisms of resistance are important components of this phenomenon.

C. MECHANISMS

OF

RESISTANCE

Because the literature is voluminous in the mechanisms of resistance99 in soybean it is not the goal of this chapter to present a comprehensive review of research in this area. Categories of insect resistance in soybean germplasm are summarized in Table 3. The antixenosis and antibiosis resistance in three widely studied PIs (171451, 227687, and 229358) appear to be chemically based. Reviews are available that chronicle progress on elucidating the chemical basis of resistance.66,121,147 Resistance in soybean is attributable to both constitutive and induced factors.65 Allelochemicals associated with feeding deterrency and antibiosis in soybean are isoflavones (i.e., plaseol, afrormosin, coumestrol, diadzein, and glyceollin),20,31,41,67,94 phenolic acids,23 and phytoalexins.41 Primary metabolites also have been implicated in resistance mechanisms.65 Tester135 reported that PI227687 and PI229358 did not accumulate as much total nitrogen or accumulate nitrogen at the rate of susceptible cultivars. Foliage of resistant soybean germplasm was found to contain lesser amounts of reducing sugars and protein.143 In addition, Elden and Kenworthy28 reported that foliar nitrogen levels were similar among Mexican bean beetle-resistant and susceptible genotypes. Although three elements (P, Ca, Fe) were lower in the resistant genotypes, no direct cause–effect relationship with resistance was observed. Wier and Boethel148-149 demonstrated reduced herbivore performance, symptomatic of antibiosis, when soybean loopers were fed foliage of non-nodulating soybean

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TABLE 3 Categories of Insect Resistance in Soybean Germplasm Resistance category

Insect Helicoverpa zea, Heliothis virescens

Antibiosis Antibiosis, antixenosis Antibiosis Undescribed

Anticarsia gemmatalis, Pseudoplusia includens, Spodoptera exigua Pseudoplusia includens

Antibiosis Antibiosis, antixenosis Tolerance Antibiosis, antixenosis Tolerance Antibiosis

Bemisia argentifolii Nezara viridula

Epilachnia varivestis

Antibiosis, antixenosis Tolerance

Germplasm

Ref.

PIs 227687,229358, ED73-375 (PI229358 x “Bragg”) PI171451

47,75

PIs 227687, 229358 PI171451

5,79,132 79

GATIR 81-296 PIs 227687, 229358

6 5,6,8,107, 121,124 74 54,57

F90-724 PIs 171444, 171451, 227687, 229358 ED73-355, “IAC-100” HC83-46-1, HC83-46-2, HC83-50-1, HC83-123-9, L76-0038, L76-0049, L76-0132, L76-0272, L76-0328, L78-608, PI227687 PIs 171451, 227687, 229358, “Shore” PI90481, “Shore”

9,25,75

54,65 38,114

30,38,62,125, 144,145 117

isolines that contained low foliar nitrogen. Thus, the mechanisms of resistance may involve secondary plant compounds or nutrient levels or both.28

D. INHERITANCE

OF

RESISTANCE

In contrast to the number of studies on mechanisms of resistance in soybean, there have been few investigations on the inheritance of resistance. Kogan62 was among the first to provide evidence that resistance to Mexican bean beetle was an inherited trait, suggesting a semidominance type of inheritance. Sisson et al.120 agreed with Kogan62 after evaluating the resistance of F3 progeny of crosses between commercial cultivars and resistant PIs 227687, 229358, and 229321. No simple Mendelian genetic ratios could be detected from frequency distributions of F3 progeny means. The results suggested a quantitative inheritance of resistance and further suggested involvement of two or three major genes controlling resistance. Luedders and Dickerson85 crossed early (MG II) cultivars with late maturity resistant PI171451 and PI229358 (MGVII) and screened the F3 and F4 progeny and parents against the cabbage looper. It was concluded that linkage between resistance and maturity genes would not be a factor in transferring the resistance from latematurity PIs to genotypes adapted in more northern latitudes.

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Kilen et al.58 crossed PI229358 and “Davis” and examined the F1, F2, and F3 progenies for resistance to soybean looper. Data suggested no dominance for resistance with the F2 distribution skewed toward the susceptible, indicative of partial dominance toward susceptibility. About 20% of the F2 plants and 27% of the F3 plants exhibited resistance equal to PI229358, which suggested only a few major genes condition resistance to the soybean looper. Kenty et al.56 also found that a few genes contributed to soybean looper resistance and that data supported quantitative inheritance of resistance. Lambert and Kilen79 compared “Davis” with PIs 171451, 227687, and 229358 and their intercrosses and examined the resistance of F1 lines and donor parents to five lepidopterous leaf-feeders. The response to F1 progeny indicated resistance could be increased through intercrossing and selection, and because the larval size on the F1 hybrids tended to be similar to the most resistant parent, the presence of slight dominance for resistance was likely. Because differences in degree of resistance to certain insect species occur among the resistant PIs 171451, 227687, and 229358, Kilen and Lambert59 conducted a study with velvetbean caterpillar to determine if these resistant genotypes have different genes controlling resistance. Similar to the results in an earlier study,58 the large number of F3 progeny exhibiting damage greater than the parents suggested only a few major genes for resistance and partially dominant inheritance of susceptibility. The highly consistent pattern of very susceptible F3 lines recovered from crosses of resistant parents also suggested that each genotype carries at least one gene for resistance that differs from the other two sources. Rufener et al.116 used an antibiosis screening technique115 to determine the inheritance of Mexican bean beetle resistance in soybean populations resulting from crosses of the susceptible cultivar “Williams” and three resistant breeding lines. The intermediate resistance of F1 plants confirmed the findings of others, suggesting additive gene action or partial dominance.58,62 There appeared to be different levels of dominance depending on the resistant parent (PI171451 or PI229358) used, which was not surprising based on findings that resistance genes in the PIs were not identical.59 Although the larval bioassay used is more precise than field screening, it is not precise enough to yield clear Mendalian ratios. Therefore, Rufener et al.116 recommended that progress in breeding for Mexican bean beetle resistance would be best obtained by treating resistance as a quantitative trait. If genetic drift is minimized, recurrent selection should be effective; however, backcrossing is unlikely to transfer the full complement of resistance genes.

E. GENOTYPE

BY

ENVIRONMENT INTERACTIONS

Various factors affect the expression of insect resistance in soybean.37 Genotype by environment interactions, especially when genotypes are evaluated for resistance in the field with naturally occurring insect populations, are a problem encountered in the development of insect-resistant cultivars.113 Factors responsible for these interactions include plant phenological stage, planting dates, leaf age and position on plant, timing of insect attack, insect species or species complex, insect population, plant nutrition, and moisture stress.

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Reynolds and Smith106 reported that the top two fully expanded leaves near the apex of PI227687 plants supported higher soybean looper growth rates than the third or lower leaves, and growth rates were not different on plants varying in age from 4 to 8 weeks old. McWilliams and Beland91 also demonstrated corn earworm preferred younger leaves on vegetative stage soybean. Mechanical wounding of PI227687 leaves decreased larval growth rates relative to those on normal PI227687 leaves supporting the existence of induced resistance, a phenomenon demonstrated in subsequent studies.23 Nault et al.92 found that corn earworm preferred older leaves of both resistant (GATIR81-296) and susceptible (“Cobb”) plants throughout the season. Nault et al.93 also reported that resistance to soybean looper and velvetbean caterpillar declined with plant maturity, and Hammond et al.37 confirmed these findings with Mexican bean beetle. Rowan et al.113 attempted to elucidate the soybean maturity effect on the expression of resistance to lepidopterous pests by examining resistant and susceptible germplasm in MGs V through VIII. When insect defoliation occurred in the vegetative or early reproductive stage of development, plant maturity was not associated with defoliation level. But, when defoliation occurred late in the season, the early maturing genotypes had greater defoliation than the late maturing genotypes, resulting in greater defoliation on resistant MG V genotypes than susceptible MG VII and VIII genotypes. Collectively, these studies revealed that the level of resistance in a genotype will vary during plant ontogeny, and genotypes at the same stage of development should be compared with a reliable assessment of resistance. Boerma et al.13 recommended stratifying genotypes by maturity in field experiments or including resistant and susceptible standards of various maturities represented by the test genotypes. Studies have shown that moisture stress can influence the expression of resistance. In one study,77 soybean grown under water deficit conditions had adverse effects on soybean looper larvae. In a subsequent study, defoliation by soybean looper of irrigated plants of both resistant (D75-10169) and susceptible (“Centennial”) genotypes was more than 50% greater than defoliation on nonirrigated plants and larvae developing on nonirrigated plants of both genotypes were smaller than those on irrigated plants.78 Hammond et al.37 observed loss of resistance to Mexican bean beetle in several resistant genotypes when grown under excessive moisture conditions.

F. COMPATIBILITY

WITH

OTHER MANAGEMENT TACTICS

Concurrent with efforts to develop insect-resistant cultivars have been studies to determine the impact of these resistant soybean genotypes on natural enemies of soybean pests. The effect of antibiotic resistance factors in soybean on parasites and microbial pathogens has perhaps been studied in greater depth than for many other crops.49 Boethel and Orr15 reviewed much of this research which documented the influence of soybean antibiosis through the third and fourth trophic levels. Studies examining egg parasites (Scelionidae), egg-larval parasitoids (Encyrtidae), larval parasitoids (Braconidae, Eulophidae, Tachinidae), and predators (Pentatomidae, Lygaeidae) revealed adverse effects on natural enemy biology such as reduced growth

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rates, survival, fecundity, and longevity. Rogers and Sullivan108,109 demonstrated that the predator, Geocoris punctipes (Say), exhibited increased development time and mortality and decreased weight gain as a result of directly feeding on foliage of PI171451 and PI229358. The same responses were seen when the predator fed on insect hosts that had fed on the resistant plants. The apparent incompatibility between pest-resistant soybean and biological control was noted by Hare39 in a review of host plant resistance-biological control interactions in applied systems. Antagonistic interactions were found in 6 of 16 cases involving parasitoids with soybean being responsible for four of these clearly antagonistic interactions. Hare39 suggested that the disruptive tritrophic interactions may be more common in newly introduced plant species or new, resistant crop varieties (e.g., soybean) than in associations with longer evolutionary histories. Although much of the research indicated that some degree of incompatibility existed between these management tactics, some studies have not shown this. Wheatley and Boethel146 examined a multi-trophic system comprised of resistant soybean, a mite herbivore, and a predaceous mite that allowed for assessment of the interactions at the population level. There was no indication that any antibiosis effect persisted through the third trophic level, and the additive effect of plant resistance and predation supported compatibility between the tactics. McCutcheon and Turnipseed89 found that resistant soybean genotypes did not influence the incidence of parasitism of lepidopterous pests in South Carolina. Despite corn earworm population density being 3.6-fold greater on densely pubescent than glabrous “Davis” isolines, there was no difference in the percent parasitism by the braconid Microplitis croceipes (Cresson).136 These results indicated that glabrous soybean and biological control with M. croceipes would be compatible in the management of corn earworm. Powell and Lambert103 found that egg predation of corn earworm by G. punctipes was not adversely affected by resistant soybean genotypes (PI229358, “Lamar”) or by isolines with reduced pubescence, a trait also less preferred by the pest. McAuslane et al.88 examined the influence of foliar pubescence on the abundance and parasitism of silverleaf whitefly. Greater numbers of immature whiteflies occurred in pubescent (D75-10169) and hirsute (D90-9220) than glabrous (D90-9216) near isolines of soybean. However, four species of aphelinid parasitoids differed in their distribution relative to pubescence. Encarsia nigricephala Dozier and E. transvena Timberlake more commonly parasitized whitefly on glabrous lines, while E. pergandiella Howard and Eretmocerus nr. californicus more often parasitized whiteflies on the hairy genotypes. The early studies that showed adverse effects on natural enemies by insectresistant soybean genotypes used the PIs that expressed high levels of antibiosis and most examined the effects on individual organisms without tracking the influence of the resistance factor on subsequent generations of the natural enemies. The case histories expounded in this chapter illustrate that studies in the field89 and at the population level,146 and with germplasm exhibiting another mechanism of resistance (antixenosis-pubescence),88,136 support the compatibility of plant resistance and biological control. The findings that whitefly parasitoid species differed in their attack based on pubescence of the plants is particularly interesting. It illustrates the uniqueness of each system with regard to the effect of plant resistance on the biologies of © 1999 by CRC Press LLC

natural enemies and makes generalizations about compatibility of the tactics difficult.16,39,98,99 Little attention has been given to interactions between insect-resistant soybean and entomopathogens, but the data available suggest compatibility. Although the fungal pathogen Nomuraea rileyi (Farlow) Sampson caused 100% mortality of corn earworm on both resistant and susceptible soybean genotypes, the larvae died at an earlier age on the resistant plants.10 Beach and Todd7 reported that Nuclear Polyhedrosis Virus (NPV) for soybean looper and velvetbean caterpillar were equally effective in reducing larvae and defoliation by these species, and NPV-infected larvae of both species consumed significantly less foliage when reared on the resistant genotype. Insect-resistant soybean germplasm can influence the toxicity of insecticides toward soybean insect pests, but the outcome of the interactions are variable depending on the insecticide and insect species involved. In laboratory studies, corn earworm larvae fed foliage of ED73-371, a genotype resistant to Mexican bean beetle, were more susceptible to methomyl and Bacillus thuringiensis Berliner (Bt) but not to methyl parathion, whereas the opposite situation occurred when soybean looper was the target species.55 The data on soybean looper and methyl parathion are interesting because the insect has been known to be resistant to methyl parathion since the 1970s.15 Whether the population examined in 1978 was actually resistant is not known but the prospects of enhancing activity against an insect resistant to an insecticide by use of a resistant plant type is exciting. In field trials, control of corn earworm with methyl parathion and Bt was greater in ED73-371 soybean plots than in plots of the susceptible germplasm.55 In yet another study, the susceptibility of velvetbean caterpillar to fenvalerate and acephate and soybean looper to acephate was enhanced by feeding on resistant PI227687.110 Incorporation of coumestrol, an isoflavonoid associated with PI227687 resistance, into artificial diet resulted in reductions in weight gain in soybean looper larvae and enhanced the toxicity of fenvalerate while reducing the toxicity of methomyl.

G. RELEASED INSECT-RESISTANT CULTIVARS Four insect-resistant cultivars, “Shore,” “Crockett,” “Lamar,” and “Lyon,” have been registered and released by breeding programs in the 26 years since research began to explore insect resistance in soybean germplasm. Their adoption has been limited for various reasons, and the hectarage on which they are grown is negligible. “Shore” soybean originated as a cross of PI80837 X “Hood”95,128 and has resistance to Mexican bean beetle.29,30 Under heavy Mexican bean beetle pressure in the U.S. Atlantic Coastal Plain region, “Shore” yielded approximately 17% more than a susceptible cultivar “York.”22,126,128 However, when Mexican bean beetles were not a problem, the yield of “Shore” did not compare favorably with other commercial cultivars.65 For this reason and the rather narrow geographic range over which it is adapted, “Shore” has not been widely accepted by growers. Three insect-resistant cultivars have been developed for southern U.S. “Crockett” (PI171451 X “Hampton”) is a MG VIII cultivar with resistance to foliar-feeding insects and moderate resistance to southern green stink bug.17 “Lamar” and “Lyon” are MG VI cultivars that are increases of F5 lines of crosses of “Tracy M” X F3 selection

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(“Centennial” X D75-10169) and D82-2218 (“Bedford” X “Tracy M”) X “Lamar,” respectively.44,45 “Lamar” has resistance to foliar feeding by soybean looper, velvetbean caterpiller, and corn earworm, with evidence of a resistance factor in the pod walls to the later species. “Lyon” has resistance to soybean looper that is equal to that of “Lamar.” Because of their late maturity, low yield, and tendency to lodge (“Crockett”), these southern adapted cultivars have not been grown on large hectarages.82 Some commercial soybean cultivars have moderate levels of resistance to soybean insects. After evaluating 46 cultivars and 10 insect-resistant checks for resistance to foliar-feeding lepidopterans, Rowan et al.112 found that some cultivars showed 50% of the resistance of the insect-resistant entries. No differences were found in levels of defoliation in MG V entries but “Coker 686” and “Deltapine 566” (VI), “Braxton” (VII), and “Coker 6738” were the most resistant genotypes in the late maturity groups.

H. BIOTECHNOLOGICAL ADVANCES A molecular map of soybean has been developed,118 and molecular markers associated with insect resistance genes have been identified.105 Restriction fragment length polymorphism (RFLP) markers have been used in plants from crosses of “Cobb” (susceptible) and PI229358. Several loci were found to be associated with reduced corn earworm defoliation with major loci grouped on Linkage Group M and H of the USDA/ISU public soybean genetic map14,105 Molecular markers can be used to identify and map major insect resistance loci to facilitate their introgression into elite soybean germplasm.12 The potential applications for marker-assisted breeding are several. Perhaps the most obvious is identification of insect-resistant lines by selecting for markers instead of extensive screenings for resistance. With the molecular map of soybean completed, molecular markers for insect resistance being identified, and the potential for avoiding the genotype by environment interactions that plague traditional screening, soybean breeders have the tools to expedite development of resistant cultivars. Soybean genetic engineering is not routine14 and only limited success has occurred in the development of plants engineered for pest resistance.13 Parrott et al.101 successfully incorporated the native Bt cryIA(b) insecticidal protein gene into a breeding line F376, selected for its rapid growth in embryonic suspensions. The engineered plants inhibited feeding, growth, and development of velvetbean caterpillar compared with nonengineered plants. The level of feeding deterrence of the genetically engineered plants was comparable to that exhibited by the resistant breeding line GATIR81-296.14 More recently, the synthetic Bt cryIAc gene was incorporated into the cultivar “Jack,” also selected because of its high embryonic capacity.129 The transgenic plants were protected from damage by the soybean looper, velvetbean caterpillar, and corn earworm. Although Stewart et al.129 acknowledged that their report was the first of soybean transformed with a biologically effective insect resistance gene, they did not feel the transgenic lines were deployable in the southeast where lepidopteran damage is most likely to occur. Survival levels are still high enough on these current transgenic lines to foster development of resistant insect populations.

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III.

CONCLUSIONS

Soybean breeders and entomologists have encountered several obstacles in their attempts to develop insect-resistant cultivars for soybean insect pest management systems. Although pubescent germplasm has been successfully used in management of potato leafhopper, the four cultivars released with resistance to foliar-feeding insects have not been widely used by growers primarily because of yield limitations. The expectation that insect-resistant cultivars yield equal to or greater than existing commercial cultivars remains today, as it did when expressed by Newsom96 years ago. Sporadic outbreaks of insect pests, efficient and generally inexpensive insecticides, and an ecologically based IPM system that has been highly effective and costconscious for nearly a quarter century demand that an insect-resistant cultivar possess high yield potential for adoption by producers. These factors have helped relegate insect resistance to rather low priority in soybean breeding programs. Depending on the environment for which a cultivar is being developed, resistance may have to be maintained for as many as 12 diseases (including 44 biotypes), 5 nematode species (6 biotypes), and tolerance to herbicides.82 Soybean breeders often have responded to these immediate concerns. There has been great difficulty recovering high yield in insect-resistant lines and several breeding cycles are necessary to recover lines that have seed yield approaching those of current cultivars.82 This could be caused by linkage of resistance genes to undesirable genes which impact on yield, or because seed yield is controlled by many genes and insect resistance is controlled by much fewer genes, and the proportion of F2 plants from any crosses that will have high yield and insect resistance is small. Thus, the small size of breeding populations may be a major limitation to identification of germplasm with insect resistance and high yield.82 Treating insect resistance as a quantitative trait and using recurrent selection to accumulate alleles has been suggested as a more effective alternative to the traditional backcross method.116 The incorporation of the Bt gene into soybean offers exciting prospects of insertion of other foreign genes for insect resistance. The concern about development of insect resistance to the Bt transgenic plants expressed by Stewart et al.129 is warranted (see Boethel et al.15 and Mascarenhas et al.86). Again, the infrequent occurrence of damaging populations, the presence of pest complexes, adequate management for most current pests, and the possibility of a cost-prohibitative technology fee make the adoption of Bt transgenic plants for management of soybean insect pests less essential than it might be for insect management on other crops. However, this new technology should not be discounted. Traditional soybean breeding programs have identified numerous genotypes and germplasm lines with resistance to multiple insect species, especially to foliarfeeding pests. Three PIs, 171451, 227687, and 229358, have been extensively examined and instrumental in the many contributions made in the arena of soybean plant resistance. However, reliance on these PIs as the resistance donor parents appears to have limited the genetic diversity in the multiple insect resistant germplasm. This chapter will close with optimism that agronomically acceptable insectresistant cultivars will be forthcoming. The investment made in traditional breeding

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programs in concert with advances in biotechnology fuel this optimism. There is a place for insect-resistant cultivars in soybean culture which will complement the current IPM programs to continue the sustained, profitable production of one of the world’s major crops.

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89. McCutcheon, G. S. and S. G. Turnipseed, Parasites of lepidopterous larvae in insect resistant and susceptible soybeans in South Carolina, Environ. Entomol., 10:69-74, 1981. 90. McKenna, T., L. Lambert, J. D. Ouzts, and T. C. Kilen, Evaluation of wild soybean, Glycine soja, for resistance to foliar feeding insects, J. Miss. Acad. Sci., 33:17-24, 1988. 91. McWilliams, J. M. and G. L. Beland, Bollworm: effect of soybean leaf age and pod maturity on development in the laboratory, Ann. Entomol. Soc. Am., 70:214-216, 1977. 92. Nault, B. A., J. N. All, and H. R. Boerma, Influence of soybean planting date and leaf age on resistance to corn earworm (Lepidoptera: Noctuidae), Environ. Entomol., 21:264-268, 1992. 93. Nault, B. A., J. N. All, and H. R. Boerma, Resistance in vegetative and reproductive stages of a soybean breeding line to three defoliating (Lepidoptera: Noctuidae) pests, J. Econ. Entomol., 85:1507-1515, 1992. 94. Neupane, F. P. and D. M. Norris, Iodoacetic acid alteration of soybean resistance to the cabbage looper (Lepidoptera: Noctuidae), Environ. Entomol., 19:215-221, 1990. 95. Newcomer, J. L. and P. E. Paz, Shore: a new soybean variety resistant to Mexican bean beetle, Univ. of Maryland Agronomy Mimeo. No. 45, College Park, 1975. 96. Newsom, L. D., Progress in integrated pest management of soybean pests, In: E. H. Smith and D. Pimental, Eds., Pest Control Strategies, Acad. Press, New York, 1978, 157. 97. Newsom, L. D., M. Kogan, F. D. Miner, R. L. Rabb, S. G. Turnipseed, and W. H. Whitcomb, General accomplishments toward better pest control in soybean, In: C. B. Huffaker, Ed., New Technology of Pest Control, John Wiley, New York, 1980, 51. 98. Orr, D. B. and D. J. Boethel, Influence of plant antibiosis through four trophic levels, Oecologia, 70:242-249, 1986. 99. Painter, R. H., Insect Resistance in Crop Plants, Univ. Press of Kansas, Lawrence, 1951. 100. Palmer, R. G., T. Hymowitz, and R. L. Nelson, Germplasm diversity within soybean, In: D. P. S. Verma and R. C. Shoemaker, Eds., Soybean: Genetics, Molecular Biology, and Biotechnology, CAB International, Wallingford, 1996, 1. 101. Parrot, W. A., J. N. All, M. J. Adang, M. A. Bailey, H. R. Boerma, and C. N. Stewart, Jr., Recovery and evaluation of soybean plants transgenic for a Bacillus thuringiensis var. kurstaki insecticidal gene, In Vitro Cell. Dev. Biol., 30P:144-149, 1994. 102. Poos, F. W. and F. F. Smith, A comparison of feeding habits of some species of Empoasca, J. Econ. Entomol., 24:361-371, 1931. 103. Powell, J. E. and L. Lambert, Soybean genotype effects on bigeyed bug feedings on corn earworm in the laboratory, Crop Sci., 33:556-559, 1993. 104. Probst, A. H. and R. W. Judd, Origin, U.S. history, and development, and world distribution, In: B. E. Caldwell, Ed., Soybeans: Improvement, Production, and Uses, ASA, CSSA, & SSSA, Madison, 1976, 1. 105. Rector, B. G., J. N. All, and H. R. Boerma, Identification of molecular markers associated with insect resistance QTLS in soybean, In:, 6th Biennial Conf. — Molecular and Cellular Biology of the Soybean, Univ. of Missouri, Columbia, (Abstract), 1996. 106. Reynolds, G. W. and C. M. Smith, Effects of leaf position, leaf wounding, and plant age of two soybean genotypes on soybean looper (Lepidoptera: Noctuidae) growth, Environ. Entomol. 14:475-478, 1985.

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107. Reynolds, G. W., C. M. Smith, and K. M. Kester, Reductions in consumption, utilization, and growth rate of soybean looper (Lepidoptera: Noctuidae) larvae fed foliage of soybean genotype PI 227687, J. Econ. Entomol., 77:1371-1375, 1984. 108. Rogers, D. J. and M. J. Sullivan, Nymphal performance of Geocoris punctipes (Hemiptera: Lygaeidae) on pest-resistant soybeans, Environ. Entomol., 15:1032-1036, 1986. 109. Rogers, D. J. and M. J. Sullivan, Growth of Geocoris punctipes (Hemiptera: Lygaeidae) in attached and detached leaves of pest-resistant soybeans, J. Entomol. Sci., 22:282-285, 1987. 110. Rose, R. L., T. C. Sparks, and C. M. Smith, Insecticide toxicity to the soybean looper and the velvetbean caterpillar (Lepidoptera: Noctuidae) as influenced by feeding a resistant soybean (PI 227687) leaves and coumestrol, J. Econ. Entomol., 81:1288-1294, 1988. 111. Rossetto, C. J., Breeding for resistance to stink bugs, In: A. J. Pascale, Ed., Proc. World Soybean Res. Conf. IV, AASOJA, Buenos Aires, 1989, 2046. 112. Rowan, G. B., H. R. Boerma, J. N. All, and J. Todd, Soybean cultivar resistance to defoliating insects, Crop Sci., 31:678-682, 1991. 113. Rowan, G. B., H. R. Boerma, J. N. All, and J. W. Todd, Soybean maturity effect on expression of resistance to lepidopterous insects, Crop Sci., 33:433-436, 1993. 114. Rufener II, G. K., R. B. Hammond, R. L. Cooper, and S. K. St. Martin, Mexican bean beetle (Coleoptera: Coccinellidae) development on resistant and susceptible soybean lines in the laboratory and relationship to field selection, J. Econ. Entomol., 79:1354-1358, 1986. 115. Rufener II, G. K., R. B. Hammond, R. L. Cooper, and S. K. St. Martin, Larval antibiosis screening techniques for Mexican bean beetle in soybean, Crop. Sci., 27:598-600, 1987. 116. Rufener II, G. K., S. K. St. Martin, R. L. Cooper, and R. B. Hammond, Genetics of antibiosis resistance to Mexican bean beetle in soybean, Crop Sci., 29:618-622, 1989. 117. Schillinger, J. A., Host plant resistance to insects in soybean, In: L. D. Hill, Ed., Proc. World Soybean Res. Conf. I., Interstate Printers & Publishers, Danville, 1976, 579. 118. Shoemaker, R. C. and J. E. Specht, Integration of soybean molecular and classical genetic linkage groups, Crop Sci., 35:436-446, 1995. 119. Singh, B. B., H. H. Hadley, and R. L. Bernard, Morphology of pubescence in soybeans and its relationship to plant vigor, Crop Sci., 11:13-16, 1971. 120. Sisson, V. A., P. A. Miller, W. V. Campbell, and J. W. Van Duyn, Evidence of inheritance of resistance to the Mexican bean beetle in soybeans, Crop Sci., 16:835-837, 1976. 121. Smith, C. M., Expression, mechanisms, and chemistry of resistance in soybean, Glycine max L. (Merr.) to the soybean looper, Pseudoplusia includens (Walker), Insect Sci. Applic., 6:243-248, 1985. 122. Smith, C. M. and C. A. Brim, Field and laboratory evaluations of soybean lines for resistance to corn earworm leaf feeding, J. Econ. Entomol. 72:78-80, 1979. 123. Smith, C. M. and C. A. Brim, Resistance to the Mexican bean beetle and corn earworm in soybean genotypes derived from PI2276787, Crop Sci., 19:313-314, 1979. 124. Smith, C. M. and D. F. Gilman, Comparative resistance to multiple insect-resistant soybean genotypes to the soybean looper, J. Econ. Entomol., 74:400-403, 1981. 125. Smith, C. M., R. F. Wilson, and C. A. Brim, Feeding behavior of Mexican bean beetle on leaf extracts of resistant and susceptible soybean genotypes, J. Econ. Entomol., 72:374-377, 1979.

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126. Smith, J. C. and M. W. Alexander, An evaluation of Mexican bean beetle resistance in 'Shore' variety soybeans in Tidewater Virginia, VA. J. Sci., 26:45, 1975. 127. Smith, K., Importance of soybeans, In: L. G. Higley and D. J. Boethel, Eds., Handbook of Soybean Insect Pests, Entomol. Soc. Am., Lanham, 1994, 3. 128. Smith, T. J., H. M. Camper, Jr., and J. A. Schillinger, Registration of Shore soybean., Crop Sci., 15:100, 1975. 129. Stewart, C. N., M. J. Adang, J. N. All, H. R. Boerma, G. Cardineau, D. Tucker, and W. A. Parrott, Genetic transformation, recovery, and characterization of fertile soybean transgenic for a synthetic Bacillus thuringiensis cryIAc gene, Plant Physiol., 112:121-129, 1996. 130. Sullivan, M. J., Resistance to insect defoliators, In: R. Shibles, Ed., Proc. World Soybean Res. Conf. III, Westview, Boulder, 1985, 400. 131. Talekar, N. S. and B. S. Chen, Identification of sources of resistance to Limabean podborer (Lepidoptera: Pyralidae) in soybean, J. Econ. Entomol., 76:38-39, 1983. 132. Talekar, N. S., H. R. Lee, and Suharsano, Resistance of soybean to four defoliator species in Taiwan, J. Econ. Entomol., 81:1469-1473, 1988. 133. Talekar, N. S. and C. P. Lin, Characterization of resistance to Limabean pod borer (Lepidoptera: Pyralidae) in soybean, J. Econ. Entomol., 87:821-825, 1994. 134. Teare, I. D. and H. F. Hodges, Soybean ecology and physiology, In: L. G. Higley and D. J. Boethel, Eds., Handbook of Soybean Insect Pests, Entomol. Soc. Am., Lanham, 1994, 4. 135. Tester, C. F., Constituents of soybean cultivars differing in insect resistance, Phytochemistry, 16:1899-1901, 1977. 136. Tillman, P. E. and L. Lambert, Influence of soybean pubescence on incidence of the corn earworm and the parasitoid, Microplitis croceipes, Southwest. Entomol., 20:181-185, 1995. 137. Todd, J. W., R. M. McPherson, and D. J. Boethel, Management tactics for soybean insects, In: L. G. Higley and D. J. Boethel, Eds., Handbook of Soybean Insect Pests, Entomol. Soc. Am., Lanham, 1994, 115. 138. Turnipseed, S. G., Management of insect pests of soybeans, In: E. V. Komarek, Ed., Proc. Tall Timbers Conf. on Ecol. Animal Control by Habitat Mgmt., Tall Timbers Res. Stn., Tallahassee, 1972, 189. 139. Turnipseed, S. G., Influence of trichome density on populations of small phytophagous insects on soybean, Environ. Entomol., 6:815-817, 1977. 140. Turnipseed, S. G. and M. Kogan, Soybean entomology, Ann. Rev. Entomol., 21:247-282, 1976. 141. Turnipseed, S. G. and M. Kogan, Integrated control of insects, In: Soybeans: Improvement, Productions, and Uses, J. R. Wilcox, Ed., ASA, CSSA, & SSSA, Madison, 1987, 779. 142. Turnipseed, S. G. and M. J. Sullivan, Plant resistance in soybean insect management, In: L. D. Hill, Ed., Proc. World Soybean Res. Conf. I, Interstate Printers & Publishers, Danville, 1976, 549. 143. Van Duyn, J. W., Investigations Concerning Host Plant Resistance to the Mexican Bean Beetle, Epilachnia varivestis Mulsant, in Soybeans, Glycine max (L.) Merrill, Ph.D. Dissertation, Clemson Univ., Clemson, 1971. 144. Van Duyn, J. W., S. G. Turnipseed, and J. D. Maxwell, Resistance in soybeans to the Mexican bean beetle. I. Sources of resistance, Crop Sci., 11:572-573, 1971. 145. Van Duyn, J. W., S. G. Turnipseed, and J. D. Maxwell, Resistance in soybeans to the Mexican bean beetle: II. Reactions of the beetle to resistant plants, Crop Sci., 12:561-562, 1972.

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146. Wheatley, J. A. and D. J. Boethel, Populations of Phytoseiulus persimilis (Acari: Phytoseiidae) and its host, Tetranychus urticae (Acari: Tetranychidae), on resistant and susceptible soybean cultivars, J. Econ. Entomol., 85:731-738, 1992. 147. Wheeler, G. S. and F. Slansky, Jr., Effect of constitutive and herbivore-induced extractables from susceptible and resistant soybean foliage on nonpest and pest noctuid caterpillars, J. Econ. Entomol., 84:1068-1079, 1991. 148. Wier, A. T. and D. J. Boethel, Feeding activity, growth, and survival of soybean looper on low-nitrogen nonnodulating soybean, Agron J., 86:1088-1091, 1994. 149. Wier, A. T. and D. J. Boethel, Feeding, growth, and survival of soybean looper (Lepidoptera: Noctuidae) in response to nitrogen fertilization of nonnodulating soybean, Environ. Entomol., 24:326-331, 1995. 150. Wolfenbarger, D. A. and J. P. Sleesman, Variation in susceptibility of soybean pubescent types, broad bean, and runner bean varieties and plant introductions to the potato leafhopper, J. Econ. Entomol., 56:895-897, 1963. 151. Yeargan, K. V., Potato leafhopper, In: L. G. Higley and D. J. Boethel, Eds., Handbook of Soybean Insect Pests, Entomol. Soc. Am., Lanham, 1994, 75. 152. Tillman, G., Personal communication. 1994.

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7

Germplasm Resources, Insect Resistance, and Grain Legume Improvement Stephen L. Clement, Massimo Cristofaro, Susan E. Cowgill, and Susanne Weigand

CONTENTS I. II. III. IV. V. VI.

Introduction World Production and Uses Insect Constraints to Production Germplasm Resources Plant Adaptation and Breeding Plant Resistance to Manage Insect Pests A. Specialist Insects B. Generalist Insects C. Genetically Engineered Plants D. New Avenues to Explore VII. Conclusions Acknowledgments References

I.

INTRODUCTION

Grain legumes or pulses, second only to the cereals in importance as food for humans and livestock, comprise a wide range of genera and species. Among species of this group used for food are warm and cool season food legumes. This chapter is devoted to cool season food legumes (CSFLs) which are generally considered to comprise four principal species: Cicer arietinum L. (chickpea), Pisum sativum L. (dry pea), Lens culinaris Medikus (lentil), and Vicia faba L. (faba bean).29,59 Phaseolus vulgaris L. (common bean), the most important warm season food legume, is the subject of Chapter 5 in this volume. Cool season food legumes have been companions of cereal grains since humans first began farming in the Near East at the close of the Pleistocene Age, 10,000 to 0-8493-2695-8/99/$0.00+$.50 © 1999 by CRC Press LLC

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12,000 years ago. Archeological evidence indicates that Near East peoples began growing lentils, peas, and chickpeas approximately 1,000 to 2,000 years after they started cultivating wheat and barley (>10,000 B.P.).27 Early civilizations apparently recognized the complementary role of cereals and legumes in their diets. This is exemplified by pea and lentil with wheat, Triticum aestivum L., and barley, Hordeum vulgare L., in the Near East, soybean, Glycine max (L.) Merr., and rice, Oryza sativa L., in the Orient, Vigna bean and sorghum, Sorghum bicolor (L.) Moench, in Africa, and Phaseolus bean and maize, Zea mays L., in the New World.96 The protein content of legumes (>20%) greatly exceeds that of the cereals and the amino acid profile of the protein is such that it complements that of cereals, thus ancient cultivation of cereals and CSFLs helped early civilizations meet their dietary requirements for essential amino acids. It remained for modern science to show that cereals and legumes are nutritionally complementary because “amino acids that are deficient in one are generally adequate in the other.”33 Lentils, peas, and chickpeas probably accompanied wheat and barley cultivation and the spread of Neolithic agriculture (9,000 B.P.) across the Mediterranean Basin and into central Europe.27,70 Faba bean appeared in Europe after chickpea, pea, and lentil (4,000 to 5,000 B.P.).8 These crops also radiated out to other areas: for example, peas to the Nile River Valley by 7,000 B.P.; peas, lentils, and chickpeas to the Indian subcontinent by 4,000 to 4,500 B.P.; chickpeas to Ethiopia 3,000 years ago; and faba bean to China during the last millennium.1,27,96 Worldwide, more than 60 insect species feed on chickpea and lentil6,65 and more than 20 species feed on faba bean and field pea.88 These insects contribute to global yield instability as a group, although only a few species are major pests.90 Among the major pests are seed beetles in the family Bruchidae. These bruchids or their chrysomelid ancestors are thought to have adapted to the seeds of leguminous plants during the Upper Jurassic or Cretaceous periods.21,39 Over evolutionary time, they developed specific host relationships with many legumes with some species becoming restricted to the seed of one legume.13,60 The Near East and adjacent areas of the Mediterranean Basin and Central Asia are centers of bruchid species diversity.9,89 From these regions, bruchids accompanied or followed the spread of CSFLs to new regions and continents where, with other insect pest groups, they attack these crops. Insecticides are often the management method of choice to control insect pests of CSFLs, especially in the developed countries.19,90 However, we should be able to endow the crops with resistance to some pests, thereby reducing the use of insecticides and the risk of environmental and health problems. Entomologists and plant breeders will have to turn to germplasm stocks should cool season food legume (CSFL) breeding programs emphasize the development of insect-resistant cultivars.

II.

WORLD PRODUCTION AND USES

World CSFL production is concentrated in temperate and subtropical climates, mainly at higher altitudes with very limited production in the warm lowland tropics. Within these geoclimatic areas, over 90% of the chickpea, lentil, dry pea, and faba bean production occurs in only 12 countries. Chickpeas and lentils are predominantly produced in the developing regions of the world (Indian subcontinent, West Asia,

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North and East Africa) and dry peas in the developed world (Europe, the Americas, Australia), with faba beans more evenly distributed (China, Europe, North Africa).59,71 World production, yield, and harvested area of chickpea, dry pea, and lentil increased modestly over the last 20 years (Table 1). By contrast, faba bean production has decreased worldwide over the last decade. Pea is the most widely cultivated of the four legumes and the world’s second most important pulse.80 Total production of the four crops increased from 18.6 mmt in 1975 to 24.7 mmt in 1985 (+33%), but only increased 6.5% to 26.3 mmt in 1995. While increasing, this production is well below the level needed to keep pace with world population growth, especially with the increased growth rates in the developing world.59 Each agricultural civilization has recognized CSFLs as important sources of natural fertilizer. By fixing atmospheric nitrogen through symbiosis with the root bacterium Rhizobium, these crops replenish the soil for subsequent cereal crops, thus reducing dependence on nitrogen fertilizers.27,96 Today, this contribution to soil fertility is a key factor in sustaining the production of cereals in the rainfed dry areas in the developing world.71 This practice also enhances yields of cereals grown in rotation with pulses in Australia and North America.59 Pea and faba bean crops are occasionally used for green manure worldwide. Winter field peas are a green manure crop in the U.S. Pacific Northwest.53 Moreover, faba bean seed is regarded as valuable animal feed in Europe and Asia96 and field peas are commonly grown and harvested for livestock feed in Australia and North America.59 Chickpeas, lentils, and faba beans are an important part of the diet of humans in developing nations, where they are mainly consumed after cooking as mature dry seeds or split cotyledons. In India, chickpea seeds are often processed into flour and combined with wheat flour to produce a better quality bread. Traditionally the pea crop was harvested to be eaten as dry peas, but today more of the crop, especially in Europe and North America, is grown for commercial canning or freezing.34 Worldwide, CSFLs are also consumed fresh in the form of immature tender pods, green seeds, or mature seeds.57

III.

INSECT CONSTRAINTS TO PRODUCTION

There are major impediments to increasing CSFL production, especially in the developing countries. Apart from poor cultivation practices, harvesting difficulties, and abiotic (e.g., low soil fertility, water deficits) and socio-economic factors (e.g., high labor costs, restrictive trade policies), CSFL production is constrained by weeds, diseases, nematodes, and insect pests. The occurrence of these biotic stresses vary across time, geography, and CSFL species. Moreover, the occurrence and severity of biotic stresses are often magnified by abiotic stresses.71,90,91 Yield and quality are also affected by insect depredation during storage.11 It is difficult to generate yield loss estimates for different biotic stresses because of high season to season variation. Weeds and diseases often rank ahead of insects as major production constraints.88 However, insect attacks in the field and during storage cause severe yield losses, quality problems, and greatly increase the cost of production. The literature provides some examples:

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TABLE 1 Trends in World Production, Yield, and Harvested Area of Cool Season Food Legumes 1975 Crop Chickpea Dry pea Faba bean Lentil

1985

1995

Area (000 ha)

Yield (kg/ha)

Production (000mt)

Area (000 ha)

Yield (kg/ha)

Production (000mt)

Area (000 ha)

Yield (kg/ha)

Production (000mt)

9,487 7,851 3,886 2,075

591 976 1,051 599

5,606 7,664 4,086 1,242

9,742 9,031 3,174 2,625

663 1,368 1,241 724

6,460 12,410 3,938 1,900

11,096 7,625 2,850 3,351

722 1,513 1,227 839

8,819 11,440 3,192 2,814

From FAOSTAT (Food Agriculture Organization of the United Nations, Production Statistics), on the World Wide Web (http://apps. fao.org/), 1997. (Ref. 25)

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1. Chickpea yield losses range from 0 to 36% in Syria91 and 10 to 40% in the former USSR41 from Liriomyza cicerina (Rondani) and losses of 14 to 20%83 and 6 to 42%65 from Helicoverpa armigera (Hübner) in India. 2. Faba bean seed losses of up to 28% have occurred from Sitona lineatus L. in Denmark54 and 10 to 90% from Bruchus dentipes Baudi in Syria,84 and abandonment of the crop in northern California because of severe damage by the seed bruchid B. rufimanus Boh.51 3. Pea seed losses of 10.6 to 71.5% in Australia31,50 and up to 63.9% in the U.S.62 from B. pisorum (L.), losses of 45.3% in Australia from H. punctigera Wallengren,50 and seed biomass reductions of 5 to 10% from S. lineatus in the U.S.93 4. Lentil seed quality problems (chalky spot) from Lygus bug feeding (Lygus spp.) in the U.S.58 5. Chickpea seed losses in storage of 7 to 70% in Syria and 24 to 100% in Jordan from Callosobruchus chinensis (L.) and C. maculatus (F.).92 Not surprisingly, the extensive range of yield loss estimates shown above reflect the differences between unsprayed and insecticide protected field plots. These estimated yield losses could be reduced through insecticide protection. Researchers have reported increased pea yields with insecticidal control of Acyrthosiphon pisum (Harris) in Burundi,55 Canada,48,81 and Ethiopia,90 and increased yields of lentil seed with insecticidal control of Sitona crinitus Herbst in Syria.35 Many of the pests discussed above are common throughout the world. Together with other important pests such as Aphis craccivora (Koch) (pest of all CSFLs), A. fabae Scop. (pest of faba bean),90 and B. lentis Frohl. (pest of lentil seed),5 these insects pose a major threat to CSFL production wherever the crops are grown.91

IV.

GERMPLASM RESOURCES

Major seed collections of CSFLs are maintained by the International Center for Agricultural Research in the Dry Areas (ICARDA), Aleppo, Syria; the International Crops Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru, India; and the U. S. Department of Agriculture, Agricultural Research Service (USDAARS), Western Regional Plant Introduction Station (WRPIS), Pullman, Washington (Table 2). The collections at the USDA-ARS National Seed Storage Laboratory, Fort Collins, Colorado, back-up the active collections (including CSFLs) at the Pullman Station and other USDA-ARS genebanks.68 Other genebanks in Ethiopia, Turkey, Italy, India, England, Mexico, The Netherlands, Sweden, Pakistan, Russia, Germany, and China store and maintain sizeable collections of one or more of the four crop species.52 Together, the world’s genebanks store approximately 36,000, 31,000, 20,000, and 14,300 accessions of Cicer, Pisum, V. faba, and Lens, respectively.52,76 Some repositories house special collections, exemplified by the Marx pea genetic stocks collection at the WRPIS genebank.75 This collection of 500 accessions of mutants and other well-characterized genetic stocks could offer potential sources of resistance to entomologists and breeders for basic studies and breeding (Chapter 12 in this volume). © 1999 by CRC Press LLC

TABLE 2 Genetic Resources of Cool Season Food Legumes at Major Genebanks and Their Evaluation for Insect Resistance

Institutea

Crop

Total no. of accessions in storage

ICARDA

Chickpea Wild chickpea Faba beans Lentil Wild lentil

9,682 292 9,703 7,477 434

ICRISAT

Chickpea

USDA-ARS-WRPIS

Chickpea Wild chickpea Faba beans Lentil Wild lentil Pea Wild pea

17,244 4,412 168 436 2,710 153 4,419 48

No. of accessions evaluated (no. found resistant or tolerant to insects) 7,000 200 952 93 0

(10) b; 6,697 (0) c (30) b; 137 (51) c (0) d; 7156 (114) e (21) f

14,800 (10) g 0 0 0 0 0 2,074 (4) h; 1,571 (10) i 33 (14) i

a

ICARDA, International Center for Agricultural Research in the Dry Areas, Aleppo, Syria; ICRISAT, International Crops Research Institute for the Semi-Arid Tropics, Andhra Pradesh, India; USDA-ARSWRPIS, United States Department of Agriculture, Agricultural Research Service, Western Regional Plant Introduction Station, Pullman, Washington, U.S. b Liriomyza cicerina Rondani, Ref. 77. c Callosobruchus chinensis (L.), Ref. 92. d Bruchus dentipes Baudi, Ref. 85. e Aphis craccivora Koch, Ref. 24. f Aphis craccivora Koch and Acyrthosiphon pisum (Harris), Ref. 36. g Helicoverpa armigera (Hübner), Ref. 47. h Sitona lineatus L., Ref. 56. i Bruchus pisorum L., Refs. 14, 17, and 62.

With cultivated forms making up the greater part of the largest CSFL collections, more emphasis should be placed on collection of the wild relatives which are underrepresented. Less than 3% of the total CSFL collections (excluding faba bean) at ICARDA, ICRISAT, and the WRPIS genebanks are composed of wild Cicer, Lens, and Pisum species (Table 2). Wild Vicia species are not included in the total because the progenitor of faba bean is unknown.96 Because only a very small proportion of the CSFL accessions are wild relatives, the full genetic variability of these species is not fully represented in these collections. The presence of insect-resistant genes in the wild relatives of many crop species indicates the likelihood of useful resistance genes in heterogenous populations of wild CSFL species.42,94

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V.

PLANT ADAPTATION AND BREEDING

Cool season food legumes, like all crop plants, evolved in parallel with human societies. As early civilizations began to plant and harvest seeds of these crops, they inadvertently began to alter the gene pool as they selected for “crop types.” By harvesting only nonshattering pods, early farmers selected for nondehiscence, more determinate growth, and increased seed production.27 Over time, farmers went from unconscious to conscious selection of plant traits that transformed wild forms into productive cultivars that provided for their specific needs. Other traits commonly associated with the process of domestication in CSFLs include elimination of seed dormancy, increased seed size, a trend toward smooth and thin seed coats, improved seed germination and seedling vigor, and the evolution of more erect growth forms.80,96 The main objective of modern plant breeding is the development of highyielding, high-quality cultivars. Disease resistance, especially in chickpea and pea, as well as adaptation to limited moisture conditions rank high among aims in CSFL breeding.52 Although entomologists and plant breeders have identified sources of resistance to several of the most important insect pests in CSFLs, the fruits of their research have yet to reach farmers in the form of insect-resistant cultivars.15 Wild grain legumes are thought to have natural chemical defenses in the form of potent toxins and antimetabolites that are inhibitory to insects and mammalian herbivores.96 Cultivars frequently lack or contain reduced amounts of these chemical defenses which increase their vulnerability to insect attack and the need for insecticidal control.96 Genes for insect resistance have probably been lost under intense selection for high yield, wider adaptability, and improved nutritional value. Consistent with this hypothesis is the view that pigmentation in chickpeas is associated with insect and disease resistance and that its loss during sustained selection for advanced kabuli chickpeas with their large and cream-colored testas has increased susceptibility to insect and disease attack.80 Indeed, some kabuli types are so susceptible to Helicoverpa attack that few or no pods survive unless the plants are chemically protected, whereas less advanced desi chickpeas with their brown to yellow seed testas are less susceptible.64

VI.

PLANT RESISTANCE TO MANAGE INSECT PESTS

Cool season food legumes are often grown without insecticide protection in developing countries because insecticides are generally unavailable or too expensive.12,65 Therefore, there is much interest in CSFL breeding for pest resistance in these countries. Insecticides are readily available in developed countries where their success in controlling pests of CSFLs has lulled farmers and researchers into a false sense of security, causing a loss of urgency in the development of insect-resistant and tolerant cultivars. However, with widespread insecticide use comes the risk of health and environmental problems, high mortality of natural enemies of pests, and, eventually, the build-up of insecticide resistance in the pests.46 Farmers also must contend with increased scrutiny and registration of pesticide use by government

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agencies. As a consequence, new demands are being placed on farmers and agricultural scientists to develop and adopt more environmentally sound and economically viable pest management programs. Plant resistance to pests is the cornerstone of such programs. Plant resistance to at least 17 field and storage insect pests of CSFLs has been identified. Searches for resistance to an individual pest have involved as few as 2, normally 6 to 140, and at times more than 14,000 germplasm accessions or entries.15 For the most part, these screening programs evaluated germplasm collections from ICARDA, ICRISAT, and WRPIS for resistance to the most important pests of CSFLs (Table 2). These pests included both specialist (restricted to one plant genus) and generalist (wide host range) insects (see below). We also must add the recent screenings of 1900 Pisum accessions from 9 germplasm collections for resistance to B. pisorum28 and 37 Pisum accessions from genebanks in Russia (Vavilov Institute) and Sweden (Svalöf, Nordic Gene Bank) for resistance to A. pisum.69

A. SPECIALIST INSECTS Resistance to the pea specialist B. pisorum was found in pea germplasm,16,62 but this antixenosis resistance has limited value because it did not hold up under field conditions in Chile and Australia.15 Genetic resistance to B. dentipes was not located in conserved faba bean germplasm.85 Searches of germplasm stocks for resistance to the faba bean specialist B. rufimanus and the lentil specialist B. lentis have not been undertaken to our knowledge. With no useable resistance to B. pisorum and B. dentipes heretofore discovered in the primary genepools of pea and faba bean, the progenitors and wild relatives of these crops may be the only source of genetic variation for the development of resistant cultivars. The recent discovery of strong antibiosis resistance in wild peas (P. fulvum Sibth. & Sm.) to B. pisorum17,28 suggests that the wild relatives of lentil (L. culinaris ssp. orientalis and ssp. odemensis, and L. nigricans ssp. nigricans and ssp. ervoides) and faba bean (V. narbonensis L. and other species in the tertiary gene pool) would be a good place to search for genetic resistance to B. lentis, B. dentipes, and B. rufimanus. It is necessary to search the tertiary gene pool of faba bean because its progenitor has not been found and there is no known species in the secondary genepool.96 Host specificity of these univoltine bruchids is largely determined by feeding of the adults on pollen and nectar of their primary hosts, which is very important for maturation of gonads and subsequent reproductive activity and oviposition. Females of B. lentis require lentil pollen and nectar for egg formation,61 both sexes of B. dentipes feed on pollen and nectar of faba beans before oviposition takes place,84 and ingestion of faba bean pollen terminates diapause and induces vitellogenesis in female B. rufimanus. Additionally, female B. rufimanus become sexually active when faba bean pods become available.87 Although pollen of a number of species other than pea promotes ovarian development in female B. pisorum, pea pollen is most effective in promoting oogenesis.63 That females of each species lay eggs only on the green pods of their hosts is further evidence of the highly specific nature of these four bruchid-host plant relationships.13,60,84,87

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It has been hypothesized that these very close relationships are the result of coevolution that ensures the synchronization of bruchid reproduction and development with host plant availability and bruchid diapause during periods of host plant unavailability in the field.32,60 Furthermore, Labeyrie44 used specific host plant relationships involving bruchids to support his view that “the more an insect is specialized, the more coevolutionary processes are likely to be important and the more the effect of the plant upon different aspects of reproduction is to be suspected.” However, without evidence for reciprocal evolutionary change, these highly specific relationships cannot be interpreted as a result of coevolutionary history.86 Yet coevolutionary assumptions of these relationships are important because they could influence decisions regarding the search for genetic resistance in plant genetic resources. The high frequency of the Np gene among wild pea accessions from regions where the B. pisorum is native and exerts high pressure on its host may reflect a counter response in P. sativum to this pressure.4 Plants with this gene respond to the presence of pea weevil eggs on pods by forming callus,4,22 which may confer a measure of resistance since the callus appears to impede larval penetration of the pod wall.4,14 Therefore, it is important to consider the possible accumulation of resistance genes over evolutionary time and their potential utility in breeding for gene-based resistance to specialist bruchids. If highly specific relationships limit the adaptive possibilities of legume bruchids, non-host species in the secondary and tertiary gene pools of grain legumes may be the best sources of resistance genes.

B. GENERALIST INSECTS Screening of over 14,000 chickpea germplasm accessions from the ICRISAT genebank (Table 2) showed that chickpea genotypes vary in their susceptibility to H. armigera,45 the dominant pest of chickpea and several other crops in India.20,65 Although entomologists and chemists have learned much about the mechanisms involved in H. armigera resistance in chickpeas20,66,82,95 and the inheritance of resistance,37 the resistant genotypes selected from germplasm and the resistant lines developed from crosses are susceptible to fusarium wilt.65,79 At ICRISAT, crosses have been made to combine fusarium wilt and H. armigera resistance, but the level of insect resistance in many of the progeny is not comparable to that in the resistant parent lines. This inability to combine insect and disease resistance has stymied the development and release of regionally adapted insect-resistant cultivars. Approximately 7000 chickpea germplasm and breeding lines have been evaluated at ICARDA for resistance to the leaf miner L. cicerina, the most important pest of chickpea in Western Asia and North Africa. Ten lines were rated moderately resistant, with three kabuli lines consistently rated resistant in subsequent trials at ICARDA.77,91 In 1994, these three lines were registered as germplasm lines and released to breeding programs by ICARDA and ICRISAT.78 An additional 200 accessions of 8 annual wild Cicer species have been evaluated for leaf miner resistance, of which 35 may be considered good sources of resistance.77 There is interest in using some of these resistant lines in the chickpea breeding program at ICARDA.75

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Conserved germplasm has been evaluated for resistance to other generalist insect pests; namely, peas to S. lineatus,56 faba beans to A. craccivora,24 and lentils to A. craccivora and A. pisum,36 with the result that some accessions were rated resistant or tolerant to these insects (Table 2). Unfortunately, efforts to develop useful levels of plant resistance in peas to S. lineatus were not successful,3 leading to a cessation of breeding efforts against this weevil in the U.S.15 The nature and practical value of aphid resistance in faba beans and lentils are unknown. However, the incorporation of high levels of antibiosis from wild Vicia spp. into faba bean cultivars was viewed by Holt and Birch30 as a possible long term solution to the development of virulent, resistance-breaking aphid biotypes. Extensive screening of Cicer spp. germplasm for variation in susceptibility of seed to the storage pest C. chinensis has identified resistance only in wild species (Table 2).91

C. GENETICALLY ENGINEERED PLANTS The first success in genetically engineering a CSFL for insect resistance followed the development of a reproducible transformation and regeneration system for the introduction of foreign genes into cultivars of peas,73 and involved the transfer of a pest-resistant trait in the seeds of common bean (P. vulgaris) into pea for resistance to the storage pests C. maculatus and C. chinensis.74 This resistance is based on a gene for a seed protein called α-amylase inhibitor in common bean, which blocks the action of the starch-digesting enzyme α-amylase and thus prevents weevil larvae from digesting starches in the seeds. Transgenic peas that expressed the bean α-amylase inhibitor gene in their seeds also exhibited resistance to the pea weevil B. pisorum.72 This research places pea at the forefront in the biotechnological improvement of legumes for insect resistance. In subsequent phases of this research, genetic engineering of pea and other CSFLs for resistance to other insect pests is envisaged. A better understanding of secondary plant compounds in legume seeds and their protective role against insect pests of CSFLs will increase the possibilities for gene introgression via biotechnology.10,26 The recent screening of several hundred bean germplasm lines and the discovery of a large diversity of α-amylase inhibitors in the seed illustrate the importance of conserved germplasm as a source of new insecticide genes for plant biotechnology.38

D. NEW AVENUES

TO

EXPLORE

What is the potential that searches of germplasm stocks will widen the arsenal of plant defensive traits for possible use in CSFL breeding? Accumulating evidence suggest the potential for widening the arsenal of plant defensive traits to pests is good (Table 3). The neoplastic pod trait in peas (Np gene) mentioned earlier in this chapter and listed in Table 3 was detected in pea germplasm from the Marx genetic stocks collection at the WRPIS genebank. It is thought the Np allele regulates the expression of a plant defensive trait (callus production on pods) in response to attack (oviposition) by B. pisorum.4,22 Likewise, pea germplasm can be a source of genes responsible for plant phenotypes that differ in their resistance to herbivores. A study found that beetle predators were more effective at finding and controlling pea aphids

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TABLE 3 Selected Plant Characters with Potential or Demonstrated Value in Moderating Insect Attacks on Pea, Chickpea, and Faba Bean Crop Pea

Chickpea

Faba bean

Plant attribute

Genes

Leafless phenotype

af, st

Semi-leafless phenotype Glossy wax mutant

af wel

Neoplastic pod

Np

Plant volatiles

?

Trichome acid exudates

?

Plant volatiles

?

Plant volatiles

?

Potential or demonstrated effect on insect Enhances ladybird beetle predation of pea aphids (Acyrthosiphon pisum) Lowers pea aphid densities Enhances ladybird beetle predation of pea aphids Alters behavior of pea weevil (Bruchus pisorum) larvae on pod surface Mediates host-selection behavior of adult pea weevils Antibiosis effects on larvae of gram pod borer (Helicoverpa armigera) Mediates host-selection behavior of larvae and adult gram pod borers Synergism with aggregation pheromone of pea leaf weevil (Sitona lineatus) for greater weevil attraction

Ref. 40 82 Chapter 12 4, 14, 22 2, 18 66, 95 67 7

on leafless phenotypes compared to normal-leafed peas.40 Other researchers found fewer pea aphids on semi-leafless peas, which they attributed to increased vulnerability of aphids on these architecturally simple plants to adverse weather and to the reduction of preferred feeding space (leaflets) for aphid population growth.81 Finally, research suggests a role for glossy wax mutants in the Marx pea collection for genetic resistance to pea aphid via enhanced beetle predation of the aphid prey (Chapter 12 in this volume). Phytochemicals play an important role in interactions between plants and phytophagous insects and should be an important factor in breeding crops for insect resistance.43 With only a few studies, it is difficult to draw any conclusions about the potential importance of CSFL phytochemicals to resistance breeding. What is apparent from these studies, however, is that volatile chemicals from pea, chickpea, and faba bean and contact chemicals (trichome exudates) from chickpea play a decisive role in regulating the behavior and host-selection of three important pests of these crops (Table 3). Because many plants produce a rich menu of volatile chemicals49 and trichome exudates,23 it seems fair to say that chemical prospecting of CSFL germplasm stocks would greatly increase our understanding of host resistance factors. Knowledge of chemical defenses in CSFLs and the genes that control their production could enhance the selection of pea, chickpea, and faba bean genotypes with reduced insect attractiveness via low production of volatile kairomones, and may also help in breeding chickpea lines with specific contact chemicals that would confer insect resistance.67

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VII.

CONCLUSIONS

The germplasm stocks of CSFLs have been indispensable for insect resistance screening and they are beginning to be searched for new plant defensive traits for potential use in breeding resistant cultivars (Table 3). Although cultivars with germplasm lines in their pedigrees have not been released, there is optimism that regionally adapted cultivars of chickpeas with H. armigera and L. cicerina resistance will eventually be developed and released to farmers in the developing world. This is based on research at ICARDA and ICRISAT, which led to the registration of chickpea germplasm lines with leafminer resistance. Beyond the primary gene pools, the wild relatives of CSFLs in genebanks appear to be fruitful sources of insect resistance genes, exemplified by resistance in wild chickpeas to L. cicerina and C. chinensis, resistance in wild peas to B. pisorum (Table 2), and resistance in wild Vicia spp. to aphids.30 Several biological and technological factors constrain the transfer of resistance-conferring genes from wild plants and other nonadapted germplasm to adapted backgrounds. These include insufficient information about the chemical and physical nature and genetic bases of plant resistance to insects, the need for research to overcome barriers to the development of cultivars with multiple insect and disease resistance, and the requirement for new and improved technology to overcome barriers to inter-specific hybridization.15,64 But in the long run, progress in insect resistance breeding in CSFLs will depend on a stronger commitment by national and international research programs. Biotechnology research in Australia and the U.S. has broadened the genetic resource base for insect resistance genes and the possibilities for transferring these genes from alien plants to cultivars of CSFLs. These efforts illustrate the power of interdisciplinary research involving molecular biologists, plant physiologists, and entomologists.72,74 Team research like this must continue and be expanded to apply biotechnological innovations to the development of insect-resistant cultivars. The world’s germplasm stocks are likely to become more important as the main supplier of germplasm for insect resistance through both conventional plant breeding and plant biotechnology. Societal pressures for more biologically based pest management practices in cropping systems will place a higher value on global genetic resources. Utilizing plant defense mechanisms is the most economical way to control insect pests and the best way to mitigate the unpredictability of insect attacks on CSFLs while at the same time boosting the yields of these crops in the face of increasing human populations.

ACKNOWLEDGMENTS The first author is grateful to L. Elberson for general assistance and C. Simon and D. Stout for information on germplasm holdings at the USDA-ARS Western Regional Plant Introduction Station. This research was supported in part by a grant from the USDA-FAS-ICD-RSED (AS37).

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39. Johnson, C. D. Seed beetle host specificity and the systematics of the Leguminosae, In: R. M. Polhill and P. H. Raven, Eds., Advances in Legume Systematics, Part 2, Royal Botanic Gardens, Kew, 1981, 995. 40. Kareiva, P. and R. Sahakian, Tritrophic effects of a simple architectural mutation in pea plants, Nature, 345:433-434, 1990. 41. Kay, D. E., Food Legumes. Crop and Product Digest No. 3, Tropical Products Institute, London, 1979. 42. Kennedy, G. G. and J. D. Barbour, Resistance variation in natural and managed systems, In: R. S. Fritz and E. L. Simms, Eds., Plant Resistance to Herbivours and Pathogens, Ecology, Evolution, and Genetics, Univ. Chicago Press, Chicago, 1992, 13. 43. Kogan, M., Natural chemicals in plant resistance to insects, Iowa State J. Res., 60:501-527, 1986. 44. Labeyrie, V., Reproduction of insects and coevolution of insects and plants, Entomol. Exp. Appl., 24:296-304, 1978. 45. Lateef, S. S., Gram pod borer Heliothis armigera (Hub.) resistance in chickpeas. Agric. Ecosyst. & Environ., 14:95-102, 1985. 46. Lateef, S. S., Scope and limitations of host plant resistance in pulses for the control of Helicoverpa, In: J. N. Sachan, Ed., First National Workshop on Heliothis Management: Current Status and Future Strategies, Division of Entomology, Directorate of Pulses Research, Kanpur, 1990, 129. 47. Lateef, S. S. and M. P. Pimbert, The search for host-plant resistance to Helicoverpa armigera in chickpea and pigeonpea at ICRISAT, In: Proc. 1st Consultative Group on Host Selection Behavior of Helicoverpa Armigera, ICRISAT, Andhra Pradesh, 1990, 14. 48. Maiteki, G. A. and R. J. Lamb, Spray timing and economic threshold for the pea aphid, Acyrthosiphon pisum (Homoptera: Aphididae), on field peas in Manitoba, J. Econ. Entomol., 78:1449-1454, 1985. 49. Metcalf, R. L. and E. R. Metcalf, Plant Kairomones in Insect Ecology and Control, Chapman and Hall, New York, 1992. 50. Michael, P. J., D. C. Hardie, G. P. Mangano, T. P. Quinn, and I. A. Pritchard, The effectiveness of chemicals against the pea weevil, Bruchus pisorum (L.), and native budworm, Helicoverpa punctigera Wallengren, on field peas, Pisum sativum L., in western Australia, In: A.M. Smith, Ed., National Pea Weevil Workshop, Victoria Department of Agriculture and Rural Affairs, Melbourne, 1990, 41. 51. Middlekauff, W. W., Field studies on the bionomics and control of the broad bean weevil, J. Econ. Entomol., 44:240-243, 1951. 52. Muehlbauer, F. J. and W. J. Kaiser, Using host plant resistance to manage biotic stresses in cool season food legumes, In: F. J. Muehlbauer and W.J. Kaiser, Eds., Expanding the Production and Use of Cool Season Food Legumes, Kluwer Academic, Dordrecht, 1994, 233. 53. Murray, G. A., K. D. Kephart, L.E. O’Keeffe, D. L. Auld, and R. H. Callihan, Dry pea, lentil and chickpea production in northern Idaho, Agric. Exp. Stn. Bull. No. 664, University of Idaho, Moscow, 1987. 54. Nielsen, B. S., Yield responses of Vicia faba in relation to infestation levels of Sitona lineatus L. (Col., Curculionidae), J. Appl. Entomol., 110:398-407, 1990. 55. Nijimbere, M., M. Kayibigi, A. Autrique, and A. D. Malithano, Factors which limit production of pea in Burundi, In: R. J. Summerfield, Ed., World Crops: Cool Season Food Legumes, Kluwer Academic, Dordrecht, 1988, 159. 56. Nouri-Ghanbalani, G., Host plant resistance to the pea leaf weevil, Sitona lineatus (L.), in pea, Pisum sativum L., and its inheritance. Ph.D. Thesis, University of Idaho, Moscow, 1977.

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57. Nwokolo, E. and J. Smartt, Eds., Food and Feed from Legumes and Oilseeds, Chapman & Hall, London, 1996. 58. O’Keeffe, L. E., H. W. Homan, and D. J. Schotzko, Chalky spot damage to lentils, Current Information Series No. 894, University of Idaho, College of Agriculture, Moscow, 1991. 59. Oram, P. A. and M. Agcaoili, Current status and future trends in supply and demand of cool season food legumes, In: F. J. Muehlbauer and W. J. Kaiser, Eds., Expanding the Production and Use of Cool Season Food Legumes, Kluwer Academic, Dordrecht, 1994, 3. 60. Pajni, H. R., Ecological status of host range and polymorphism in Bruchidae, In: E. Donahaye and S. Navarro, Eds., Proc. 4th Int. Working Conf. Stored Product Protection, Agricultural Research Organization, Bet Dagan, 1986, 506. 61. Pajni, H. R. and K. Mittal, Observations on infestation of Lens culinaris by Bruchus lentis in the Chandigarh area of India, LENS Newsl., 11:27-28, 1984. 62. Pesho, G. R., F. J. Muehlbauer, and W. H. Harberts, Resistance of pea introductions to the pea weevil, J. Econ. Entomol., 70:30-33, 1977. 63. Pesho, G. R. and R. J . van Houten, Pollen and sexual maturation of the pea weevil (Coleoptera: Bruchidae), Ann. Entomol. Soc. Am., 75:439-443, 1982. 64. Reed, W., C. Cardona, S. S. Lateef, and S. I. Bishara, Screening and breeding for insect resistance in pea, lentil, faba bean and chickpea, In R. J. Summerfield, Ed., World Crops: Cool Season Food Legumes, Kluwer Academic, Dordrecht, 1988, 107. 65. Reed, W., C. Cardona, S. Sithanantham, and S. S. Lateef, Chickpea insect pests and their control, In: M. C. Saxena and K. B. Singh, Eds., The Chickpea, Kluwer Academic, Dordrecht, 1987, 283. 66. Rembold, H., Malic acid in chickpea exudate-a marker for Heliothis resistance, Int. Chickpea Newsl., 4:18-19, 1981. 67. Rembold, H., P. Wallner, A. Köhne, S. S. Lateef, M. Grüne, and Ch. Weigner, Mechanisms of host-plant resistance with special emphasis on biochemical factors, In: H. A. van Rheenen, M. C. Saxena, B. J. Walby, and S. D. Hall, Eds., Chickpea in the Nineties, ICRISAT, Andhra Pradesh, 1990, 191. 68. Roos, E. E., Long-term seed storage, In: J. Janick, Ed., Plant Breeding Reviews, Volume 7, The National Plant Germplasm System of the United States, Timber Press, Portland, 1989, 129. 69. Sandström, J., High variation in host adaptation among clones of the pea aphid, Acyrthosiphon pisum, on peas, Pisum sativum, Entomol. Exp. Appl., 71:245-256, 1994. 70. Sauer, J. D., Historical Geography of Crop Plants, CRC Press, Boca Raton, 1993. 71. Saxena, M. C., The challenge of developing biotic and abiotic stress resistance in cool-season food legumes, In: K. B. Singh and M. C. Saxena, Eds., Breeding for Stress Tolerance in Cool-Season Food Legumes, Wiley, Chichester, 1993, 3. 72. Schroeder, H. E., S. Gollasch, A. Moore, L. M. Tabe, S. Craig, D. C. Hardie, M. J. Chrispeels, D. Spencer, and T. J. V. Higgins, Bean-amylase inhibitor resistance to the pea weevil (Bruchus pisorum) in transgenic peas (Pisum sativum L.), Plant Physiol., 107:1233-1239, 1995. 73. Schroeder, H. E., A. H. Schotz, T. Wardley-Richardson, D. Spencer, and T. J. V. Higgins, Transformation and regeneration of two cultivars of pea (Pisum sativum L.), Plant Physiol., 101:751-757, 1993. 74. Shade, R. E., H. E. Schroeder, J. J. Pueyo, L. M. Tabe, L. L. Murdock, T. J. V. Higgins, and M. J. Chrispeels, Transgenic pea seeds expressing the alpha-amylase inhibitor of the common bean are resistant to bruchid beetles, Bio/Tech., 12:793-796, 1994.

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75. Simon, C., Current status of the USDA Pisum P.I. and genetic stocks collections, Abstr., Biennial Meeting Nat. Pea Improv. Assoc., The National Pea Improvement Association, East Lansing, 1995, 16. 76. SINGER (System-wide Information Network on Genetic Resources), Consultative Group on International Agricultural Research, on the World Wide Web (http://nocl. cgiar.org/cgintrus.htm), 1997. 77. Singh, K. B. and S. Weigand, Identification of resistant sources in Cicer species to Liriomyza cicerina, Genetic Resources and Crop Evolution, 41:75-79, 1994. 78. Singh, K. B. and S. Weigand, Registration of three leafminer-resistant chickpea germplasm lines: ILC 3800, ILC 5901, and ILC 7738, Crop Sci., 36:472, 1996. 79. Singh, O., S. C. Sethi, S. S. Lateef, and C. L. L. Gowda, Registration of ICCV 7 chickpea germplasm, Crop Sci., 37:295, 1997. 80. Smartt, J., Grain Legumes, Evolution and Genetic Resources, Cambridge University Press, Cambridge, 1990. 81. Soroka, J. J. and P. A. MacKay, Seasonal occurrence of the pea aphid, Acyrthosiphon pisum (Harris) (Homoptera: Aphididae), on cultivars of field peas in Manitoba and its effect on pea growth and yield, Can. Entomol., 122:503-513, 1990. 82. Srivastava, C. P. and R. P. Srivastava, Screening for resistance to the gram pod borer, H. armigera, in chickpea: genotypes and observations on its mechanism of resistance in India, Insect Sci. Appl., 10:255-258, 1989. 83. Srivastava, C. P. and R. P. Srivastava, Estimation of avoidable loss in chickpea (Cicer arietinum) due to gram-pod borer (Heliothis armigera) in Rajasthan. Indian J. Agric. Sci., 60:494-496, 1990. 84. Tahhan, O. and H. F. van Emden, Biology of Bruchus dentipes Baudi (Coleoptera: Bruchidae) on Vicia faba and a method to obtain gravid females during the imaginal quiescence period, Bull. Entomol. Res., 79:201-210, 1989. 85. Tahhan, O. and H. F. van Emden, Resistance of faba bean, Vicia faba, to Bruchus dentipes Baudi (Coleoptera: Bruchidae), Bull. Entomol. Res., 79:211-218, 1989. 86. Thompson, J. N., Interaction and Coevolution, John Wiley, New York, 1982. 87. Tran, B. and J. Huignard, Interactions between photoperiod and food affect the termination of reproductive diapause in Bruchus rufimanus (Boh.) (Coleoptera: Bruchidae), J. Insect Physiol., 8:633-642, 1992. 88. van Emden, H. F., S. L. Ball, and M. R. Rao, Pest, disease and weed problems in pea, lentil, faba bean and chickpea, In: R. J. Summerfield, Ed., World Crops: Cool Season Food Legumes, Kluwer Academic, Dordrecht, 1988, 519. 89. Watanabe, N., Diversity in life cycle patterns of bruchids occurring in Japan (Coleoptera: Bruchidae), In: K. Fujii, A. M. R. Gatehouse, C. D. Johnson, R. Mitchel, and T. Yoshiuda, Eds., Bruchids and Legumes: Economics, Ecology and Coevolution, Kluwer Academic, Dordrecht, 1990, 141. 90. Weigand, S., S. S. Lateef, N. El-Din Sharaf El-Din, S. F. Mahmoud, K. Ahmed, and K. Ali, Integrated control of insect pests of cool season food legumes, In: F. J. Muehlbauer and W. J. Kaiser, Eds., Expanding the Production and Use of Cool Season Food Legumes, Kluwer Academic, Dordrecht, 1994, 679. 91. Weigand, S. and M. P. Pimbert, Screening and selection criteria for insect resistance in cool-season food legumes, In: K. B. Singh, and M. C. Saxena, Eds., Breeding for Stress Tolerance in Cool-Season Food Legumes, John Wiley, Chichester, 1993, 145. 92. Weigand, S. and O. Tahhan, Chickpea insect pests in the Mediterranean zones and new approaches to their management, In: H. A. van Rheenen, M. C. Saxena, B. J. Walby, and S. D. Hall, Eds., Chickpea in the Nineties, ICRISAT, Andhra Pradesh, 1990, 169.

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93. Williams, L., III, D. J. Schotzko, and L. E. O’Keeffe, Pea leaf weevil herbivory on pea seedlings, Entomol. Exp. Appl., 76:255-269, 1995. 94. Yencho, G. C., M. W. Bonierbale, W. M. Tingey, R. L. Plaisted, and S. D. Tanksley, Molecular markers locate genes for resistance to the Colorado potato beetle, Leptinotarsa decemlineata, in hybrid Solanum tuberosum x S. berthaultii potato progenies, Entomol. Exp. Appl., 81:141-154, 1996. 95. Yoshida, M., S. E. Cowgill, and J. A. Wightman, Mechanisms of resistance to Helicoverpa armigera (Lepidoptera: Noctuidae) in chickpea: role of oxalic acid in leaf exudate as an antibiotic factor, J. Econ. Entomol., 88:1783-1786, 1995. 96. Zohary, D. and M. Hopf, Domestication of Plants in the Old World, Clarendon Press, Oxford, 1993.

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8

Alfalfa Germplasm Resources and Insect Resistance George R. Manglitz and Edgar L. Sorensen

CONTENTS I. II. III. IV.

Introduction The Genus Medicago Plant Introductions and Germplasm Resources Insect Resistance A. Nature of Resistance B. Breeding Techniques C. Specific Insects 1. Potato Leafhopper 2. Aphids 3. Alfalfa Weevil 4. Other Insects V. Conclusions References

I.

INTRODUCTION

It is thought that alfalfa, Medicago sativa L., originated in Asia Minor, Transcaucasia, Iran, and the highlands of Turkmenistan. Alfalfa was cultivated before recorded history and is now found growing wild from China to Spain and from Sweden to North Africa. In addition, it has become acclimatized in South Africa, Australia, New Zealand, and North and South America.81 Alfalfa is concentrated in certain zones within the northern hemisphere, i.e., U.S., Canada, Italy, France, China, southern Russian Federation, and in selected countries in the southern hemisphere, i.e., Argentina, Chile, South Africa, Australia, and New Zealand. Alfalfa can be grown as far as the 60° N Lat but it is essentially a crop of temperate regions. The U.S., Russian Federation, and Argentina contribute about 70% to the world estimate of approximately 32 million ha. France, Italy, Canada, and China combine to account for an additional 17%. Michaud et al.81 list 50 countries and the hectares of alfalfa in each.

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Alfalfa is an extremely energy efficient crop to grow, providing high yields of forage of exceptional nutritional value whether consumed by livestock as pasturage, green chop, hay, silage, or incorporated into pellets or other feeds. It improves soil structure and tilth and is important for soil conservation and reclamation. Alfalfa may have increasingly larger roles in low input sustainable agriculture.100 A large number of insect species are associated with alfalfa fields, many of which are beneficial to the crop. The beneficials include insects essential to the pollination of alfalfa flowers and predators and parasitoids of herbivorous species. Nevertheless, more than 100 species damage the crop in the U.S.,2 northeast Africa, and southeast Asia.36 Nielson and Lehman90 list 32 of the more important insect pests of alfalfa along with their worldwide distribution and approximate pest status. A general review of the pest insects of alfalfa in the U.S. was published in 1988.74 In 1972, Sorensen et al.126 reported cultivars with resistance to the spotted alfalfa aphid, Therioaphis trifolii (Monell), and the pea aphid, Acyrthosiphon pisum (Harris), and that progress had been made in locating sources of resistance to the meadow spittlebug Philaenus spumarius (L.), potato leafhopper Empoasca fabae (Harris), and alfalfa weevil Hypera postica (Gyllenhal). Eight years later, Nielson and Lehman90 reported additional progress with these and other insects and discussed the emerging problem of resistance breaking biotypes of insects, particularly the spotted alfalfa aphid. After another eight years, Sorensen et al.110 reported more than 100 cultivars with resistance to the pea aphid, the spotted alfalfa aphid, or both. Some of these cultivars also carry resistance to the blue alfalfa aphid, Acyrthosiphon rondoi Shinji. The use of insect-resistant cultivars to control insects that damage both quantity and quality of the alfalfa crop is a well-established practice. Insect control is achieved without additional costs to the producer and without additional manipulations, such as the treatment of the crop with insecticides. The use of insect-resistant alfalfas is also compatible with most other methods of insect control including the use of parasites and predators. The complete control of insects and their damage also is not as important in the case of a forage crop as it would be with a fruit or vegetable crop where consumers may demand an insect-free product (Chapter 9 in this volume).

II.

THE GENUS MEDICAGO

The genus Medicago contains more than 60 species, two-thirds of which are annuals and one-third perennials. The perennial species are cross-fertilized, whereas the annual species are self-fertilized. In conformity with their breeding system, progenies of annual plants are remarkably uniform.97,98 Conversely, most perennial species are allogamous with different degrees of self-incompatibility. Although some selftripping occurs in these species, they rely on insect pollinators to activate their flower tripping and pollination. Most Medicago species are quite polymorphic because of their high degree of outcrossing. Their pollinating agents include numerous species of bees. Diploid, tetraploid, and hexaploid species are found in Medicago. Most of the diploids are of the 2N = 16 type with a basic chromosome number of X = 8. Five species of diploids have 2N = 14 chromosomes, namely M. constricta Dur., M. praecox D.C., M. polymorpha L., M. rigidula (L.) All., and M. murex Willd. Two annual species, M. scutellata (L.) Mill. and M. rugosa Desr., have 2N = 30. The © 1999 by CRC Press LLC

hexaploids are M. cancellata M.B. and M. saxatilis M.B., both perennials. A number of perennial species have both diploid and tetraploid forms. The tetraploid species probably arose via unreduced gametes of the diploids. It is thought that perennials with a woody growth habit are phylogenetically the oldest, followed by herbaceous perennials with annuals the youngest.69,97 Taxa that hybridize freely with alfalfa constitute the M. sativa complex. They are classified as subspecies of M. sativa on the basis of morphological and ploidy criteria. Quiros and Bauchan97 included the following: ssp. sativa (L.) L. & L., ssp. coerulea Schmalh., ssp. falcata Arcengeli, ssp. varia Arcengeli, ssp. x hemicycla, ssp. x polychroa, ssp. x tunetana, and ssp. glutinosa M.B. The species M. glomerata Balb. and M. prostrata Jacq. are closely related to this complex.

III.

PLANT INTRODUCTIONS AND GERMPLASM RESOURCES

Alfalfa was introduced to eastern U.S. as early as 1736 but failed to become a major crop. Then in 1897, the first official plant explorer, N. E. Hansen of the U.S. Department of Agriculture (USDA), obtained 18 accessions of alfalfa from the Russian Federation. These plant introductions (PI) acquired early in the twentieth century had a major effect on alfalfa production in the U.S. Some were released as cultivars: “Cossack” (PI20714, PI20716), “Ladak” (PI26927, PI30433), “Orenburg” (PI23625, PI28071), “Semipalatinsk” (PI24455, PI28070), and “Turkestan” (PI469, PI25277-79, PI25805, PI25807). Naturalized populations such as “Grimm” (PI230223, PI452172) and “Ontario Variegated” were widely distributed in the U.S. and Canada.100 At least three registered alfalfa cultivars were selected directly from PI lines in the alfalfa germplasm collection of the U.S. Department of Agriculture (USDA): “Desert” from PI279958,96 “Mesa-Sirsa” from PI235736,104 and “Teton” from a cross between PI20711 and PI24455.38 In 1850, Spanish alfalfa types were introduced into southwestern U.S. from South America. Three winter hardy germplasm sources from Europe and Russia were brought into the upper midwestern U.S. and eastern Canada between 1858 and 1910. Intermediate winter-hardy germplasm from a broad area in the Near East (southern Russia, Iran, Afghanistan, and Turkey) was introduced between 1898 and 1925 and from France in 1947. Nonwinter hardy types were introduced from Peru (1899), India (1913, 1956), and Africa (1924). Essentially all germplasm used in the development of North American cultivars can be traced to these sources of alfalfa.3,5 The U.S. National Plant Germplasm System (NPGS) stores data on individual PIs and permits queries for information and seed from interested scientists. The NPGS alfalfa collection is maintained by the USDA-Agricultural Research Service (ARS), Western Regional Plant Introduction Station, Washington State University, Pullman, Washington. The U.S. Alfalfa Crop Germplasm Committee develops descriptor lists, defines descriptors, and sets priorities for germplasm collections and research on PI material. Plant explorations initiated by the committee have covered major alfalfa distribution centers worldwide. The goal has been to collect native germplasm before local ecotypes are lost or contaminated with introduced germplasm. Beginning in 1983, a program was established to evaluate PI material for resistance to nine insects.4 © 1999 by CRC Press LLC

IV. A. NATURE

OF

INSECT RESISTANCE

RESISTANCE

Trichomes vary widely in structure and function and in distribution over plant parts.133 They are important in plant defense against phytophagous insects. In numerous studies, trichome density is negatively correlated with insect feeding, ovipositional responses, and larval nutrition.70 Some insect resistance in alfalfa is associated with trichomes. Trichomes in Medicago consist of two types: glandular hairs (erect and procumbent) and simple hairs. The stalk of erect trichomes is composed of one to six cells including an enlarged basal cell arising from the epidermis. Gland heads contain numerous cells arranged in distinct tiers. The procumbent trichomes are generally composed of a basal cell embedded in the epidermis, a single stalk cell, and a gland head of four cells in two tiers. The small procumbent glands are typically the only kind appearing on diploid and tetraploid alfalfa.56 The erect capitate glands produce copious secretions apparently on an intermittent basis during the life of the shoot.57 These secretions are probably produced by secretory cell plastids. Procumbent glands produce only a small amount of secretion, which tends to become dry and hard. The procumbent glands and their secretions are not associated with insect resistance. The compounds found in the exudate samples of M. scutellata and M. sativa ssp. sativa (PI346919, M. glutinosa Bieb. in some literature) are straight chain esters (length C17 and greater), straight chain alkanes (C30 and greater), straight chain aldehydes (length C12 and greater), and a straight chain acid (C17). These compounds are large molecules, are nonvolatile, and contribute to the viscous nature of the exudate. Young weevil larvae and leafhopper nymphs are immobilized (Figure 1) by the sticky secretion. No toxic compounds were found in the sticky secretion.132 Erect glandular hairs did not affect the digestibility or crude protein content of alfalfa forage in quality studies.66 Likewise, they did not affect concentrations of neutral or acid detergent fibers, hemicellulose, lignin, or cellulose of leaves or stems within the species and hybrids tested.67 In addition, the presence of erect glandular hairs did not negatively affect forage preference by sheep.68 Simple hairs observed on Medicago species are unicellular and unbranched. Simple hairs have been implicated in resistance of several plant species to insects.70 The hairs apparently provide resistance by interfering with feeding and oviposition. Although very few anatomical features of alfalfa plants have been suspected as causes of resistance to aphids, Manglitz and Kehr73 suspected that a heavy covering of simple trichomes on the seedling stems of PI406295 and PI406296 was responsible for low levels of spotted alfalfa aphid resistance in these lines. Subsequent studies comparing seedlings of this accession with alfalfas having an equal covering of trichomes indicated that the trichomes were not responsible.18 However, glandular trichomes could impart aphid resistance.106 Chemical factors are the primary cause of resistance to insects in several crops. Lignin is an insoluble amorphous micromolecule that encrusts and strengthens fragile polysaccharide constituents of cell walls. This material prevents fiber digestion by

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FIGURE 1 Scanning electron micrograph of a fresh specimen of a first-instar larva of the alfalfa weevil, Hypera postica, after immobilization on a stem of M. scutellata × 100.

livestock through protection of cellulose from enzyme activity, and prevention of swelling of the fiber to a condition suitable for penetration by microbial polysaccharidases.42 Lignin increases the resistance of cell walls to pressure and protects the cellulose fibrils from becoming creased. Lignin has been implicated in resistance of alfalfa to insects.10,11,127 Spotted alfalfa aphids have been observed on resistant cultivars, but studies indicated that the aphids on resistant plants were starving to death and not being killed by a toxin.52,53,76 In one study, aphids appeared unable to ingest plant sap after their stylets reached the phloem on resistant plants.89 Plant chemicals that stimulate and deter T. trifolii feeding have been identified but an association with plant resistance has not been demonstrated.54 The plant chemical saponin may play a role in pea aphid resistance.58 However, saponins, particularly medicagenic acid, were not considered responsible for A. pisum resistance in two studies.8,9 Investigations of the relationship between aphid resistance and sugar and amino acid concentrations of phloem sap indicate that resistance is not a simple nutritional effect but that the amino acid balance may contribute to resistance exhibited by some cultivars.35,37,99 Investigation of epicuticular lipids in relation to T. trifolii feeding indicates that these compounds may play a role in the resistance of one plant line but not in another.7 Miles and Oertli82 speculate that plants respond to wounding, including insect feeding, then mobilize and oxidize phenolic compounds to deter insects. These researchers

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demonstrated reduced reproductive rates by spotted and blue alfalfa aphids when the stems on which they were feeding were infiltrated with ascorbate or glutathione.

B. BREEDING TECHNIQUES Alfalfa genetic behavior and breeding procedures appropriate for developing alfalfa cultivars were described by Rumbaugh et al.101 A wide range of intra- and interpopulation breeding methods have been used but phenotypic recurrent selection in random mating populations has been especially successful in developing resistance to insects.110 For insect resistance breeding, it is necessary to have information on genetic variation in the insect, host reaction to feeding, and the extent to which environmental conditions affect plant resistance and insect behavior. Success of a breeding program also depends on effective screening methods. Access to germplasm resources is essential because large numbers of plants must be included in the breeding program to minimize inbreeding and to conserve genetic variability for other desirable traits. Standard tests for evaluating resistance in alfalfa to important insect pests were published by the North American Alfalfa Improvement Conference.1 The entomological techniques for development of insect-resistant cultivars include mass screening of plant populations to identify resistance categories (antibiosis, nonpreference, tolerance).95 Painter’s nonpreference95 has been replaced with antixenosis.55,72 The final selection of individual plants is based on performance under varied infestation pressures of the test insect and on the level and type of resistance desired. The progeny are tested to determine if the level of resistance is adequate. Strain crosses may be useful in developing multiple pest-resistant cultivars and populations.34 Desirable genes identified in conserved germplasm during evaluation may also be transferred one at a time into selected clones by new techniques.130 Erect glandular hairs on Medicago species have been implicated in resistance to insects that attack alfalfa. The hairs have been transferred from tiny wild diploid plants to tetraploid hay-type alfalfa via 2N gametes.115

C. SPECIFIC INSECTS 1. Potato Leafhopper The potato leafhopper causes extensive losses to the second and successive crops of alfalfa in much of the midwestern and eastern U.S. Both adults and nymphs pierce leaves and stems and ingest plant juices. Injury is characterized by a yellowing, reddening, or purpling of leaves, stunting of plants, and reduced forage quality and yields. Potato leafhoppers overwinter in the U.S. Gulf States and migrate northward each year in spring with the aid of warm air currents. Females deposit eggs in small stems and leaf veins of alfalfa. Eggs hatch in 6 to 9 days into whitish nymphs that soon turn yellowish green. There are several generations per year, with a 3-week interval in the development of adults from eggs.74 Breeding programs first emphasized resistance to yellowing in the field as the resistance criterion. As a result, resistance to yellowing was observed among plants and cultivars.110 In general, M. sativa was more susceptible than M. falcata L.

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(= M. sativa ssp. falcata). Research also discovered that cultivars with falcata germplasm were useful in selecting plants for resistance to yellowing. Recurrent phenotypic selection for resistance to yellowing was effective in the laboratory111,139,140 and in the field.29 The first cultivar (“Cherokee”) developed for resistance to yellowing resulted from seven cycles of selection in a broad-based population.28 It has greatly improved resistance to yellowing compared to that of the parent population. At present, many cultivars with leafhopper resistance are commercially available. Indeed, over 70 cultivars and 19 germplasms were available in 1983.83 This resistance results mainly from plant tolerance. Antixenosis caused by simple hairs was noted in “Hairy Arabian”131 and in “Hairy Peruvian” (“San Pedro”).41 An inheritance study involving clones from “Arc” and B16 (a synthetic involving two “Hairy Peruvian” PIs, 247789 and 247790) found that stem and leaf pubescence could be increased in populations derived from B16.31 Significant correlations for all crosses and clones were found between stem pubescence and both feeding damage and nymphal populations. Several studies identified antibiosis in alfalfa,32,88,103 but highly resistant germplasm lines have not been developed. Entrapment in the sticky exudate of glandular hairs caused almost complete mortality of first-instar nymphs on several annual Medicago species with glandular hairs.105 By contrast, entrapment of adults did not occur on perennial Medicago species, yet antibiosis or strong antixenosis (100% mortality) was detected.10 If the exudate functioned in resistance to adults, it probably restricted feeding or acted as a tactile, olfactory, or gustatory repellent. Glandular hairs contributed to resistance because oviposition on glandular-haired PI346919 clones decreased as hair density increased.10 In a field study, Danielson et al.24 found that potato leafhoppers were less abundant on glandular-haired M. glandulosa David and M. glutinosa than on alfalfa cultivars. Less damage was observed on M. glandulosa than on any other entry. Clones from M. prostrata (UAG 1862) and M. glutinosa (PI346919) were antibiotic to adult leafhoppers and poor substrates for oviposition. Moreover, Elden and McCaslin33 reported that stem glandular hair density was associated with leafhopper resistance and suggested that glandular hairs are associated with a toxic or repellent compound. Brewer et al.11 attributed antixenosis to dense glandular hairs on stems and leaves and noted that stems were tough. Glandular-haired cultivars with a high level of resistance to the potato leafhopper are now available to growers. Genes for resistance were provided by glandular-haired germplasm, including PI346919.108,112,113,125 Commercial breeding programs focused on combining high levels of potato leafhopper resistance with high yield potential, winter hardiness, and multiple disease resistance. Several years of backcrossing to alfalfa and field and greenhouse screening for improved vigor, disease, and potato leafhopper resistance led to the release of the cultivars.94 2. Aphids The pea, the spotted, and the blue alfalfa aphids are the three most important aphid pests of alfalfa. The geographic distribution of the first two aphids appears to coincide with the current distribution of alfalfa in the temperate regions of the world. The

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pea aphid, a native of Europe and a legume specialist, was first observed attacking alfalfa in the U.S. in the late 1800s. The spotted and the blue aphids, both from Asia, were not recognized pests of alfalfa until they appeared in the U.S. southwest, the former in the 1950s and the latter species in the 1970s. The blue aphid continues to spread in the U.S. (Maryland)62 and South America (Argentina138 and Brazil93). The host range of the blue aphid includes several leguminous species, while the spotted alfalfa aphid is largely restricted to the genus Medicago.74 Cultivar development for aphid resistance was greatly encouraged by the fortuitous discovery of the spotted alfalfa aphid resistance in the cultivar “Lahonton” in the 1950s. What followed were other cultivars with spotted alfalfa aphid and pea aphid resistance, and eventually blue alfalfa aphid resistance. It has been possible to develop resistance to two or three species, although resistance to each species appears to be inherited independently. During screening for aphid resistance in PIs, it was apparent that resistance occurs in low levels in plant material from around the world.71 There seems to be a relationship between pea aphid and blue alfalfa aphid resistance, based perhaps on close genetic relationships or pleiotropy.102 The sources of resistance in germplasm releases with resistance to one or more of the aphid species are varied (Table 1). Recently registered cultivars are summarized in Table 2. The influence of the three aphid species on cultivar development and use is demonstrated by a situation in Australia where numerous resistant cultivars have replaced “Hunter River,” a widely grown cultivar that proved to be susceptible to all of the aphids after their immigration to Australia.39 All three mechanisms of resistance have been implicated from the first discovery of spotted alfalfa aphid resistance. Antibiosis and antixenosis have received the greatest amount of attention but recent studies have revealed the possibility of selecting for plant tolerance that is heritable. In these studies, tolerant plants were equivalent to susceptible plants in the numbers of aphids supported but equivalent to resistant plants in regard to the amount of aphid-induced damage. Thus, tolerance was calculated by a damage index that considered chlorosis, stem growth, and number of leaves expressed as:

Damage index =

stem length + number of trifoliolates % leaves with clorosis

Heritability of the tolerance characters was estimated at 25% in selfed progeny of the selected tolerant plants and 20% in polycross progeny. Reduction in heritability of tolerance in these progeny suggests that dominant effects may control expression of this trait.6,46,47 Aphid resistance is heritable and relatively stable but its expression can be temporarily altered by factors such as temperature. For example, “CUF-101” growing under 20°C was highly resistant to the blue alfalfa aphid but resistance was lost at 15°C.128 Frequent monitoring of blue alfalfa aphid populations for increases above the economic threshold is necessary during unusually cool springs in California.129 Low laboratory temperature (16°C) was partially responsible for a reduction in

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TABLE 1 Germplasm Releases (1986–present) with Resistance to Blue Alfalfa Aphid, Pea Aphid, and Spotted Alfalfa Aphid Germplasm

Aphid

OK 51, PI555451 OK 85, PI555452 UC 176, PI552451 UC 195, PI552543 UC 196, PI552544 UC 222, PI552545 UC 263, PI552548 P-3, PI525455 EPA-4, PI508085 KS 189 KS 204, PI520597 KS 208, PI527336 KS 207, PI522240 KS 153 BA3 P4, PI536014 KS 219, PI555667

BAA, BAA, BAA BAA BAA BAA BAA SAA SAA BAA, BAA, BAA, BAA, BAA, BAA,

SAA SAA

KS KS KS KS KS

BAA, BAA, BAA, BAA, BAA,

PA, PA, PA, PA, PA,

220, 221, 222, 223, 224,

PI561845 PI564166 PI564167 PI568157 PI572543

PA, SAA PA, SAA PA, SAA PA, SAA SAA PA, SAA SAA SAA SAA SAA SAA

Source

Ref.

WL 314 Various cultivars “UC Cargo,” “CUF 101” 4 germplasm pools UC Cargo, CUF 101 Wide based germplasm Wide based germplasm Various cultivars EUAN5 x EUPH5 “Sirsa 9” NC 83-2 Various cultivars Various cultivars KS 70 GrahamM3P3AN3 X RileyM4P4AN2 NC 83-2 BIC “Anchor” KS 63 KS 186

15 16 63 64 63 65 65 77 78 116 119 117 114 118 122 120 121 123 124 125

Note: BAA = blue alfalfa aphid; PA = pea aphid; SAA = spotted alfalfa aphid.

resistance to spotted alfalfa aphid in Australia,44 which may explain the observed reduction in field resistance to the spotted alfalfa aphid in the U.S. during 1981. Although a number of resistance breaking biotypes of the spotted alfalfa aphid have been identified by their reaction to the parent clones of “Moapa,” 90 this is not a problem with current resistant cultivars. Presumably other sources of resistance are more difficult for the aphids to overcome. A recent report from Oklahoma documents a new biotype (BAOK 90) of the blue alfalfa aphid which is much more virulent on resistant cultivars than the previously occurring biotype.151 The fact that aphid populations contain a great deal of genetic variation is illustrated by an Australian study in which 12 collections each of blue alfalfa aphid and pea aphid were studied on three cultivars. It was speculated that the 12 blue alfalfa aphid collections were composed of 3 to 7 clones and that the pea aphid collections were composed of 3 to 4 clones.75 These authors suggest that because there is variation in the way plants respond to aphids, cultivars with potential resistance must be exposed to a large number of aphid clones to confirm the presence of useful aphid resistance.

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TABLE 2 Cultivars Registered (1984–present) with Resistance to Blue Alfalfa Aphid, Pea Aphid, and Spotted Alfalfa Aphid Cultivar OK 49, PI547906 WL 525 HQ, PI584332 WL 457, PI550724 “Alfanafa” “Grasslands Oranga” 5444, PI509535 WL 322 HQ, PI549097 WL 225, PI550722 WL 317, PI550723 WL 323, PI586638 “Malone,” PI522242 “Wilson,” PI522243 “Sabre,” PI584992 “Majestic,” PI584991 “Victory,” PI584990 5423 5331, PI522244 555, PI522245 5364, PI533625 5262, PI533624 5311, PI549105 5888, PI549108 5246, PI561715

Aphid

Country

Ref.

SAA BAA, PA, SAA BAA, PA, SAA PA BAA, SAA SAA PA, SAA PA, SAA PA, SAA PA, SAA SAA PA, SAA PA BAA, PA, SAA BAA, PA, SAA PA, SAA PA, SAA PA, SAA PA, SAA PA, SAA PA BAA, PA PA

U.S. U.S. U.S. Australia New Zealand U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S. U.S.

14 20 19 27 30 148 45 60 59 61 79 80 135 136 137 147 142 146 144 145 143 150 149

Note: BAA = blue alfalfa aphid; PA = pea aphid; SAA = spotted alfalfa aphid.

3. Alfalfa Weevil The alfalfa weevil is an Old World insect that was accidentally introduced into the U.S. on three separate occasions (Salt Lake City, Utah, 1904; Yuma, Arizona, 1939; and Baltimore, Maryland, 1951). It now occurs throughout North America. The three weevil introductions are nearly indistinguishable in morphology, cytogenetics, and allozyme patterns and are considered to be strains (western, Egyptian, eastern) of the same species. A distinct difference is the endosymbiotic rickettsia Wolbachia postica which is found in the western strain but not in the Egyptian and eastern strains. While the Egyptian and eastern strains are reproductively compatible, crosses between males of the western and females of the Egyptian or eastern weevils produce infertile eggs.43 The principal damage to alfalfa caused by the alfalfa weevil is the feeding of larvae on interveinal tissue, sometimes leaving only the leaf veins. Heavily infested

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fields may have a whitish appearance. In most areas this beetle produces one generation per year and overwinters as an adult. The greatest abundance of the weevil and the greatest crop damage occur during the first growth of alfalfa in the spring.74 Differences in weevil damage occur among experimental germplasm lines and commercial cultivars. Most progress in breeding has resulted from selection in the field. The first weevil-resistant cultivar (“Team”) was released in 1969.5 Its parentage traces to Kansas and Nebraska synthetics and “Atlantic,” “Dupuits,” “Narragansett,” and “Rhizoma” cultivars. “Arc” and “Liberty” released in 197425 and 1977, respectively, were developed from the same germplasm as “Team” by phenotypic recurrent selection. “Weevilchek,” released in 1971, was derived from M. sativa ssp. falcata and Indian breeding lines. At present, several resistant cultivars are available to growers. This resistance is mostly attributed to plant tolerance expressed as heavy terminals and axillary branching. Well-developed axillary buds continue to grow after the stem terminals have been destroyed by larval feeding. This level of resistance has been effective in integrated control programs that combine cultural, chemical, and biological methods.152 Resistance to oviposition and feeding would greatly increase the value of resistant cultivars. Stem morphology, particularly small stem diameter, has been associated with resistance to weevil oviposition in some M. sativa populations and Medicago species but resistant cultivars have not been developed.12,17,26,92,134 Stem size was not related to oviposition resistance in annual species.49,134 Resistance to weevil feeding was noted in glandular-haired annual species by Shade et al.107 and Johnson et al.,48,50,51 but inability to hybridize annual species with hay-type M. sativa precluded the use of this high level of resistance in cultivar development. The potential of glandular hairs as a resistance mechanism to young alfalfa weevil larvae and other small insects and mites offers promise. Exudate from glandular hairs is gluelike and immobilizes small insects (Figure 1). No toxic materials were found in the exudate of M. scutellata L. Mill.132 Plants selected from glandularhaired diploid species, M. prostrata and M. glandulosa, were highly resistant to alfalfa weevil larvae feeding,23 development,21 and adult oviposition.22 Perennial glandularhaired germplasm that can be crossed with hay-type alfalfa cultivars has been released to breeders for use in the development of insect-resistant cultivars.108,112,114,125 4. Other Insects The threecornered alfalfa hopper, Spissistilus festinus (Say), became a more serious pest on alfalfa in Louisiana when increases in soybean acreage led to increases in hopper populations. This membracid causes significant changes in alfalfa quality and dry-matter yield during the sixth cutting period.85,141 Additional damage is caused by its association with Fusarium crown rot.87 In greenhouse studies, there were clear differences in the developmental rate of the hopper on six alfalfa cultivars.84 Under field conditions there were significant differences among the cultivars for insect numbers and plant damage.86 The alfalfa seed chalcid, Bruchophagus roddi (Gussakovsky), occurs wherever alfalfa seed is produced and has been responsible for reductions in seed yields approaching 85%.110 Low levels of alfalfa resistance exist, but the resistance is

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inadequate in current cultivars. Nielson and Schonhorst91 reported in 1967 that “Lahonton” and several PIs were good sources of resistance. “Lahonton” and other cultivars of Turkestan origin were less susceptible to field infestations than “Hunter River” in a field study in Australia.40 In a field nursery in Iowa, tightly coiled pods were much less infested with seed chalcids than were straight or lightly coiled pods.109 In the most recent study, annual Medicago species with the thickest pod walls experienced the lowest infestation levels compared with a variety of cultivars and species.127 This suggests that breeding for increased pod-wall lignification may reduce seed losses by B. roddi. Resistance to the Meadow spittle bug and clover root curculio, Sitona hispidulus (F.), have been investigated.110 One cultivar, “Culver,” has been released with resistance to the meadow spittle bug, but there have been no cultivar releases with resistance to the clover root curculio. However, a recent report indicates <10% Sitona damage on tap roots of PI183060 and PI183263 compared with 20 to 25% damage for most entries.13

V.

CONCLUSIONS

The development and use of insect-resistant alfalfa cultivars for the control of pest insects is a well-established practice worldwide. The conservation and use of alfalfa germplasm has been vital to the development of these cultivars. A number of cultivars have been developed with resistance to one or more aphid species, including the spotted alfalfa, pea, and blue alfalfa aphids. While resistance breaking biotypes of aphids have been reported, they have not become a serious problem because more stable forms of plant resistance have been located and utilized. In addition, cultivars have been developed with resistance to the potato leafhopper, the alfalfa weevil, and the meadow spittle bug. The development of glandular-haired cultivars with high levels of resistance to the potato leafhopper is a classic example of the use of exotic and conserved germplasm for improving pest management practices for alfalfa production. Glandular-haired germplasm may prove useful in the development of alfalfa resistance to other insect pests.

REFERENCES 1. Anonymous, Standard tests to characterize alfalfa cultivars, North America Alfalfa Improv. Conf., Beltsville, 1996. 2. App, B. A. and G. R. Manglitz, Insects and mites, In C. H. Hanson, Ed., Alfalfa Science and Technology, ASA, CSSA, & SSSA, Madison, 1972, 527. 3. Barnes, D. K., E. T. Bingham, R. P. Murphy, O. J. Hunt, D. F. Beard, W. H. Skrdla, and L. R. Teuber, Alfalfa germplasm in the United States: genetic vulnerability, use, improvement, and maintenance, USDA-ARS Tech. Bull. 1571, U.S. Department of Agriculture, Agricultural Research Service, Hyattsville, 1977. 4. Barnes, D. K., G. P. Goplen, and J. E. Baylor, Highlights in the USA and Canada, In: A. A. Hanson, D. K. Barnes, and R. R. Hill, Eds., Alfalfa and Alfalfa Improvement, Agron. Monogr. 29, ASA, CSSA, & SSSA, Madison, 1988, 1.

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5. Barnes, D. K., C. H. Hanson, R. H. Ratcliffe, T. H. Busbice, J. A. Schillinger, G. R. Buss, W. V. Campbell, R. W. Hemken, and C. C Blickenstaff, The development and performance of Team alfalfa: A multiple pest resistance alfalfa with moderate resistance to the alfalfa weevil, USDA-ARS 34-115, U.S. Government Printing Office, Washington, D.C., 1970. 6. Berberet, R. C., R. W. McNew, J. W. Dillwith, and J. L. Caddel, Within-plant patterns of Therioaphis maculata on resistant, tolerant and susceptible plants, Environ. Entomol., 20:551-555, 1991. 7. Bergeman, D. K., J. W. Dillwith, J. W. Zarrabi, A. A. Caddel, and R. C. Berberet, Epicuticular lipids of alfalfa relative to its susceptibility to spotted alfalfa aphids (Homoptera: Aphididae), Environ. Entomol., 20:781-785, 1991. 8. Bournoville, R., Relationship between the pea aphid and the saponin content of lucerne cultivars, Fourrages, 136:547-554, 1993. 9. Bournoville, R., Net reproductive rate of the pea aphid (Acyrthosiphon pisum Harris) and medicagenic acid content of lucerne, Agronomie, 16:89-94, 1996. 10. Brewer, G. J., E. K. Horber, and E. L. Sorensen, Potato leafhopper (Homoptera: Cicadellidae) antixenosis and antibiosis in Medicago species, J. Econ. Entomol., 79:421-425, 1986. 11. Brewer, G. J., E. L. Sorensen, E. Horber, and G. L. Kreitner, Stem anatomy and potato leafhopper (Homoptera: Cicadellidae) resistance, J. Econ. Entomol., 79:1249-1253, 1986. 12. Busbice, T. H., W. V. Campbell, J. O. Rawlings, D. K. Barnes, R. H. Ratcliffe, and C. H. Hanson, Developing alfalfa resistance to alfalfa weevil oviposition, Crop Sci., 8:762-767, 1968. 13. Byers, R. A., W. A. Kendall, R. N. Peaden, and D. W. Viands, Field and laboratory selection of plant introductions for resistance to the clover root curculio (Coleoptera: Curculionidae), J. Econ. Entomol., 89:1033-1039, 1996. 14. Caddel, J. L., R. C. Berberet, A. A. Zarrabi, and K. T. Shelton, Registration of ‘OK 49’ alfalfa, Crop Sci., 32:280, 1992. 15. Caddel, J. L., R. C. Berberet, A. A. Zarrabi, and K. T. Shelton, Registration of OK 51 germplasm, Crop Sci., 32:839, 1992. 16. Caddel, J. L., R. C. Berberet, A. A. Zarrabi, and K. T. Shelton, Registration of OK 85 germplasm, Crop Sci., 32:839-840, 1992. 17. Campbell, W. V. and J. W. Dudley, Differences among Medicago species in resistance to oviposition by the alfalfa weevil, J. Econ. Entomol., 58:245-248, 1965. 18. Carter, M. R., G. R. Manglitz, and E. L. Sorensen, Resistance to the spotted alfalfa aphid (Homoptera: Aphididae) in simple-haired alfalfa plant introductions, J. Econ. Entomol., 81:1760-1764, 1988. 19. Cluff, G. J., F. L. Bedard, and M. A. Peterson, Registration of ‘WL 457’ alfalfa, Crop Sci., 31:1700-1701, 1991. 20. Cluff, G. J., F. L. Bedard, D. A. Schnebbe, and M. A. Peterson, Registration of ‘WL 525 HQ’ alfalfa, Crop Sci., 35:1501, 1995. 21. Danielson, S. D, G. R. Manglitz, and E. L. Sorensen, Development of alfalfa weevil larvae when reared on perennial glandular-haired Medicago species in the greenhouse, Environ. Entomol., 15:396-398, 1986. 22. Danielson, S. D, G. R. Manglitz, and E. L. Sorensen, Resistance of perennial glandular-haired Medicago species to oviposition by alfalfa weevils (Coleoptera: Curculionidae), Environ. Entomol., 16:195-197, 1987.

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118. Sorensen, E. L., D. L. Stuteville, and E. K. Horber, Registration of KS153BA3P4 alfalfa germplasm, Crop Sci., 30:750, 1990. 119. Sorensen, E. L., D. L. Stuteville, E. K. Horber, and O. J. Dickerson, Registration of KS204 alfalfa germplasm with resistance to five diseases, three insects, and the stem nematode, Crop Sci., 29:239, 1989. 120. Sorensen, E. L., D. L. Stuteville, E. K. Horber, R. N. Peaden, and D. Z. Skinner, Registration of KS220 multiple pest-resistant alfalfa germplasm, Crop Sci., 33:218, 1993. 121. Sorensen, E. L., D. L. Stuteville, E. K. Horber, R. N. Peaden, and D. Z Skinner, Registration of KS221 multiple pest-resistant alfalfa germplasm, Crop Sci., 33:886, 1993. 122. Sorensen, E. L., D. L. Stuteville, E. K. Horber, and D. Z. Skinner, Registration of KS219 alfalfa germplasm with resistance to eight pests, Crop Sci., 32:502-503, 1992. 123. Sorensen, E. L., D. L. Stuteville, E. K. Horber, and D. Z. Skinner, Registration of KS222 multiple pest-resistant alfalfa germplasm derived from ‘Anchor’ alfalfa, Crop Sci., 33:886-887, 1993. 124. Sorensen, E. L., D. L. Stuteville, E. K. Horber, and D. Z. Skinner, Registration of KS223 alfalfa germplasm with resistance to four diseases and three insects, Crop Sci., 34:312, 1994. 125. Sorensen, E. L., D. L. Stuteville, E. K. Horber, and D. Z. Skinner, Registration of KS224 glandular-haired alfalfa germplasm with multiple pest resistance, Crop Sci., 34:544, 1994. 126. Sorensen, E. L., M. C. Wilson, and G. R. Manglitz, Breeding for insect resistance, In: C. H. Hanson, Ed., Alfalfa Science and Technology, ASA, CSSA, & SSSA, Madison, 1972, 371. 127. Springer, T. L., S. D. Kindler, and E. L. Sorensen, Comparison of pod-wall characteristics with seed damage and resistance to the alfalfa seed chalcid (Hymenoptera: Eurytomidae) in Medicago species, Environ. Entomol., 19:1614-1617, 1990. 128. Summers, C. G., Cultivar and temperature influence on development, survival, and fecundity in four successive generations of Acyrthosiphon kondoi (Homoptera: Aphididae), J. Econ. Entomol., 81:515-521, 1988. 129. Summers, C. G. and A. S. Newton, Low temperature decreases CUF 101 alfalfa resistance to blue alfalfa aphid, Calif. Agric., 41:11-12, 1987. 130. Tabe, L. M., T. Wadly-Richardson, and T. J. V. Higgins, A biochemical approach to improving the nutritive value of alfalfa, J. Animal Sci., 73:2752, 1995. 131. Taylor, N. L., Pubescence inheritance and leafhopper resistance relationships in alfalfa, Agron. J., 48:78-81, 1956. 132. Triebe, D. C., C. E. Meloan, and E. L. Sorensen, The chemical identification of the glandular hair exudate from Medicago scutelata, In: Rep. 27th Alfalfa Improv. Conf., USDA-ARS, ARM-NC-19, U.S. Department of Agriculture, Peoria, 1981, 52. 133. Uphof, J. C. Th., Plant hairs, In: W. Zimmermann and P. C. Ozenda, Eds., Encyclopedia of Plant Anatomy, Vol. 4, Part 5, Gebruder Borntraeger, Berlin, 1962, 1. 134. VanDenburgh, R. S., B. L. Norwood, C. C. Blickenstaff, and C. H. Hanson, Factors affecting resistance of alfalfa clones to adult feeding and oviposition of the alfalfa weevil in the laboratory, J. Econ. Entomol., 59:1193-1198, 1966. 135. Viands, D. R., J. L. Hansen, and C. C. Lowe, Registration of ‘Sabre’ alfalfa, Crop Sci., 35:1502, 1995. 136. Viands, D. R., J. L. Hansen, and C. C. Lowe, Registration of ‘Majestic’ alfalfa, Crop Sci., 35:1503, 1995.

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137. Viands, D. R., J. L. Hansen, and C. C. Lowe, Registration of ‘Victory’ alfalfa, Crop Sci., 35:1502, 1995. 138. Vincente, H., M. W. de Holgado, and A. Suarez, New alfalfa cultivars evaluated for cutting in the Republic of Argentina, Informativo de Tecnologia Agropecuaria para la Region Semiarida Pampeana, 90:5-7, 1989. 139. Webster, J. A., E. L. Sorensen, and R. H. Painter, Temperature, plant growth stage, and insect-population effects on seedling survival of resistant and susceptible alfalfa infested with potato leafhoppers, J. Econ. Entomol., 61:142-145, 1968. 140. Webster, J. A., E. L. Sorensen, and R. H. Painter, Resistance of alfalfa varieties to the potato leafhopper: seedling survival and field damage after infestation, Crop Sci., 8:15-17, 1968. 141. Wilson, H. K. and S. S. Quisenberry, Impact of feeding by three cornered alfalfa hopper (Homoptera: Membracidae): greenhouse and field study, J. Econ. Entomol., 80:185-189, 1987. 142. Woodward, W. T., G. E. Hoard, D. L. Jessen, D. J. Miller, L. M. Nash, R. Salter, L. D. Satterlee, and M. A. Smith, Registration of ‘5331’ alfalfa, Crop Sci., 29:486, 1989. 143. Woodward, W. T., G. E. Hoard, D. L. Jessen, D. J. Miller, E. F. Poynor, L. D. Satterlee, and M. A. Smith, Registration of ‘5311’ alfalfa, Crop Sci., 33:874, 1993. 144. Woodward, W. T., G. E. Hoard, D. L. Jessen, D. J. Miller, L. D. Satterlee, and M. A. Smith, Registration of ‘5364’ alfalfa, Crop Sci., 30:230, 1990. 145. Woodward, W. T., G. E. Hoard, D. L. Jessen, D. J. Miller, L. D. Satterlee, and M. A. Smith, Registration of ‘5262’ alfalfa, Crop Sci., 30:229, 1990. 146. Woodward, W. T. and J. W. Miller, Registration of ‘555’ alfalfa, Crop Sci., 29:486-487, 1989. 147. Woodward, W. T., J. W. Miller, L. D. Eckman, L. K. Edmunds, G. E. Hoard, B. J. Hartman, L. M. Nash, and E. F. Poynor, Registration of ‘5432’ alfalfa, Crop Sci., 27:1305-1306, 1987. 148. Woodward, W. T., J. W. Miller, L. D. Eckman, L. K. Edmunds, G. E. Hoard, B. J. Hartman, L. M. Nash, and E. F. Poynor, Registration of ‘5444’ alfalfa, Crop Sci., 27:1306, 1987. 149. Woodward, W. T., D. J. Miller, G. E. Hoard, D. L. Jessen, E. F. Poynor, R. Salter, L. D. Satterlee, and M. A. Smith, Registration of ‘5246’ alfalfa, Crop Sci., 33:1103, 1993. 150. Woodward, W. T., L. D. Satterlee, G. E. Hoard, D. L. Jessen, E. F. Poynor, D. J. Miller, and M. A. Smith, Registration ‘5888’ alfalfa, Crop Sci., 33:1102, 1993. 151. Zarrabi, A. A., R. C. Berberet, and J. L. Caddel, New biotype of Acyrthosiphon kondoi (Homoptera: Aphididae) on alfalfa in Oklahoma, J. Econ. Entomol., 88:1461-1465, 1995. 152. Zavaleta, L. R. and W. G. Ruesink, Expected benefits from nonchemical methods of alfalfa weevil control, Am. J. Agric. Econ., 62:801-805, 1980.

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Section III Vegetable Crops

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9

Vegetable Crops: Search for Arthropod Resistance in Genetic Resources James D. Barbour

CONTENTS I. II. III. IV.

Introduction Value of Germplasm is Context Dependent Need for Germplasm Resources Search for Resistant Germplasm A. Apiaceae 1. Celery (Apium spp.) 2. Carrot (Daucus carota L.) B. Asteraceae 1. Lettuce (Lactuca sativa L.) C. Cucurbitaceae 1. Cucumber (Cucumis sativus L.) 2. Cantaloupe (Cucumis melo var. cantalupensis Naudin) D. Solanaceae V. Conclusions References

I.

INTRODUCTION

Vegetable crops for human consumption are grown for the fresh and processing food markets. The edible portion of a vegetable may be mature and immature fruits (e.g., cucumber Cucumis sativus L., tomato Lycopersicon esculentum Miller), flowers (e.g., sprouting broccoli Brassica oleracea var. “italica” Plenck), leaves (e.g., lettuce Lactuca sativa L., celery Apium graveolens L.), stems (e.g., asparagus Asparagus officinalis L.), roots (e.g., carrot Daucus carota L., cassava, Manihot esculenta Crantz), tubers (e.g., potato Solanum tuberosum L.), or bulbs (e.g., onion Alium cepa L.). Vegetables are produced in tropical, subtropical, and temperate zones throughout the world and are important sources of vitamins, minerals, starches, and proteins. Some vegetables are produced on a scale approaching that of the major cereal crops. For example, potato ranks fourth in production (8.5% of world food production)

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behind wheat Triticum aestivum L., rice Oryza sativa L., and maize Zea mays L., and is a dietary staple for people in many parts of the world.21,69 Others, such as cassava and taro, Colocasia esculenta L., are produced on a smaller scale but are major sources of dietary energy for local populations in parts of the developing world. Many vegetables (e.g., asparagus, rhubarb Rheum rhabarbarum L.), though important to those who produce and consume them, are produced on low hectarage and are considered minor crops. Vegetables are attacked by a large number of arthropods,53,55,105 which are usually controlled with insecticides. Over the last 30 years, however, increasing demands for improved quality of food and environment have resulted in market and legislative forces severely restricting the amounts and kinds of pesticides used in agriculture. The nutritional needs of a rapidly expanding world population require that agricultural production continue to increase at the same time. Meeting these needs will require the development of arthropod management methods that fill the void created by limited pesticide use.81,82 Plant resistance is a principal tactic in integrated pest management64,72 and has a long history as a successful pest management tool.64,87 Plant resistance holds promise in helping to meet our future agricultural needs. However, the challenge of incorporating plant resistance into cultivars has not been well documented for vegetables. Smith103 recorded releases of more than 460 cultivars and germplasm and parent lines in the U.S. from 1973 to 1988. Most of these releases (>85%) were in grains,35 namely wheat,32 corn,86 and sorghum Sorghum bicolor (L.) Moench,107 as well as alfalfa Medicago sativa L.83 and cotton Gossypium hirsutum L.84 Only 12 cultivars and 27 germplasm lines (<10%) were vegetable releases. Kennedy55 reported 9 and Stoner105 reported 50 releases of insect-resistant vegetable germplasms from 1966 to 1977 and 1978 to 1991, respectively. Not all germplasm releases, especially those outside of the U.S., are reported in the literature. These accounts, therefore, represent minimum estimates of the releases of insect-resistant vegetable germplasm. They also document the relatively slow rate at which insectresistant germplasm is being incorporated into vegetable crops. A number of factors contribute to the relatively slow adoption of insect-resistant cultivars into vegetable production.98,105 The most important is the extremely low tolerance by industry and consumers for the presence of arthropods and arthropod damage in vegetable crops. This intolerance can result in very low economic thresholds, effectively raising almost any pest utilizing the harvestable portion of a crop to the status of key pest. Thus, even high levels of plant resistance may fail to provide effective protection. The effect of low economic thresholds for insect pests in high value crops such as vegetables is that insecticides and miticides become economical and thus are frequently used. Most vegetable crops also are attacked by multiple arthropod pests. This poses difficulties for breeding programs because the incorporation of multiple pest resistance into a single cultivar is more difficult than the incorporation of single pest resistance. This increases the time and cost of producing resistant cultivars. This chapter attempts to provoke discussion on the value of germplasm in programs searching and/or breeding for resistance to arthropods in vegetable crops. Because of the great diversity of vegetable crops in terms of their number and © 1999 by CRC Press LLC

taxonomic breadth, variation in their agricultural, economical, and nutritional importance, and limitations of space and available information, a comprehensive treatment is not attempted. Instead, the value of arthropod-resistant vegetable germplasm in a cropping system context is discussed followed by a review of arthropod-resistant germplasm for selected vegetable crops.

II.

VALUE OF GERMPLASM IS CONTEXT DEPENDENT

The value of a resistant cultivar is ultimately measured by the benefits that result from its deployment. Whether or not this value is realized and the extent to which it is realized depends on how the resistant germplasm is matched to the needs of the intended production system. Indeed, the expected benefits of deployment constitute the justification for including selection for arthropod resistance in a breeding program. Because resistance results from an interaction between plant, target arthropod, and environment, a large number of factors may need to be considered. These include the number of pests, pest biology, damaging stages of the pest, the portion (and value) of the crop damaged, the cost of alternative control measures, the type and level of resistance, the potential for development of resistance-breaking insect biotypes, and the effects of resistance on natural enemies of the target pest, secondary pests in the target crop, and non-target pests (weeds, diseases, other arthropods) in the target and adjacent crops. These factors are not unique to vegetable crops, but are worth considering here.17,59 Specific resistance mechanisms may be more valuable than others in a given pest-crop system. Consider, for example, the cowpea, Vigna ungiculata (L.), which is produced as a vegetable crop for the immature seeds. Infested or damaged seeds are unacceptable in the U.S. The cowpea curculio, Chalcodermis aeneus Boheman, feeds on cowpea pods and oviposits on seeds developing within the pods. Three types of resistance to C. aeneus are available: (1) antibiosis prolongs development and increases mortality of larvae feeding on the seeds, (2) antixenosis inhibits adult feeding on pods, and (3) a pod factor inhibits penetration of the pod wall and oviposition.9 In this case, antixenosis resistance and the pod factor, which act to reduce cosmetic damage to the seeds, are more important than antibiosis resistance which does not reduce the frequency of infested and damaged seed. The level of insect resistance expressed by cultivars must be appropriate to the cropping system. It is difficult to imagine useful levels of tolerance for insects that feed on the harvested portion of a vegetable crop. However, the ability to produce a second crop when the first has been destroyed may be advantageous for a crop that is not required to meet a restrictive marketing window.56 Different levels of antibiosis or antixenosis resistance may be more or less suited to the needs of the pest-crop system under consideration. In the case of cowpea and the cowpea curculio, the decision concerning the most appropriate type of resistance was not difficult. However, determining the level of resistance befitting this situation may be more challenging. For pests that feed on the harvested portion of a crop or for itinerant pests that attack crops in large numbers from sources outside the crop, even high levels of resistance may provide inadequate protection. Consider tomato in North Carolina © 1999 by CRC Press LLC

(U.S.), which is attacked by tomato fruitworm, Helicoverpa zea (Boddie).58 Corn is the primary acreage host supporting H. zea populations during the spring and early summer months. As corn senesces in mid-summer, large numbers of moths emigrate to blooming crops, including tomato. Tremendous numbers of eggs may be laid in these crops. Most or all of the fruit is commonly destroyed in unprotected plantings.59 Under these conditions, low to moderate levels of antibiosis or antixenosis would contribute little to crop protection. Levels of antixenosis or antibiosis approaching immunity would be required to significantly reduce the need for insecticides. High levels of resistance preventing oviposition or causing high mortality to young larvae before they entered fruits might contribute to a management program by facilitating insecticidal control.58 On the other hand, more flexibility in terms of resistance levels might exist for the production of processing tomatoes or for insects and mites not attacking tomato fruits or flowers. Lower levels of resistance also may be appropriate for pests whose densities are initially low, but whose numbers increase slowly over several generations in a tomato crop.58 Resistance to one pest may be negatively correlated to resistance to other pests attacking a crop. Cucurbitacins are tetracyclic terpenoid compounds that act as powerful feeding stimulants for cucumber beetles, Diabrotica spp.7 While the lack of cucurbitacins in cucumber conditions antixenosis resistance to cucumber beetles, their presence confers antibiosis resistance to twospotted spider mite, Tetranychus urticae Koch.13,19,85 Resistance may also be associated with negative effects on natural enemies of the pest under consideration, or on natural enemies of other pests in a cropping system.2,3,22,44 Under such circumstances, plant breeders must carefully consider the relative importance of the various pests and the contribution of natural enemies to pest control. The durability of resistance is another important consideration. Research and development leading to deployment of an arthropod-resistant cultivar requires a 3- to 15-year commitment of resources.37,64 Development of pest biotypes capable of overcoming resistance within a few years after release can seriously reduce the value and effectiveness of resistant germplasm. A number of factors, e.g., pest biology, resistance mechanism, physiological/behavioral effect of resistance on a pest, and inheritance of resistance, can affect the rate at which selection for resistant biotypes occurs.39,58,113 Antibiosis resistance is often assumed to result in greater selection pressure than antixenosis-based resistance. Yet, antibiosis resistance that simply slows development of a univoltine pest may have no effect on the pest’s fitness.39 The intensity of selection imposed by antixenosis resistance depends on the degree of hostspecificity of a pest, its mobility, and the availability of alternate hosts. Antixenosis resistance may impose considerable selection pressure on pests when alternate hosts are not located within the range of the pest’s mobility. The rate of development of resistance-breaking pest biotypes may be affected by the strategy used to deploy a resistant cultivar.40,41,113 Development of resistant biotypes in vegetable crops is uncommon, but has been reported for pea aphid, Acyrthosiphon pisum (Harris), on pea, Pisum sativum L.;91 bean aphid, Aphis fabae Scopoli, on faba bean, Vicia faba L.;48 and cabbage aphid, Brevicoryne brassicae (L.), on brussels sprouts, B. oleracea var. “gemmifera” Zenker,25 and swede (rutabaga), B. napus var. “napobrassica” (L.) Reichb.67,68 © 1999 by CRC Press LLC

III.

NEED FOR GERMPLASM RESOURCES

In view of the above considerations, it seems clear that plant resistance should be evaluated in the context of the cropping system of interest. However, germplasm that exhibits the type and level of resistance best fitting the anticipated needs of the targeted cropping system is seldom readily available. In fact, the quality of many germplasm collections, with respect to arthropod resistance, is unknown because efficient and reliable screening methods are not available.46 This lack of efficient screening methods is also a hindrance to research programs seeking to improve crop cultivars by evaluating large numbers of germplasm for resistance. The existence of sufficient genetic diversity is a fundamental requirement if programs breeding for resistance and other crop improvements are to meet future crop production needs. For a number of reasons, genetic diversity is at risk. Habitat destruction reduces plant genetic diversity in many parts of the world.34,49,79,80 There also has been a continuous movement of genetic diversity out of farmer fields and into gene banks since the early 1900s.1,34 This trend was exacerbated by the green revolution of the 1960s and 1970s, in which traditional varieties and landraces were replaced by newer high-yielding cultivars that were often inappropriate to local agriculture and required high production inputs to achieve higher yields.5,34,45,46 Fowler34 estimated this decline by comparing the U. S. Department of Agriculture (USDA) inventory of vegetable cultivars being sold commercially in the U.S. in 1903 with inventories of the same cultivars held in the USDA-Agricultural Research Service (ARS), National Seed Storage Laboratory, Fort Collins, Colorado, in 1983. For vegetable crops, 83 to 100% of the cultivars listed in 1903 were not listed in the 1983 inventory. The exception was chives, Allium schoenoprasum L., which had the same cultivar listed in both the 1903 and 1983 inventories. Although Fowler’s34 estimates do not account for new cultivars inventoried since 1983, they do indicate potentially serious losses of genetic diversity in vegetable germplasm over an 80-year period.

IV.

SEARCH FOR RESISTANT GERMPLASM

Theories regarding the evolutionary origin of resistance are central to the question of where to search for useful sources of resistant germplasm.46,47,71 Leppik71 argued that resistance to insects and other pests is most likely to be found in areas where the pest and the plant are sympatric. Specifically, the search for resistance should focus on centers of genetic diversity of cultivated plants. Leppik’s approach assumes the existence of an evolutionary relationship between plant and pest, which over time results in an accumulation of pest-resistant genes in the plant. For example, resistance to spotted alfalfa aphid, Therioaphis maculata (Buckton), in alfalfa seems to be concentrated in accessions from the Turkestan region.71 Although Leppik’s focus was on high levels of single gene resistance to plant pathogens, there are many examples of plant resistance to insects involving plant/pest sympatry (Chapter 4 in this volume).46,47 High levels of resistance to arthropods have been documented from areas where the crop and pest are not sympatric, i.e., allopatric resistance.47 Smith103 recorded

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resistance in 10 crop species to 13 species of arthropods that has apparently evolved in the absence of the insect (Chapter 4 in this volume). A number of mechanisms may explain the existence of allopatric resistance; namely, pleiotropic effects of nonresistance genes, native pest-resistance genes conferring resistance to non-native pests, and genetic drift.47,56 Harris47 argued that allopatric resistance is more likely to be polygenic and, therefore, less likely to be overcome genetically by a pest.50,51 Resistance appears to be “where you find it” and in the absence of a single theory to unite or replace these two contrasting hypotheses, the best approach is to maintain a broad germplasm base in repositories.46 A review of arthropod-resistant germplasm for several major vegetable crop species is provided below. The focus is on sources (landraces, wild species, etc.) of resistance rather than resistance mechanisms or the genetic basis of resistance. A summary of arthropod-resistant accessions and cultivars for the vegetable crops discussed herein is provided in Table 1.

A. APIACEAE 1. Celery (Apium spp.) The genus Apium consists of 15 species that are distributed throughout the world. The center of origin of Apium is thought to be the Mediterranean region.92 All cultivars belong to a single species, A. graveolens, of which celery, A. graveolens var. “dulce” (Miller) Pers., is the most economically important. Although grown as a medicinal crop since 6000 B.P., cultivation of celery as a vegetable crop is thought to have begun in the 1500s in Italy.92 The three horticultural cultivars of celery, A. graveolens var. “dulce” (celery), A. graveolens var. “rapaceum” (Miller) Gaudin (celeriac), and A. graveolens var. “secalinum” Alef. (smallage), result from selection for solid petioles, hollow petioles, and enlarged hypocotyls, respectively.92 All modern celery cultivars apparently derive from the cultivars “White Plume” and “Giant Pascal,” both introduced into the U.S. in 1887.92 Celery is produced largely in the Americas and Europe. Production in the U.S. is more than 711,200 mt annually,73 mostly (67%) in California where yields average 65.02 mt/acre. Production figures are not available for other countries.92 Insect resistance has been found in celery cultivars (primary genepool) and in related Apium species (tertiary genepool). Cultivars of celery, celeriac, and smallage have been evaluated for resistance to the lepidopterans Spodoptera exigua (Hübner) and Trichoplusia ni (Hübner).43,77 No resistance to T. ni was detected, but resistance to S. exigua was detected in cultivars of celeriac. Plant Introduction (PI) 223333 was less suitable than PI357333 as a host for S. exigua as measured by survival, time to pupation, and pupal weight. Resistance has been detected in related Apium species evaluated against the field pests Liriomyza trifolii (Burgess) and S. exigua. Resistance to S. exigua in wild A. prostratum Labill did not appear to be associated with furanocoumarin levels in the host, which are associated with contact dermatitis in humans. Therefore, this resistance might be suitable for transfer to commercial celery cultivars.20 Resistance to L. trifolii was found in accessions of wild A. prostratum, A. chilense Hook and

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TABLE 1 Arthropod Resistance in Cultivars and Accessions of Selected Vegetable Crops and Their Wild Relatives Plant Celeriac

Species

Carrot

Apium graveolens var. “rapaceum” A. chilense A. panul A. prostratum A. nodiflorium Daucus carota

Lettuce Wild lettuce

Lactuca sativa L. virosa

Cucumber

L. saligna L. serriola L. perennis Cucumis sativus

Wild celery

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Arthropod

Accession, line, or cultivar

Ref.

Spodoptera exigua

PI223333

20, 43, 77

Liriomyza trifolii

A73 A160 A230 PI279829 Various cultivars Various cultivars “Berlikum” “Imperial 45634-m” PIVT 280 PIVT 280, various accessions Various accessions PI261653 PI274372 PI279684, PI253299, PI281876 “Hybrid Long Green Pickle,” “Ohio M.R.,” “Taipei No. 1,” “Robin 50,” “Aodai,” PI220860, PI178885, PI163222, PI218036

20, 110 20 20, 110 109 99,100 10-12, 28,29 24 23 27 26, 95 53, 89 62, 63, 88, 89, 112

Lygus spp. Psila rosae Cavariella aegopodii Pemphigus bursarius Nasonovia ribisnigri Myzus persicae Macrosiphum euphorbiae Trichoplusia ni

Tetranychus urticae

8, 14-16, 19, 65

TABLE 1 (continued) Arthropod Resistance in Cultivars and Accessions of Selected Vegetable Crops and Their Wild Relatives Plant Wild cucumber

Cantaloupe

Tomato Wild tomato

Species C. africanus C. anguria C. myriocarpus C. angolensis C. asper C. deteri Cucumis melo

Lycopersicon esculentum Seven species

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Arthropod T. urticae

Accession, line, or cultivar Various accessions

Trialeurodes vaporarorium

Aphis gossypii

Ref. 8, 14-16, 19, 65

66

Diabrotica spp. Acalymma spp. Liromyza sativa Š12 species

PI161375, PI164320, PI371795, PI414723, 90234 (from PI371795), “Kanro makuwa,” “Ginsen makuwa,” “Shiroubi okayama,” AR Hales Best Jumbo, AR5, AR Topmark 91213 (from PI371795) C922-174-B PI282448, PI313970 Ý100 accessions, lines, or cultivars

Š20 species

Š50 accessions

4, 18, 60, 75, 90

61, 76 57 33

Arn, and A. panul (DC.) Reiche.110 Crosses between A. chilense (accession A73) or A. panul (A160) and A. graveolens (“Tall Utah,” A41) express antibiosis resistance to L. trifolii, but have low fertility, making their use in a breeding program difficult. Apium nodiflorum (L.) Lag. (PI279829) exhibited effective antibiosis against L. trifolii while possessing low levels of furanocoumarins, making it a suitable candidate for backcrossing with commercial cultivars for possible insect resistance.109 Hybridization of A. graveolens (A41) with A. prostratum (A230) also has been successful,92 resulting in the expression of antixenosis resistance at a level approaching immunity to L. trifolii. The University of California, Davis maintains a large working collection of A. graveolens cultivars and accessions of several wild species. Approximately 204 accession of A. graveolens (cultivars and landraces) are stored and maintained at the USDA-ARS Northeast Regional Plant Introduction Station (NRPIS), Geneva, New York.42 A directory of Apium germplasm collections is available.108 2. Carrot (Daucus carota L) Carrot, an open pollinated crop originating from the region around Afghanistan,101 spread to the Mediterranean region and was widely cultivated in Europe by the Middle Ages.69 It was introduced to the U.S. (Virginia) in 1607. Carrot is important to worldwide agriculture with a total production of more than 12 mmt annually. Average U.S. production is about 889,000 mt, placing carrots ninth in terms of vegetable crop production.73 Carrot is also an important source of provitamin A carotenes.101 All insect resistance in carrot has been found in cultivars. Resistance has been documented for Lygus spp., the carrot aphid, Cavariella aegopodii (Scop.), and the carrot rust (root) fly, Psila rosae (F). Resistance to Lygus spp. among cultivars and among plants within cultivars results from increased mortality (antibiosis).99 This resistance could be increased through selecting and selfing plants within cultivars.100 Dunn24 documented resistance variation among cultivars against C. aegopodii, with “Berlikum” the least susceptible to attack. Resistance among eight carrot cultivars to P. rosae has been reported.28,29 There is a positive relationship between the concentration of chlorogenic acid in carrot roots and damage by P. rosae, such that fly attacks result in higher concentrations of chlorogenic acid, higher fly infestations, and more damage.10-12 Relatively little carrot germplasm has been collected and preserved in genebanks for distribution. Approximately 710 accessions are stored and maintained at the USDA-ARS North Central Plant Introduction Station, Ames, Iowa.42 Although cultivated carrot crosses readily with wild carrot, D. carota L. ssp. carota, (Queen Anne’s Lace), the extent of genetic variability in wild carrot and other wild Daucus species for insect resistance has not been examined.101

B. ASTERACEAE 1. Lettuce (Lactuca sativa L.) Lettuce, a cool season leafy vegetable, has been cultivated for over 4,500 years.69 The largest producing areas are in the U.S. and Europe with annual hectarages of 91,000 and 80,000, respectively. However, lettuce is produced on all continents with substantial production in Australia, Japan, China, Israel, Mexico, Chile, Argentina, Brazil, © 1999 by CRC Press LLC

and Peru.89 Two-thirds of the U.S. production is in California. Lettuce is the most valuable vegetable crop in the U.K., and ranks second behind tomato in the U.S. Although the nutritional value of lettuce is not high, its consumption places it fourth behind tomato, citrus, and potato in terms of its nutritional contribution to the consumer diet.96 Insect resistance has been found in L. sativa cultivars and in related wild species. The lettuce root aphid, Pemphigas bursarius (L.), is a serious pest that threatens outdoor crisphead lettuce production in the U.K. Resistance to this aphid has been found in several cultivars derived from “Imperial 45634-m.”23 The lettuce leaf aphid, Nasonovia ribisnigri (Mosley), is the most common aphid pest in The Netherlands. Resistance amounting to near immunity to N. ribisnigri was found in accession PIVT 280 of the wild lettuce L. virosa L. and transferred to L. sativa using another wild lettuce, L. serriola L., as a bridge species.27 Resistance to N. ribisnigri is controlled by a single dominant gene (Nr) that also conditions partial resistance to the green peach aphid, Myzus persicae (Sulzer).95 Additionally, resistance to M. persicae26 and Macrosiphum euphorbiae (Thomas)94 has been found in lettuce cultivars.53,89 The cabbage looper, T. ni, is a serious pest of lettuce in the U.S. where as many as 15 insecticide applications per season may be required to control it in Arizona and California production areas.62 Wild relatives of L. sativa appear to hold valuable sources of resistance to this pest. For example, Pesho et al.88 screened 433 plant introductions representing six Lactuca species and found that three species were highly resistant: L. saligna L. (PI261653), L. serriola L. (PI274372), and L. perennis L. (PI279684). Further research indicated that the resistance in PI261653 and PI274372 resulted from reduced oviposition.63 Examination of progenies from crosses between L. sativa and L. serriola (PI274372) indicated that antixenosis resistance was independent of agronomic characters.62 Although this resistance was transferred to L. sativa, it has proven ineffective under field conditions.89 Antibiosis resistance resulting in delayed development of T. ni larvae was also documented in PI261653 and in two accessions (PI253299, PI281876) of L. perennis.112 The U.S. Lactuca collection is stored and maintained by the USDA-ARS Western Regional Plant Introduction Station, Pullman, Washington (Ý1309 accessions).42 The Center for Genetic Resources, Wageningen, The Netherlands (Ý2118 accessions),6 and Horticulture Research International, Wellesbourne, U.K. (Ý1000 accessions),114 maintain large collections in Europe.

C. CUCURBITACEAE The genus Cucumis originated in northeastern Africa (watermelon, Citrullus lanatus [Thunb.] Matsum. and Nakai var. “lanatus”), Arabia and the eastern Mediterranean (melon, Cucumis melo L.), and India (cucumber, Cucumis sativus L.).70 Southern Africa is the secondary site of genetic diversity for C. melo. The C. sativus Asiatic group has centers of diversity in India and China.102,104,106

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1. Cucumber (Cucumis sativus L.) Cucumber was originally cultivated in India and has been in cultivation for over 5,000 years.106 Cucumber was brought to the New World by Columbus in 1494 and is now cultivated in temperate, tropical, and subtropical zones throughout the world.70 World production (13 mmt on over 800,000 ha) ranks fourth behind tomato, cabbage, and onion.106 More than half of the world production is consumed in Asia.106 Arthropods are a major factor limiting production of Cucumis worldwide. Differences in chromosome number between species of the Asian group (2N = 14), including C. sativus, and the African group (2N = 24), including C. melo, have made resistance to arthropods in species of one group generally unavailable to species of the other group. However, an interspecific cross between C. sativus and C. hystrix Chakr. may form a bridge between these groups, allowing reciprocal gene flow.8 This bridge may be important because pests such as twospotted spider mite, T. urticae, and cucumber beetles, Diabrotica spp. and Acalymma spp., attack both C. sativus and C. melo, and other cultivated and wild cucurbit species.53,78 Resistance in C. sativus to T. urticae has been attributed to antibiosis conditioned by the presence of the tetracyclic terpenoid cucurbitacins.7,19,85 Because cucurbitacins act as powerful feeding stimulants for cucumber beetles,13,78 germplasm resistant to T. urticae, but not containing cucurbitacins, could prove valuable. Resistance to T. urticae has been observed in non-bitter (lacking cucurbitacins) cucumber cultivars18 and, conversely, some bitter cucumber lines (containing cucurbitacins) are known to be susceptible to T. urticae.38 Research in The Netherlands indicated that resistance to T. urticae was not correlated with resistance to Diabrotica spp.16,18 Several lines resistant to T. urticae, but lacking cucurbitacins, were produced from crosses of a non-bitter slicing cucumber line (G6) with F3 and F5 selections from crosses of “Hybrid Long Green Pickle” and “Robin 50,” and “Hybrid Long Green Pickle” and “Varamin,” respectively.18 These lines were resistant to T. urticae in greenhouses in The Netherlands, but not in the field in the U.S.19 Kooistra65 and DePonti14-16 screened 800 cucumber cultigens (hybrids, landraces, PIs) and 6 related Cucumis species for resistance to T. urticae and found resistance in 9 C. sativus cultivars and accessions. The resistance, in decreasing order, was in PI220860, “Hybrid Long Green Pickle,” PI178885, “Ohio M.R. 200,” “Taipei no. 1,” “Robin 50,” “Aodai,” PI163222, and PI218036. Resistance also was found in C. africanus Lindly, C. anguria L., and C. myriocarpus Naud. Use of C. hystrix as a bridge species may make this resistance exploitable to C. sativus and other related Cucumis species.8 Resistance to the whitefly, Trialeurodes vaporariorum Westw., a serious pest of cucumber in greenhouse production, has been found in wild C. angolensis Hook, C. asper Cogn., and C. deteri Cogn.66 A screening of more than 1000 C. sativus germplasm lines for resistance to pickleworm, Diaphania nitidalis Stoll, failed to detect useful levels of resistance.30,111

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2. Cantaloupe (Cucumis melo var. cantalupensis Naudin) Accurate data on melon production are difficult to obtain because there are many cultivated types. McCreight et al.76 cited figures for cantaloupe production in Israel (1988) and cantaloupe and honeydew for the U.S. (1984) of 4,500 ha and 318,000 ha, respectively. Production figures from this same source for various melons in Spain (1988) and France (1990) were 62,000 ha and 16,000 ha, respectively. Conserved germplasm is a source of resistance to a number of arthropods attacking melon. Resistance to melon aphid, Aphis gossypii (Glover), has been documented in C. melo PIs 161375, 164320, 371795, and 4147234,60,90 and in C. melo cultivars from Japan, namely “Kanro makuwa,” “Ginsen makuwa,” and “Shiroubi Okayama.” 90 Three breeding lines, AR Hales Best Jumbo, AR5, and AR Topmark, expressing tolerance and antixenosis derived from PI371795 have been released.75 In addition, the breeding line 91213 derived from PI371795 exhibits high levels of resistance to Diabrotica spp. and Acalymma spp.61,76 Resistance to cucumber beetles, D. undecimpunctata howardii Barber, D. balteata LeConte, and A. vittata (F.), was found in a breeding line (C922-174-B) lacking cucurbitacins.85 Because Nugent et al.85 found that bitter and non-bitter cucumber lines were equally susceptible to T. urticae, they endorsed DePonti and Garretsen’s18 hypothesis of no correlation between resistance to spider mite and cucumber beetle. Wild C. melo accessions PI282448 (Africa) and PI313970 (India) were resistant to the leaf miner Liromyza sativae Blanchard.57 Major holdings for Cucumis and Citrullus germplasm are located in genebanks of the U.S. National Plant Germplasm System (approximately 7000 accessions)42 and in Russia, India, and Japan.31

D. SOLANACEAE Tomato is among the world’s most important vegetable crops and is produced on over 2.6 million ha (63 mmt). Major production centers are in Europe, Japan, China, and U.S. Tomato is the second most important U.S. vegetable crop in terms of its monetary value (>$800 million [U.S.] annually), and ranks first in dietetic contribution to total vitamins and minerals.54 Tomato is native to Central and South America and was domesticated in Mexico.54 According to Rick,97 Lycopersicon is comprised of nine species: (1) L. esculentum, (2) L. cheesmanii Riley, (3) L. parviflorum Rick, (4) L. pimpinellifolium (Juls.) Miller, (5) L. chmielewskii (Rick et al.), (6) L. hirsutum Humb. and Bonpl., (7) L.(= Solanum) pennellii (Corr.) D’Arcy, (8) L. peruvianum (L.) Miller, and (9) L. chilense Dunal. Although unilateral incompatibility is common in Lycopersicon, all species except L. peruvianum hybridize readily with L. esculentum.54 Hybridization of L. peruvianum with L. esculentum is possible only with embryo culture techniques.54 Tomato germplasm collections contain a great deal of genetic diversity for arthropod resistance in cultivated and wild species. Farrar and Kennedy33 reported results of studies evaluating nearly 100 L. esculentum germplasm lines. Resistance was documented to at least 12 arthropod species, including Helicoverpa armigera (Hübner), H. zea, Keiferia lycopersicella (Wals.), Liriomyza sativa (Blanchard), L. trifolii (Burgess), two forms of Microsiphum euphorbiae (Thomas), Spodoptera eridania

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(Cramer), Tetranychus cinnabarinus (Boisd.), T. urticae, and Trialeurodes vaporariorum (Westw.). Sixteen sources were resistant to two or more arthropods. Resistance to four arthropods, H. zea, M. euphorbiae, S. exigua, and T. cinnabarinus, was found in three accessions of L. esculentum var. “cerasiforme” (Dunal) A. Gray.36,52,93 Several wild species contain resistance to arthropods33 with levels of resistance higher than in cultivated species. Much research has focused on accessions of L. hirsutum, which occurs in two distinct forms, L. hirsutum f. typicum and L. hirsutum f. glabratum C.H. Muller. High levels of resistance to more than 20 different arthropod species have been reported in accessions of these two species alone. Many accessions contain resistance to multiple pests. For example, PI134417 (H. hirsutum f. glabratum) is resistant to Aphis gossypii Glover, Epitrix hirtipennis, H. armigera, H. zea, K. lycopersicella, Leptinotarsa decemlineata (Say), L. sativa, Manduca sexta L., Myzus persicae (Sulzer), S. exigua, T. cinnabarinus, and T. urticae.33 In addition, resistance has been documented in accessions of L. cheesmanii (2 accessions), L. chilense (1), L. chmielewskii (1), L. pennellii (Š 9), L. peruvianum (16), and L. pimpinellifolium (3).33 The NRPIS (5795 accessions) and the Tomato Genetic Stock Center, University of California, Davis (approximately 2904 accessions), U.S., store and maintain major Lycopersicon germplasm collections (Chapter 12 in this volume).42

V.

CONCLUSIONS

Many instances of the successful use of plant resistance for managing arthropod pests can be cited.64,74,103 These cases indicate both the real and potential value of plant resistance as a tool for managing arthropod pests. Plant breeders, entomologists, and others have invested tremendous amounts of effort and resources toward the development and deployment of vegetable crops resistant to arthropods. Despite the difficulties inherent in incorporating arthropod resistance into vegetable crops, the return on the investment has been promising. However, this promise falls short of that anticipated for plant resistance in vegetable crops.55,98,105 For plant resistance to meet its full potential in vegetable crop production, resistant germplasm incorporated into cultivars must simplify (sensu Kennedy et al.58) the overall management of the crop. Germplasm that is closely matched to the needs of a cropping system is more likely to simplify crop management than germplasm that is less closely matched to these needs. Therefore, teams of multidisciplinary scientists working in vegetable improvement programs are needed to establish and screen the world’s germplasm stocks for useful germplasm. Arthropodresistant germplasm has been found in crop cultivars and in wild and related species of the crop plants. However, the types and levels of resistance best suited to a cropping system may not be available in conserved germplasm. This situation is exacerbated by market and agronomic considerations that further narrow the range of acceptable germplasm for vegetable crop improvement.

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REFERENCES 1. Anonymous, Summary of the Second Crop Advisory Committee Workshop, U.S. Department of Agriculture, Agricultural Research Service, Beltsville, 1988. 2. Barbosa, P., Natural enemies and herbivore-plant interactions: influence of plant allelochemicals and host specificity, In: P. Barbosa and D. K. Letourneau, Eds., Novel Aspects of Insect-Plant Interactions, Wiley, New York, 1988, 201. 3. Bergman, J. M. and W. M. Tingey, Aspects of interaction between plant genotypes and biological control, Bull. Entomol. Soc. Am., 25:275-279, 1979. 4. Bohn, G. W., A. N. Kishaba, and H. H. Toga, Mechanisms of resistance to melon aphid in a muskmelon line, HortSci., 7:281-282, 1972. 5. Brown, A. H., O. H. Frankel, D. R. Marshall, and J. T. Williams, The Use of Plant Genetic Resources, Cambridge University Press, Cambridge, 1988, 423. 6. CGN (Center for Genetic Resources, The Netherlands), on the World Wide Web (http://www/bib/wau.nl/cgn.html), 1997. 7. Chambliss, O. and L. C. M. Jones, Chemical and genetic basis for insect resistance in cucurbits, J. Hort. Sci., 89:394-405, 1966. 8. Chen, J. F., J. E. Staub, and Y. Tashiro, Regeneration of interspecific hybrids between Cucumis sativus L. x C. hystrix Chakr. by direct embryo culture, Cucurbits Genetics Coop. Rep., 19:13-16, 1996. 9. Cuthbert, F. P., Jr., R. L. Fery, and O. L. Chambliss, Breeding for resistance to cowpea curculio in southern peas, HortSci., 9:69-70, 1974. 10. Cole, R. A., Relationship between the concentration of chlorogenic acid in carrot roots and the incidence of carrot fly larval damage, Ann. Appl. Bio., 106:211-217, 1985. 11. Cole, R. A., K. Phelps, J. A. Ellis, and J. A. Hardman, The effects of time of sowing and harvest on carrot biochemistry and the resistance of carrots to carrot fly, Ann. Appl. Biol., 110:135-143, 1987. 12. Cole, R. A., K. Phelps, J. A. Ellis, and S. A. Rollason, Further studies relating chlorogenic acid concentration in carrots to carrot fly damage, Ann. Appl. Biol., 112:13-18, 1988. 13. DaCosta, C. P. and C. M. Jones, Cucumber beetle resistance and mite susceptibility controlled by the bitter gene in Cucumis sativus L., Science, 172:145-1146, 1971. 14. DePonti, O. M. B., Resistance in Cucumis sativus L. to Tetranychus urticae Koch. 3. Search for sources of resistance, Euphytica, 27:167-176, 1978. 15. DePonti, O. M. B., Resistance in Cucumis sativus L. to Tetranychus urticae Koch. 4. The genuineness of the resistance, Euphytica, 27:435-439, 1978. 16. DePonti, O. M. B., Resistance in Cucumis sativus L. to Tetranychus urticae Koch. 5. Raising the resistance level by the exploitation of transgression, Euphytica, 28: 569-577, 1979. 17. DePonti, O. M. B., Plant resistance to insects: a challenge to plant breeders and entomologists, In: J. H. Visser and A. K. Minks, Eds., Proc. 5th Int. Symp. Insect Plant Relationships, Pudoc, Wageningen, 1982, 337. 18. DePonti, O. M. B. and F. Garretsen, Resistance in Cucumis sativus L. to Tetranychus urticae Koch. 7: the inheritance of resistance and bitterness and the relation between these characters, Euphytica, 29:513-523, 1980. 19. DePonti, O. M. B., G. G. Kennedy, and F. Gould, Different resistance of non-bitter cucumbers to Tetranychus urticae in The Netherlands and the USA, Cucurbit Genetics Coop. Rep., No. 6, 27-28, 1983.

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20. Diawara, N. M., J. T. Trumble, C. F. Quiros, and J. G. Millar, Resistance to Spodoptera exigua in Apium prostratum, Entomol. Exp. Appl., 64:125-133, 1992. 21. Douches, D. S. and K. Jastrzebski, Potato (Solanum tuberosum L.), In: G. Kalloo and B. O. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pergamon Press, Oxford, 1993, 605. 22. Duffey, S. S. and K. A. Bloem, Plant defense-herbivore-parasite interactions and biological control, In: M. Kogan, Ed., Ecological Theory and Integrated Pest Management Practice, Wiley, New York, 1986, 135. 23. Dunn, J. A., Varietal resistance of lettuce to attack by lettuce root aphid, Pemphigus bursarius (L.), Ann. Appl. Biol., 76:764-767, 1960. 24. Dunn, J. A., The susceptibility of varieties of carrot to attack by the aphid, Cavariella aegopodii (Scop.), Ann. Appl. Biol., 76:301, 1970. 25. Dunn, J. A. and D. P. H. Kempton, Resistance to attack by Brevicoryne brassicae among plants of brussels sprouts, Ann. Appl. Biol., 72:1-11, 1972. 26. Eenink, E. H. and F. L. Dieleman, Screening Lactuca for resistance to Myzus persicae, Neth. J. Pl. Pathol., 83:139-151, 1977. 27. Eenink, A. H., F. L. Dieleman, and R. Groenwold, Resistance of lettuce (Lactuca) to the leaf aphid Nasonovia ribisnigri. 2. Inheritance of resistance, Euphytica, 31:301-304, 1982. 28. Ellis, P. R., G. H. Freeman, and J. A. Hardman, Differences in the relative resistance of two carrot cultivars to carrot fly attack over five seasons, Ann. Appl. Biol., 105:557-564, 1984. 29. Ellis, P. R., and J. A. Hardman, The consistency of the resistance of eight carrot cultivars to carrot fly at several centers in Europe, Ann. Appl. Biol., 98:491-497, 1981. 30. Elsey, K. D. and E. V. Wann, Differences in infestation of pubescent and glabrous forms of cucumber by pickleworms and melonworms, HortSci., 17:253-254, 1982. 31. Equinas-Alcazar, J. T. and P. J. Gulick, Genetic Resources of Cucurbitacae — a Global Report, IBPGR, Rome, 1983. 32. Everson, E. H. and R. L. Gallun, Breeding approaches in wheat, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 513. 33. Farrar, R. R., Jr. and G. G. Kennedy, Insect and mite resistance in tomato, In: G. Kalloo, Ed., Genetic Improvements of Tomato, Monographs on Theoretical and Applied Genetics 14, Springer-Verlag, Berlin, Heidelberg, 1991, 121. 34. Fowler, C., Unnatural Selection: Technology, Politics, and Plant Evolution, Vol. 6, Gordon and Breach, Yverdon, 1994. 35. Gallun, R. L., K. J. Starks, and W. D. Guthrie, Plant resistance to insects attacking cereal crops, Ann. Rev. Entomol., 20:337-357, 1975. 36. Gentile, A. G. and A. K. Stoner, Resistance in Lycopersicon spp. to the tobacco flea beetle, J. Econ. Entomol., 61:1347-1349. 37. Gould, F., Predicting future resistance of crop varieties to pest populations: a case study of mites and cucumbers, Environ. Entomol., 7:622-626, 1978. 38. Gould, F., Resistance of cucumber varieties to Tetranychus urticae: genetic and environmental determinants, J. Econ. Entomol., 71: 680-683, 1978. 39. Gould, F., Genetics of plant-herbivore systems: interactions between applied and basic studies, In: R. F. Denno and M. S. McClure, Eds., Variable Plants and Herbivores in Natural and Managed Systems, Academic Press, New York, 1983, 599. 40. Gould, F., Simulation models for prediction durability of insect resistant germplasm: a deterministic diploid, two-locus model, Environ. Entomol., 15:1-10, 1986.

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41. Gould, F., Simulation models for prediction durability of insect resistant germplasm: Hessian fly-resistant winter wheat, Environ. Entomol., 15:11-23, 1986. 42. GRIN (Genetic Resources Information Network), United States Department of Agriculture, Agricultural Research Service, on the World Wide Web (http://www.arsgrin.gov/npgs/), 1997. 43. Griswold, M. J. and J. T. Trumble, Consumption and utilization of celery, Apium graveolens, by the beet armyworm, Spodoptera exigua, Entomol. Exp. Appl., 38:73-79, 1985. 44. Hare, J. D., Interactions between host-plant resistance and biological control, In: R. S. Fritz and E. S. Simms, Eds., Plant Resistance to Herbivores and Pathogens: Ecology, Evolution, and Genetics, University of Chicago Press, Chicago, 1992, 280. 45. Harlan, J. R., Our vanishing genetic resources, Science, 188:618-621, 1975. 46. Harlan, J. R. and K. J. Starks, Germplasm resources and needs, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 253. 47. Harris, M. K., Allopatric resistance: searching for sources of insect resistance for use in agriculture, Environ. Entomol., 4:661-669, 1975. 48. Holt, J. and S. D. Wratten, Components of resistance to Aphis fabae in faba bean cultivars, Entomol. Exp. Appl., 40:35-40, 1986. 49. Ingram, G. B. and J. T. Williams, In situ conservation of wild relatives of crops, In: J. W. Holden and J. T. Williams, Eds., Crop Genetic Resources: Conservation and Evaluation, George Allen and Unwin., London, 1988. 50. Jermy, T., Insect-host plant relationship: coevolution or sequential evolution, In: T. Jermy, Ed., The Host plant in Relationship to Insect Behavior and Reproduction, Plenum Press, New York, 1976. 51. Jermy, T., Evolution of insect-host plant relationships, Amer. Nat., 124:609-630, 1984. 52. Juvik, J. A. and M. A. Stevens, Inheritance of foliar α-tomatine content in tomatoes, J. Amer. Soc. Hort. Sci., 107:1061-1065, 1982. 53. Kalloo, G., Vegetable Breeding, Vol. II, CRC Press, Boca Raton, 1988. 54. Kalloo, G., Tomato (Lycopersicon esculentum Miller), In: G. Kalloo and B. O. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pergamon Press, Oxford, 1993, 645. 55. Kennedy, G. G., Recent advances in insect resistance of vegetable and fruit crops in North America: 1966-1977, Bull. Entomol. Soc. Am., 24:375-384, 1978. 56. Kennedy, G. G. and J. D. Barbour, Resistance variation in natural and managed systems, In: R. S. Fritz and E. S. Simms, Eds., Plant Resistance to Herbivores and Pathogens: Ecology, Evolution, and Genetics, University of Chicago Press, Chicago, 1992, 13. 57. Kennedy, G. G., G. W. Bohn, A. K. Stoner, and R. E. Webb, Leafminer resistance in muskmelon, J. Amer. Soc. Hort. Sci., 103:571-574, 1978. 58. Kennedy, G. G., F. Gould, O. M. B. DePonti, and R. E. Stinner, Ecological, agricultural, genetic, and commercial considerations in the deployment of insectresistant germplasm, Environ. Entomol., 16:327-338, 1987. 59. Kennedy, G. G., L. R. Romanow, S. F. Jenkins, and D. C. Sanders, Insects and diseases damaging tomato fruits in the coastal plain of North Carolina, J. Econ. Entomol., 76:168-173, 1983. 60. Kishaba, A. N., G. W. Bohn, and H.H. Toba, Resistance to Aphis gossypii in muskmelon, J. Econ. Entomol., 64:935-937, 1971.

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61. Kishaba, A. N., S. Castle, D. L. Coudriet, and J. D. McCreight, Field evaluation of melon aphid resistant cantaloupe breeding lines for susceptibility to the cucumber beetle complex, Cucurbit Genet. Coop. Rep., 6:41-45, 1983. 62. Kishaba, A. N., J. D. McCreight, D. L. Coudriet, T. W. Whitaker, and G. R. Pesho, Studies of ovipositional preference of cabbage looper on progenies from a cross between cultivated lettuce and prickly lettuce, J. Amer. Soc. Hort. Sci., 105:890-892, 1980. 63. Kishaba, A. N., T. W. Whitaker, L. Berry, and H.H. Toba, Cabbage looper oviposition and survival of progeny on leafy vegetables, HortSci., 11:216-217, 1976. 64. Kogan, M., Plant resistance in pest management, In: R. L. Metcalf and W. H. Luckmann, Eds., Introduction to Insect Pest Management, Wiley, New York, 1994, 73. 65. Kooistra, E., Red spider mite tolerance in cucumber, Euphytica, 20:47-50, 1971. 66. Kowalewski, P. A. and R. W. Robinson, White fly resistance in Cucumis, Bull. IOBC/WPRC, 77:3, 1977. 67. Lammerink, J., A new biotype of the cabbage aphid, Brevicoryne brassicae, on aphid resistant rape (Brassica napus L.), N. Z. J. Agric., 11:341-344, 1968. 68. Lammerink, J., and R. W. Hart, “Tina,” a new swede cultivar with resistance to dry rot and clubroot, N. Z. J. Agric., 13:417-420, 1985. 69. Langer, R. H. M. and G. G. Hill, Agricultural Plants, Cambridge University Press, Cambridge, 1991. 70. Leppik, E. E., Searching gene centers of the genus Cucumis through host-parasite relationship, Euphytica, 15:323-328, 1966. 71. Leppik, E. E., Gene centers of plants as sources of disease resistance, Ann. Rev. Phytopathology, 8:323-344, 1970. 72. Luckmann, W. H. and R. L. Metcalf, The pest-management concept, In: R. L. Metcalf and W. H. Luckmann, Eds., Introduction to Insect Pest Management, Wiley, New York, 1994, 1. 73. Magness, J. R., G. M. Markle, and C. C. Compton, Food and feed crops of the United States, Interregional Research Project IR-4, IR Bull.1, New Jersey Agr. Exp. Stn. Bull. 828, New Brunswick, 1971, on the World Wide Web (http://www.hort.purdue.edu/newcrop/ Crops), 1997. 74. Maxwell, F. G., and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980. 75. McCreight, J. D., A. N. Kishaba, and G. W. Bohn, AR Hale’s Best Jumbo, AR 5, and AR Topmark: melon aphid-resistant muskmelon breeding lines, HortSci., 19:309-310, 1984. 76. McCreight, J. D., H. Nerson, and R. Grumet, R., Melon (Cucumis melo L.), In: G. Kalloo and O. B. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pergamon Press, Oxford, 1993, 267. 77. Meade, T. and J. D. Hare, Differential performance of beet armyworm and cabbage looper (Lepidoptera: Noctuidae) larvae on selected Apium graveolens cultivars, Environ. Entomol., 20:1636-1644, 1991. 78. Metcalf, R. L. and A. M. Rhodes, Coevolution of the Cucurbitaceae and Luperini (Coleoptera: Chrysomelidae): basic and applied aspects, In: D. M. Bates, R. W. Robinson, and C. Jeffrey, Eds., Biology and Utilization of the Cucurbitaceae, Cornell University Press, New York, 1990, 167. 79. Myers, N., The Sinking Ark: A New Look at the Problem of Disappearing Species, Pergamon Press, Oxford, 1980.

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80. Namkoong, G., Preserving natural diversity, In: C. M. Schonewald-Cox, S. M. Chambers, B. MacBride, and W. L. Thomas, Eds., Genetics and Conservation, Benjamin Cummings, Menlo Park, 1983. 81. National Academy of Sciences, Alternative Agriculture, Committee on the Role of Alternative Farming Methods in Modern Production Agriculture, National Research Council, Washington, D.C., 1989. 82. National Academy of Sciences, Ecologically Based Pest Management: New Solutions for a New Century, National Research Council, Board on Agriculture, Washington, D.C., 1996. 83. Nielson, M. W. and W. F. Lehman, Breeding approaches in alfalfa, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 277. 84. Niles, G. A., Breeding cotton for resistance to insect pests, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 337. 85. Nugent, P. E., F. P. Cuthbert, Jr., and J. C. Hoffman, Two genes for cucumber beetle resistance in muskmelon, J. Am. Soc. Hortic. Sci., 109:756-759, 1984. 86. Ortega, A., K. S. Vasal, J. Mihm, and J. C. Hershey, Breeding for insect resistance in maize, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 371. 87. Ortman, E. E. and D. C. Peters, Introduction, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 3. 88. Pesho, G. R., L. W. Hudson, and T. N. Ingham, Differential feeding of cabbage loopers on Lactuca introductions, Special Report W-6, USDA-ARS Entomol. Res. Div., Grain and Forage Insects Res. Branch, Insect Resistance in New Crops Investigations, U.S. Department of Agriculture, Washington, D.C., 1969. 89. Pink, D. A. C. and E. A. Keane, Lettuce, Lactuca sativa L., In: G. Kalloo and O. B. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pergamon Press, Oxford, 1993, 543. 90. Pitrat, M. and H. Lecoq, Inheritance of resistance to cucumber mosaic virus transmission by Aphis gossypii in Cucumis melo, Phytopathology, 70:958-961, 1980. 91. Posylaeva, G. A., Trophic patterns in Acyrthosiphon pisum and breeding pea for resistance to the pest, Sel. Semenovo (Kiev), 68:76-79, 1990, (in Russian). 92. Quiros, C. F., Celery (Apium graveolens L.), In: G. Kalloo and O. B. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pregamon Press, Oxford, 1993, 523. 93. Quiros, C. F., M. A. Stevens, C. M. Rick, and M. L. Kok-Yokomi, Resistance in tomato to the pink form of the potato aphid (Macrosiphum euphorbiae Thomas); the role of anatomy, epidermal hairs, and foliage composition, J. Am. Soc. Hortic. Sci., 102:166-171, 1977. 94. Reinink, K. and F. L. Dieleman, Comparison of sources of resistance to leaf aphids in lettuce (Lactuca sativa L.), Euphytica, 40:21-29, 1989. 95. Reinink, K., F. L. Dieleman, J. Jansen, and A. M. Montenarie, Interactions between plant and aphid genotypes in resistance of lettuce to Myzus persicae and Macrosiphum euphorbiae, Euphytica, 43:215-222, 1989. 96. Rick, C. M., The tomato, Sci. Amer., 23:67-78, 1978. 97. Rick, C. M., Biosynthetic studies in Lycopersicon and closely related species of Solanum, In: J. G. Hawkes, R. N. Lester, and A. D. Skelding, Eds., The Biology and Taxonomy of the Solanaceae, Academic Press, New York, 1979. 98. Schalk, J. M., Plant resistance to insects in vegetables for the southern United States, Florida Entomol., 73:396-410, 1990.

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99. Scott, D. R., Lygus bugs feeding on developing carrot seed: plant resistance at that feeding, J. Econ. Entomol., 63:959-961, 1970. 100. Scott, D. R., Selection for Lygus bug resistance in carrot, HortSci., 12:452, 1977. 101. Simon, P. W., Carrot (Daucus carota L.), In: G. Kalloo and O. B. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pregamon Press, Oxford, 1993, 479. 102. Singh, A. K., Cytogenetics and evolution in the Cucurbitaceae, In: D. M. Bates, R. W. Robinson, and C. Jeffrey, Eds., Biology and Utilization of the Cucurbitaceae, Cornell University Press, New York, 1990, 10. 103. Smith, C. M., Plant Resistance to Insects: A Fundamental Approach, Wiley, New York, 1989. 104. Staub, J. E., J. Box, V. Meglic, T. Horejsi, and J. D. Mcreight, Comparison of isozyme and random amplified polynorphic DNA data for determining intraspecific variation in Cucumis, Genet. Resources and Crop Evol., in press. 105. Stoner, K. A., Bibliography of plant resistance to arthropods in vegetables, 1977-1991, Phytoparasitica, 20:125-180, 1992. 106. Tatlioglu, T., Cucumber (Cucumis sativus L.), In: G. Kalloo and O. B. Bergh, Eds., Genetic Improvement of Vegetable Crops, Pregamon Press, Oxford, 1993, 197. 107. Teetes, G. L., Breeding sorghums resistant to insects, In: F. G. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, Wiley, New York, 1980, 457. 108. Toll, J. and D. H. van Sloten, Directory of Germplasm Collections, 4. Vegetables, IBPGR Secretariat, Rome, 1982. 109. Trumble, J. T., W. Derecks, C. F. Quiros, and R. C. Beier, Host plant resistance and linear furanocoumarin content of Apium accessions, J. Econ. Entomol., 83:519-525, 1990. 110. Trumble, J. T. and C. F. Quiros, Antixenotic and antibiotic resistance in Apium species to Liriomyza trifolii (Diptera: Agromyzidae), J. Econ. Entomol., 81:602-607, 1988. 111. Wehner, T. C., K. D. Elsey, and G. G. Kennedy, Screening for cucumber antibiosis to pickleworm, HortSci., 20:1117-1119, 1985. 112. Whitaker, T. W., A. N. Kishaba, and H. H. Toba, Host-parasite interrelations of Lactuca saligna L. and the cabbage looper, Trichoplusia ni (Hübner), J. Am. Soc. Hort. Sci., 99:74-78, 1974. 113. Wilhoit, L. R., Evolution of herbivore virulence to plant resistance: influence of variety mixtures, In: R. S. Fritz and E. S. Simms, Eds., Plant Resistance to Herbivores and Pathogens: Ecology, Evolution, and Genetics, University of Chicago Press, Chicago, 1992, 91. 114. Astley, D., Personal communication, 1997.

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Section IV Root and Tuber Crops

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10

Utilization of Sweetpotato Genetic Resources to Develop Insect Resistance Wanda W. Collins, Edward E. Carey, Il-Gen Mok, Paul Thompson, and Zhang Da Peng

CONTENTS I. Introduction II. Characteristics of Sweetpotato A. Taxonomy B. Breeding Behavior C. Genetic Diversity III. Insect Pests IV. Using Ipomoea Genetic Resources for Insect Management A. Sweetpotato Weevil (Cylas spp.) B. Wireworm-Diabrotica-Systena Complex V. Conclusions References

I. INTRODUCTION Sweetpotato, Ipomoea batatas (L.) Lam., is one of the world’s major food crops.13 Although global production declined from 134 mmt in the late 1970s and early 1980s to 127 mmt in the early to mid-1990s, yield per hectare increased modestly over this time period (Table 1). China accounts for approximately 85% of the global production.13 Sweetpotato is used mainly for pig feeding in China. It also provides raw material for a number of processed and industrial products and serves as a basic food staple in some areas.15,40 Sweetpotato in Africa is very important in subsistence farming, especially in areas at high risk for food insecurity. It can grow and produce edible storage roots in marginal environments, where many food crops fail, and is a valuable crop for small, resource-constrained farmers.41 Sweetpotato is a regionally important crop in the southern U.S., where conditions for growth are most suitable.45 Sweetpotato is a perennial vine belonging to the family Convolvulaceae.2 It is cultivated in most areas of the world as an annual, producing a harvestable crop of fleshy roots in as few as 85 to 90 days. However, in areas of favorable conditions, roots will continue to grow and enlarge as long as they are left in the soil. The roots are

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TABLE 1 Global Area, Yield, and Production of Sweetpotatoes Region 1992–94 Average Africa North/Central America South America Asia Europe Oceania World 1992–94 average World 1979–81 average Percent change 79/81–92/94

Area harvested (000 ha)

Yield (mt/ha)

Production (mmt)

1,366 168 113 7,267 5 120 9,036 10,870 –17

5 7 11 16 13 5 14 12 14

6.79 1.11 1.22 117.52 0.06 0.59 127.25 134.23 –6

From FAOSTAT (Food Agriculture Organization of the United Nations, Production Statistics), on the World Wide Web (http://apps.fao.org/lim500/agri_db.pl), 1997.

highly perishable unless properly treated with heat and humidity (cured) and then stored at low temperatures.45 They can be stored in the ground until needed in tropical areas.41 Nutritionally, sweetpotato roots are high in energy and contain many essential vitamins and minerals.47 Orange-fleshed types, preferred in most of the developed world, are especially rich in vitamin A precursor because of the moderate to high carotene content. White-fleshed types, preferred in the developing world, are generally very low in carotene content.47 While protein content is minimal in the roots (1 to 10%), the net nitrogen availability from the protein is among the highest of all food crops.9,10 Constraints to sweetpotato production in developing countries are numerous, but it has been shown that investments in research to address problems yield high rates of return.18 Social and biological scientists at the International Potato Center (CIP), Lima, Peru, surveyed national programs, which are responsible for approximately 99% of the production in developing countries, to determine the most serious problems.20 Major constraints were postharvest quality and marketing problems. Insect damage accounts for increased postharvest quality problems and a decline in the quality of new planting material. Over 270 insects attack and damage sweetpotato roots throughout the world.37 On a global basis, the most important of these insect pests are Cylas sweetpotato weevils with crop losses of 50 to 100%.6 Other insect pests are usually important on a restricted regional or local basis.

II.

CHARACTERISTICS OF SWEETPOTATO

A. TAXONOMY Sweetpotato is a member of genus Ipomoea section Batatas (Choisy) Grisebach. The section contains 12 other species which are related to I. batatas at varying

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degrees.2 These species are primarily diploids (2N = 30), with the exception of the tetraploid I. tiliacea (Willd.) Choisy (2N = 60). Tetraploid races of I. batatas have been identified but not studied thoroughly.5 Taxonomists and plant breeders generally agree that I. trifida (H.B.K.) G. Don is the species most closely related to I. batatas. In 1956, Japanese researchers collected samples of I. trifida, which they determined to be hexaploid.32 However, Jones24 questioned the identity of this material and suggested it was feral I. batatas. The purported I. trifida hexaploids have been used extensively in crosses between I. batatas and wild species, and may have been the vehicle used to transfer insect resistance, among other traits, to I. batatas from a wild species. However, it is difficult to determine if the crosses represent an introgression of traits from a wild species or simply a wide-based cross within I. batatas.28 Sweetpotato is characterized by high ploidy level and a large number of very small chromosomes.29 It is a hexaploid with 90 chromosomes averaging 3.3 picograms of DNA per nucleus1 and has three genomes of uncertain origin and uncertain relatedness. Researchers generally agree that two of the genomes are related. The third may also be related but this has not been confirmed.29 Within the section Batatas, I. batatas is the only hexaploid and species characterized by the production of storage roots.34 There is evidence that I. trifida can produce rudimentary storage roots if given sufficient time and proper nutrition in culture17 which may further substantiate the relationship between the two taxa.

B. BREEDING BEHAVIOR Polyploidy was an ancient event in sweetpotato as evidenced by the high degree of “diploidization” in meiosis.29 Functionally, and for breeding purposes, meiotic chromosome behavior mimics a regular diploid. Even though cytogenetic examination confirms the diploid behavior, sweetpotato is still characterized by a high degree of sterility, perhaps owing to polyploid relics of meiosis (chromosome laggards and fragments). Sweetpotato is an out-crosser and it is self-incompatible. Several incompatibility groups exist, which are not fully understood, thereby preventing crossing between certain genotypes of the species. Incompatibility appears to be controlled sporophytically; however, plant breeders are not overly concerned about this incompatibility factor because the high degree of genetic diversity within the species facilitates their ability to create new combinations of useful genes.29 Ipomoea batatas is genetically isolated from other species in the section Batatas.33 Controlled crosses are successful with a few other Batatus species but at extremely low frequencies. The degree of crossability has been misstated in the past because of problems associated with incorrect species identifications. Researchers at CIP have crossed I. batatas with its nearest relative I. trifida and produced tetraploid progeny.21,34 Diaz et al.12 devised a polygon crossing scheme that uses I. trifida and I. x leucantha Jacq. as bridge species for gene flow from wild Ipomoea species to the genepool of sweetpotato. Flowering in I. batatas can be a problem because the species has a short-day flowering habit.29 The species can flower sparsely or not at all under ideal conditions. The reasons for this problem are unknown and breeders generally graft onto a flowering rootstock of other morning glory species to induce flowering. Because of

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the wide diversity available for most traits in sweetpotato, breeders usually discard non-flowering types and use flowering types that exhibit the traits of interest.

C. GENETIC DIVERSITY Sweetpotato, in almost all traits studied thus far, is characterized by tremendous genetic diversity. Inheritance of traits is most often quantitative with only a few traits controlled by one or two genes.29 Many of the agronomically important traits (e.g., resistance to Fusarium wilt disease, root-knot nematodes) are controlled by additive gene action and are easily amenable to recurrent selection. Others, such as yield, have non-additive inheritance patterns which makes it more difficult to improve. However, the inherent genetic diversity in sweetpotato has not been fully exploited to develop agronomically superior genotypes. Rao et al.39 estimated that 26,020 accessions of Ipomoea spp., mostly I. batatas, are held by 83 organizations throughout the world. The CIP stores and maintains the international collection of I. batatas consisting of over 5,500 Ipomoea accessions collected from all parts of the world. Over 3,000 accessions are I. batatas cultivars and approximately 1,500 are breeding lines. Additionally, eight related species in section Batatas are represented by over 800 accessions. The CIP collection also includes germplasm of 48 species from other sections of the genus.39 The U.S. collection is maintained by the U.S. Department of Agriculture, Agricultural Research Service (USDA-ARS), Southern Regional Plant Introduction Station, Griffin, Georgia, which holds approximately 1,106 Ipomoea accessions.14 Many of the world’s collections have not been adequately or systematically characterized and evaluated. Evaluations tend to be carried out for specific traits such as yield, eating or cooking, and disease resistance. Characterization and evaluation of sweetpotato germplasm are limited because sweetpotato is usually accorded low priority in decisions for resource allocations. Because most traits are inherited in a quantitative fashion and are unstable over environments, adequate evaluations would require multi-year and -location testing. These types of tests are often costly and beyond the means of researchers in developing countries. Germplasm characterization and evaluation studies indicate that considerable variation exists in morphological traits and growth habit.4,15,17,20-22,28,29 Extensive variation has been reported in horticultural traits such as root dry matter, starch, carotene, sugar, and protein;10 disease and insect resistance;25 and fiber, root weight, shape, cracking, and sprouting.27 Genetic diversity studies of similar traits in related species are precluded by their lack of storage roots. An exception is the variability reported in Japanese accessions of I. trifida at various ploidy levels, including the purported hexaploid.

III.

INSECT PESTS

Worldwide, the most devastating pests of sweetpotato are Cylas sweetpotato weevils.6,41 The damage to sweetpotato by all other insect pests combined may exceed that caused by Cylas weevils alone, but no one pest causes as much damage as Cylas

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FIGURE 1 Resistance reaction of sweetpotato clones to sweetpotato weevil, Cylas formicarius, in dry and rainy season testing in Indonesia.

weevils. Although several genera and species of “sweetpotato weevil” exist, the term usually refers to Cylas formicarius F., C. brunneus F., and C. puncticollis Bohe. The three species occur in Africa, with C. brunneus and C. puncticollis known to occur only on the continent.16,41,46 The distribution and damage caused by each species also vary within African countries. Cylas formicarius is circumglobal in distribution and is the single most important insect pest of sweetpotato.6,41 Sweetpotato weevils attack stems, crowns, and roots of sweetpotato plants.41 Adults feed directly on leaves and vines. Damage to fleshy storage roots is severe under dry conditions (Figure 1) as cracks in the soil provide adult weevils with access to the roots for egg laying. Subsequently, larvae cause significant damage by tunneling into the roots. Many Ipomoea species, mostly unrelated to sweetpotato, serve as alternate hosts to weevils.3 There are several other weevils of lesser importance on a regional basis. The West Indian sweetpotato weevil, Euscepes postfasciatus Fairmaire, is important in South America and in the Caribbean.38 The rough weevil, Blosyrus spp., and the striped weevil, Alcidodes dentipes Oli, are destructive pests of sweetpotato in East Africa.41 A group of insects collectively known as The Wireworm–Diabrotica–Systena (WDS) complex is of regional importance. These insects are the southern potato wireworm Conoderus falli Lane, tobacco wireworm C. vespertinus F., banded cucumber beetle Diabrotica balteata LeConte, spotted cucumber beetle D. undecimpunctata howardi Barber, elongate flea beetle Systena elongata F., pale-striped flea beetle S. blanda Melsheimer, and the flea beetle S. frontalis F.11,25 This group is treated as a complex because damage caused by any one of the species is difficult

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to distinguish. The WDS causes significant surface damage to sweetpotato roots in the southern U.S. Damage is characterized by small feeding holes or scars that are enhanced by subsequent growth of the root.25 This damage is severe and economic because of high U.S. market quality standards. Roots with surface damage caused by WDS do not meet market acceptability standards and must be discarded or sold at drastically reduced prices for processing or animal feed. A large number of other insects, i.e., stem borers, leaf miners, and webbing insects, seriously damage sweetpotato in some environments.36,41 These pests have not been the subject of long-term and systematic research on host-plant resistance. Reports of localized evaluations or observations are numerous and are valuable for making local decisions for control measures. However, the magnitude of damage resulting from these minor insect infestations, on an annual and global basis, is insignificant compared with that caused by Cylas weevils, and they will not be discussed here.

IV.

USING IPOMOEA GENETIC RESOURCES FOR INSECT MANAGEMENT

Sweetpotato germplasm, and that of its closest relatives, has been well collected on a global basis. Many accessions have been locally observed or evaluated for their reaction to insect pests. But these efforts often have not been systematic or undertaken as part of a focused breeding program to improve resistance to insects. Therefore, results from these observations and studies have limited value in establishing the level of consistent genetic variability which might be useful to a breeding program. In general, the lack of sustained, long-term testing can be attributed to several causes including: difficulty in developing and refining effective, consistent, and reliable evaluation methods in the laboratory and field; the low levels of resistance heretofore identified by researchers; the quantitative mode of the inheritance of resistance; the long-term nature of the breeding effort required to work with quantitative inheritance of low levels of resistance; and the lack of funding and commitment to long-term breeding objectives. The two exceptions include the longterm evaluation and breeding programs for resistance to Cylas weevils in several countries and the WDS complex in the U.S.

A. SWEETPOTATO WEEVIL (CYLAS

SPP.)

Field evaluations often lead to high experimental error because of the uneven distribution of insects and their ability to move quickly.44 Changes in environmental conditions, such as rain or drought, also have significant impact on results as shown in a CIP study of the reaction of sweetpotato clones to infestation by C. formicarius in Southeast Asia (Figure 1). Clones responded differently in wet and dry seasons with almost all clones rated highly susceptible in the dry season. Laboratory tests can be more unreliable than field tests and the results often do not correlate well with field observations. For example, the results from laboratory feeding tests did not correspond well with the results from field tests in the Philippines.4

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Despite the problems with field testing, a number of international centers have used this approach to generate data on the reaction of sweetpotato genotypes to weevils. The Asian Vegetable Research and Development Center, Taiwan, Republic of China, field-tested over 1,200 sweetpotato accessions for resistance to C. formicarius. Some genotypes showed good resistance initially and were selected for further study; however, the resistance was ineffective in subsequent tests. Therefore, stable sources of resistance have not been located.43 The lack of consistency led to the conclusion that resistance was probably not present in sweetpotato germplasm.42 Years of similar research at the International Institute for Tropical Agriculture (IITA), Nigeria, produced much the same result with the African sweetpotato weevil, C. puncticollis, although very low levels of resistance were identified.16 However, the IITA resistance breeding program has been discontinued. Researchers at CIP evaluated sweetpotato germplasm in the 1990s in several regions of the world, including East Africa and Southeast Asia where weevil infestations are high and damage is severe. Because levels of resistance were low, it was concluded that sweetpotato had no useful resistance to Cylas weevils.22 Consequently, CIP changed its approach from screening germplasm for resistance to Cylas weevils to one employing gene transfer with more modern methods.22 Sweetpotato has been transformed through both Agrobacterium-mediated and bombardment protocols,35,36 and clones of I. batatas transformed with the Bt-resistant gene are being field-tested by CIP.48 Transgenic plants expressing genes for cowpea trypsin inhibitor and snowdrop lectin also have been produced but have not been field-tested for insect resistance (see Chapter 13 in this volume).31 The introduction and expression of these genes opens an avenue for incorporating insect resistance into sweetpotatoes which might not be accomplished otherwise. Researchers at CIP also have evaluated species related to sweetpotato for reaction to Cylas. However, the lack of storage root formation in wild species has limited the evaluations to stem characteristics. Although some variability in reaction was seen very early in tetraploid hybrids between I. batatas and I. trifida at CIP, researchers concluded that useable levels of resistance do not exist in the wild relatives of I. batatas.22 It would be worthwhile to evaluate the naturally occurring I. batatas tetraploid for Cylas resistance if this germplasm has not been evaluated. Several national programs have evaluated clones for resistance to C. formicarius. In the U.S., Mullen et al.30 screened genotypes using a laboratory feeding procedure and identified low levels of resistance. Two cultivars and six breeding lines with moderate levels of weevil resistance were released in Georgia and South Carolina.26 However, researchers in Florida found that the resistant cultivars were susceptible under growing conditions in naturally infested fields, demonstrating once again the inconsistency in resistance over time and location.23 Thompson et al.44 reported a moderate genetic gain via intermating clones with low levels of resistance. Based on this research, 100 plant introductions of various origin were evaluated in 1993–1994 in an attempt to find additional sources of resistance to combine with an earlier population.49 This effort showed that genetic variability for resistance to Cylas is available in plant introductions (Figure 2), although at low levels. However,

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FIGURE 2 Index of damage to roots and stems of 100 sweetpotato U.S. Plant Introductions by the sweetpotato weevil, Cylas formicarius.

a number of introductions exhibited resistance levels higher than the highest level recorded previously with an intermated population. Evaluation of local germplasm has been conducted in many developing countries. Tests were usually carried out in the field with natural insect infestations and a simple rating procedure to determine damage. While differences between genotypes were usually discernible each year, it was unclear whether the variability was caused by environmental effects, insect distribution, or true genetic resistance. Only Philippine researchers have successfully increased the levels of resistance through intermating and selection.4 They observed gradual increases in levels of resistance, but the levels are too low for field control of weevils. Cultivars have been developed, released, and used in an integrated approach to weevil management. Given the lack of moderate levels of weevil resistance in sweetpotato germplasm, researchers are increasingly advocating the integrated approach to weevil management as demonstrated in the Philippines.4,8,21,41,43 This approach emphasizes cultural methods, biological control organisms which are now available and moderately successful, low levels of genetic resistance, and pesticides when absolutely necessary. Based on the evaluations and breeding efforts of many researchers, it appears that variability exists among sweetpotato germplasm for resistance to Cylas weevils, as first reported 50 years ago by Cockerman and Deen.7 However, the collective evidence shows that resistance levels are low and sweetpotato reactions are inconsistent and unstable across environments. Nothing less than a long-term, sustained, and systematic breeding effort will be required to develop resistance to a level that can be used in an integrated approach to weevil management.8

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FIGURE 3 Frequency of Wireworm-Diabrotica-Systena (WDS) injury to selected (solid line) and non-selected (broken line) sweetpotato clones. Area marked by “a” represents original (non-selected) clones with resistance, “b” represents portions of selected clones with resistance levels exceeding those found in the original clones, and the area marked “c” represents the additional selection potential for resistance in the selected population. (Modified from Cuthbert, F.P., Jr. and A. Jones, Resistance in sweet potatoes to Coleoptera increased by recurrent selection, J. Econ. Entomol., 65:1655-1658, 1972. [Ref. 11])

B. WIREWORM-DIABROTICA-SYSTENA COMPLEX The WDS complex is a serious problem in the southern region of the U.S. where sweetpotato is a major vegetable crop. While yields are seldom affected per se, significant economic damage results from unmarketable roots showing insect damage. Efforts to improve sweetpotato resistance to the WDS complex through selection and breeding were conducted at the USDA-ARS Vegetable Laboratory, Charleston, South Carolina. This effort represents the most successful use of I. batatas genetic resources for developing insect resistance.11,25 Selection began on a population that was derived from 19 accessions of diverse origin. An open-pollinated, randommating population was established from this population. Recurrent selection for insect resistance, based on the aggregate damage of the complex of insects, continued for several generations by choosing parents, intermating them, testing the seedlings, re-selecting, and beginning the process again. After four generations of recurrent selection, the improved (selected) population was compared with the control (nonselected) population to determine if genetic gain had been achieved. Indeed, the selected population had resistance levels higher than those in the non-selected population (Figure 3). Moreover, there was a significant decrease in the mean level of damage in the selected population. The results of this systematic approach demonstrate

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that breeding for quantitative resistance can be an effective mechanism to increase insect resistance using naturally occurring genes.11 It also implies that a similar breeding procedure for sweetpotato weevil resistance might be possible.

V.

CONCLUSIONS

Insects pose a serious constraint to global production of sweetpotatoes. The magnitude of damage caused by C. formicarius, the most important insect pest, in global sweetpotato production areas justifies the continued search for resistant germplasm. It is possible that the low levels of resistance heretofore detected in sweetpotato germplasm could be increased through a directed intercrossing and selection program using the vast array of genetic diversity of I. batatus stored in genebanks. This breeding approach would require a consistent and ongoing commitment of resources for a number of years, an unlikely scenario in an era of declining resources for agricultural research. An alternative approach is the use of biotechnological methods to develop insectresistant sweetpotatoes. Transgenic plants containing insect resistance genes are now in various stages of field testing. If this approach proves successful, it might be the preferred and more economical approach for incorporating resistance to various insect pests in this genetically complex crop. However, continued field evaluation and observation of the conserved genetic resources of I. batatas must continue. Such efforts might identify more sources of resistant germplasm which could facilitate the development of an integrated pest management approach. New and more efficient and cost-effective evaluation methods must be developed to assist in the breeding of insect-resistant cultivars. This will require added investments in sweetpotato research on a global basis.

REFERENCES 1. Armuganathan, K. and E. D. Earle, Nuclear DNA content of some important plant species, Plant Mol. Biol. Rep., 9:208-218, 1991. 2. Austin, D. F., The Ipomoea batatas complex-I. Taxonomy, Bull. Torrey Bot. Club 105:114-129, 1978. 3. Austin, D. F., Associations between the plant family Convolvulaceae and Cylas weevils, In: R. K. Jansson and K. V. Raman, Eds., Sweet Potato Pest Management: A Global Perspective, Westview Press, Boulder, 1991, 45. 4. Bacusmo, J. L., V. Z. Acedo, A. M. Mariscal, and M. Z. Oracion, Sweetpotato genetic resources in the Philippines, In: MAFF Int. Workshop on Genetic Resources: Root and Tuber Crops, Tsukuba, Ministry of Agriculture, Forestry and Fisheries, Tokyo, 1994, 103. 5. Bohac, J. R., D. R. Austin, and A. Jones, Discovery of wild tetraploid sweetpotatoes. Econ. Bot., 47:193-201, 1993. 6. Carey, E. E., E. Chujoy, T. Dayal, H. M. Kidanemariam, H. A. Mendoza, and Il-Gen Mok, Helping meet varietal needs of the developing world: the International Potato Center’s strategic approach to sweetpotato breeding, In: W. A. Hill, C. K. Bonsi, and P. A. Loretan, Eds., Sweetpotato Technology for the 21st Century, Tuskegee University, Tuskegee, 1992, 521.

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7. Cockerham, K. L. and O. T. Deen, Resistance of new sweet potato seedlings and varieties to attack by the sweetpotato weevil, J. Econ. Entomol., 40:439-441, 1947. 8. Collins, W. W., Variability in sweet potato germplasm for food quality characteristics, In: R. H. Howeler, Ed., Proc. 8th Int. Symp. Tropical Root and Tuber Crops, Bangkok, 1990, 724. 9. Collins, W. W., A. Jones, M. A. Mullen, N. S. Talekar, and F. W. Martin, Breeding sweet potato for insect resistance: a global overview, In: R. K. Jansson and K. V. Raman, Eds., Sweet Potato Pest Management: A Global Perspective, Westview Press, Boulder, 1991, 379. 10. Collins, W. W. and W. M. Walter, Jr., Potential for increasing nutritional value of sweetpotatoes, In: R. L. Villareal and T. D. Griggs, Eds., Sweet Potato, Proc. 1st Int. Symp., Asian Vegetable Research and Development Center (AVRDC), Shanhua, Taiwan, 1982, 355. 11. Cuthbert, F. P., Jr. and A. Jones, Resistance in sweet potatoes to Coleoptera increased by recurrent selection, J. Econ. Entomol., 65:1655-1658, 1972. 12. Diaz, J., P. Schmiediche, and D. F. Austin, Polygon of crossability between eleven species of Ipomoea: section Batatas (Convolvulaceae), Euphytica, 88:18-200, 1996. 13. FAOSTAT (Food Agriculture Organization of the United Nations, Production Statistics), on the World Wide Web (http://apps.fao.org/lim500/agri_db.pl), 1997. 14. GRIN (Genetic Resources Information Network), United States Department of Agriculture, Agricultural Research Service, on the World Wide Web (http://www.arsgrin.gov/npgs/), 1997. 15. Guo, X. D., Y. H. Wang, J. Y. Wu, and J. L. Sheng, Maintenance and use of sweetpotato germplasm in China, In: MAFF Int. Workshop on Genetic Resources: Root and Tuber Crops, Tsukuba, Ministry of Agriculture, Forestry and Fisheries, Tokyo, 1994, 121. 16. Hahn, S. K. and K. Leuschner, Resistance of sweetpotato cultivars to African sweetpotato weevil, Crop Sci., 21:499-503, 1981. 17. Hambali, G., Tuberization in diploid Ipomoea trifida from Citatah, West Java, Indonesia, In: W. A. Hill, C. K. Bonsi, and P. A. Loretan, Eds., Sweetpotato Technology for the 21st Century, Tuskegee University, Tuskegee, 1992, 469. 18. Ho, T. V., Root and tubers crop genetic resources in Vietnam, In: MAFF Int. Workshop on Genetic Resources: Root and Tuber Crops, Tsukuba, Ministry of Agriculture, Forestry and Fisheries, Tokyo, 1994, 167. 19. Horton, D., G. Prain, and P. Gregory, High level investment returns for global sweet potato research and development, CIP Circular 17, Centro Internacional de la Papa (CIP), Lima, 1989. 20. International Potato Center, The International Potato Center Annual Report, Centro Internacional de la Papa (CIP), Lima, 1991, 130. 21. International Potato Center, Program Report 1993-1994, Centro Internacional de la Papa (CIP), Lima, 1995. 22. International Potato Center, CIP in 1995: The International Potato Center Annual Report, Centro Internacional de la Papa (CIP), Lima, 1996. 23. Jansson, R. K., H. H. Bryan, and K. A. Sorensen, Within-vine distribution and damage of sweetpotato weevil, Cylas formicarius elegantulus (Coleoptera: Curculionidae) on four cultivars of sweet potato in southern Florida, Florida Ent., 70:523-526, 1987. 24. Jones, A., Should Nishiyama’s K123 (Ipomoea trifida) be designated I. batatas?, Econ. Bot., 21:163-166, 1967.

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25. Jones, A., P. D. Dukes, and F. P. Cuthbert, Jr., Mass selection in sweet potato: breeding for resistance to insects and diseases and for horticultural characteristics, J. Am. Soc. Hort., 101:701-704, 1976. 26. Jones, A., P. D. Dukes, J. M. Schalk, M. A. Mullen, M. G. Hamilton, D. R. Paterson, and T. E. Boswell, W-71, W-115, W-119, W-125, W-149 and W-154 sweet potato germplasm with multiple insect and disease resistances, HortSci., 15:835-836, 1980. 27. Jones, A., M. G. Hamilton, and P. D. Dukes, Heritability estimates for fiber content, root weight, shape, cracking and sprouting in sweetpotato, J. Am. Soc. Hort. Sci., 103:374-376, 1978. 28. Komaki, K., Sweetpotato genetic resources and breeding in Japan, In: MAFF Int. Workshop on Genetic Resources: Root and Tuber Crops, Tsukuba, Ministry of Agriculture, Forestry and Fisheries, Tokyo, 1994, 115. 29. Martin, F. W. and A. Jones, Breeding sweetpotatoes, Plant Breed. Rev., 4:313-345, 1986. 30. Mullen, M. A., A. Jones, D. R. Paterson, and T. E. Boswell, Resistance in sweetpotatoes to the sweetpotato weevil, Cylas formicarius elegantulus (Summers), J. Entomol. Sci., 20:345-350, 1985. 31. Newell, C. A., J. M. Lowe, A. Merryweather, L. M. Rooke, and W. D. O. Hamilton, Transformation of sweet potato (Ipomoea batatas L.) Lam.) with Agrobacterium tumefaciens and regeneration of plants expressing cowpea trypsin inhibitor and snowdrop lectin, Plant Sci., 107:215-227, 1995. 32. Nishiyama, I., Evolution and domestication of the sweet potato, Bot. Mag. Tokyo, 84:377-387, 1971. 33. Oracion, M. J. and I. Shiotani, Cytology and fertility of sweetpotato x diploid Ipomoea trifida F1 hybrids and their potential in analytic breeding of sweetpotato, In: W. A. Hill, C. K. Bonsi, and P. A. Loretan, Eds., Sweetpotato Technology for the 21st Century, Tuskegee University, Tuskegee, 1992, 540. 34. Orjeda, G., M. Iwanaga, and R. Freyre, Use of Ipomoea trifida germplasm for sweetpotato improvement: evaluation of storage root initiators, In: W. A. Hill, C. K. Bonsi, and P. A. Loretan, Eds., Sweetpotato Technology for the 21st Century, Tuskegee University, Tuskegee, 1992, 546. 35. Prakash, C. S. and U. Varadarajan, Expression of foreign genes in transgenic sweetpotatoes, In: Proc. Int. Soc. Plant. Mol. Biol., Tucson, 1991, (Abstract). 36. Prakash, C. S. and U. Varadarajan, Genetic transformation of sweet potato by particle bombardment, Plant Cell Rep., 11:53-57, 1992. 37. Raman, K. V., Major sweetpotato insect pests and selection for resistance to the sweetpotato weevil Euscepes postfasciatus (Fairmaire), In: Improvement of Sweet Potato (Ipomoea batatas) in East Africa, Rep. Workshop on Sweet Potato Improv. Africa, ILRAD, Nairobi, Kenya, 1987, International Potato Center (CIP), Lima, 1988, 208. 38. Raman, K. V. and E. H. Alleyne, Biology and management of the West Indian sweetpotato weevil, Euscepes postfaciatus, In: R. K. Jansson and K. V. Raman, Eds., Sweet Potato Pest Management: A Global Perspective, Westview Press, Boulder, 1991, 263. 39. Rao, R. V., D. G. Debouck, and M. Iwanaga, The role of international organizations in root and tuber crops conservation, In: MAFF Int. Workshop on Genetic Resources: Root and Tuber Crops, Tsukuba, Ministry of Agriculture, Forestry and Fisheries, Tokyo, 1994, 7.

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40. Sheng, J. L., Q. H. Xue, and D. P. Zhang, Sweetpotato breeding, production and utilization in China, In: W. A. Hill, C. K. Bonsi, and P. A. Loretan, Eds., Sweetpotato Technology for the 21st Century, Tuskegee University, Tuskegee, 1992, 153. 41. Smit, N. E. J. M., Integrated Pest Management for Sweetpotato in Eastern Africa. Grafisch Service Centrum Van Gils, B. V., Wageningen, 1997. 42. Talekar, N. S., Feasibility of the use of resistant cultivar in sweetpotato weevil control, Insect Sci. Appl., 8:815-817, 1987. 43. Talekar, N. S., Development and testing of an integrated pest management technique to control sweet potato weevil, In: Improvement of Sweet Potato (Ipomoea batatas) in Asia, Rep. Workshop Sweet Potato Improv. Asia, The International Potato Center, Lima, 1989, 256. 44. Thompson, P. G., J. C. Schneider, and B. Graves, Genetic variance components and heritability estimates of freedom from weevil injury to sweetpotato, J. Am. Soc. Hort. Sci., 119:620-623, 1994. 45. Wilson, L. G., Growing and marketing quality sweetpotatoes. North Carolina Agric. Ext. Serv. Bull. AG09, N. C. State University Agricultural Extension Service, Raleigh, 1989, 28. 46. Wolfe, G. W., The origin and dispersal of the pest species of Cylas with a key to the pest species groups of the world, In: R. K. Jansson and K. V. Raman, Eds., Sweet Potato Pest Management: A Global Perspective, Westview Press, Boulder, 1991, 13. 47. Woolfe, J. A., Sweet Potato: An Untapped Food Resource, Cambridge University Press, New York, 1992. 48. Peng, Z.D., Personal communication, 1997. 49. Thompson, P., Personal communication, 1997.

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11

The Potato: Genetic Resources and Insect Resistance Kathy L. Flanders, Silvia Arnone, and Edward B. Radcliffe

CONTENTS I. Introduction II. Germplasm Resources III. Potato Breeding IV. Pest Resistance in Exotic Potatoes V. Use of Resistant Germplasm in Breeding VI. Conclusions Acknowledgments References

I.

INTRODUCTION

Potato, Solanum tuberosum ssp. tuberosum L., is the fourth most important food crop in terms of production and cash value.62 Tubers contain high quality protein and substantial amounts of essential vitamins, minerals, and trace elements. Potato yields more useable energy per unit area of production than any other major food crop. Raw cereals and beans yield more protein than potato, but the protein content of cooked potato is almost equivalent to beans and exceeds cereals. Potato originated, botanically and as an agricultural crop, in the Andean highlands around Lake Titicaca.49 It was introduced to Europe by the Spanish in the latter half of the 16th century. However, for the next 200 years potato remained little more than a botanical curiosity because the introductions of tropical origin were not adapted to tuberize under the long summer days of western Europe. With continual selection for early tuberization, potato eventually became Europe’s most important food crop. In the mid-1960s, almost 87% of the world’s potato production occurred in central, northern, and eastern Europe.31 Today, the crop is grown in more than 130 countries, which are home to three-quarters of the world’s population. In the past 35 years, production in the developing world has increased more rapidly than

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any major food crop and at twice the rate of total food production. Factors driving expansion in developing countries include high yields of energy and protein possible per hectare under adverse circumstances, practicality of incorporating potato production into the multiple cropping systems of subsistence farmers, strong consumer demand for potatoes offering potential commercial profitability, and rapid technological advances in cultivar development, seed production systems, pest control, and post-harvest storage and processing.62 Numerous potential insect pests occur in potato ecosystems. Boiteau14 collected 565 taxa in a 3-year study of potato fields in New Brunswick, Canada. Radcliffe et al.96 listed 170 arthropod pest species in North America and that list was certainly incomplete, especially with respect to the aphid vectors of potato potyviruses and the soil inhabiting species of occasional or localized importance on stems and tubers. But, the greatest diversity of arthropods associated with potato occurs in South and Central America where potato and its wild relatives originated.98 A few insect species are potato pests of worldwide importance. Green peach aphid, Myzus persicae (Sulzer), is the most important vector of potato viruses worldwide. Indeed, more than half of all the studies on potato insect pests relate to this species. The next most important pest is Colorado potato beetle, Leptinotarsa decemlineata (Say). This insect has received more attention in North America and Europe over the past 30 years than any other potato pest because of its propensity to develop insecticide resistance. In the subtropics and tropics, potato tuber moth (= potato tuberworm), Phthorimaea operculella (Zeller), is often the most damaging pest, especially where potatoes are held in traditional storages. In each region and production system the potato pest complex is somewhat different, but invariably important. There are few, if any, places in the world in which potatoes are grown commercially without the routine use of both insecticides and fungicides. Insect-resistant potatoes are needed as the global need for food increases and as insect pests develop resistance to insecticides. This chapter examines how potato germplasm is being used to create insect pest resistant potatoes, with emphasis on M. persicae, L. decemlineata, and P. operculella. Information on nematode resistance, disease resistance, and stress tolerance is also included.

II.

GERMPLASM RESOURCES

The potato is unique among crop plants in the diversity of its wild relatives. There are approximately 230 species of potatoes (Solanaceae, Section Petota).49,90 The systematic interpretation of Hawkes, used throughout this chapter, groups the potatoes into 21 series of more closely related species.49 The series Tuberosa is the largest series with 104 species, including S. tuberosum. Wild potatoes are native to the area from southwestern U.S., southward through Mexico and Central America, along the Andes of South America, and into the plains of Argentina, Paraguay, and Uruguay.49 Each wild potato species has its own range of ecogeographic adaptation and few species occur over a wide geographic area. Wild potatoes possess a wide variety of defense mechanisms against insects, including glandular trichomes,48,124 foliar and tuber glycoalkaloids,32,123 and hypersensitive © 1999 by CRC Press LLC

reaction to the presence of insect eggs.7 Studies in the last two decades have increased our understanding of the plant mechanisms conferring insect resistance.125 Potato germplasm, in the form of true potato seed, tubers, or in vitro plantlets, is held in genebanks throughout the world. The most important collections are Central Colombia Collection, Tibaitata, Colombia; Collection of Tuberous Solanum of Argentina, Balcarce, Argentina; Commonwealth Potato Collection, Pentlandfield, Scotland;109 Brunswick Genetic Resources Centre, Braunschweig, Germany;22 Centre for Genetic Resources, Wageningen, The Netherlands;26 Gross Lusewitz Potato Species Collection of the Institut für Pflanzengenetik und Kulturpflanzenforshung Gatersleben, Gross Lusewitz, Germany;61 International Potato Collection (CIP), Lima, Peru;64 U.S. National Research Support Program 6 (NRSP-6), Potato Introduction Project Collection, Sturgeon Bay, Wisconsin, U.S.;84 Vavilov Institute of Plant Industry Collection, Leningrad, Russia;130 and the Chilean Tuberous Solanum Collection, Valdivia, Chile. Genebanks maintain databases with information about each accession. Passport data, such as the site and date of collection and original collector identification number, is usually available for each accession. Information on pest resistance and environmental stress tolerance varies with genebank. The Intergenebank Potato Database serves as a global clearing house of information on potato germplasm,60 and for 8,600 accessions belonging to 195 Solanum species it describes many traits, including resistance to insect pests.

III.

POTATO BREEDING

Indigenous potatoes cultivated in South America include: diploids (2N = 2X = 24) S. phureja Juz. et Buk., S. stenotomum Juz. et Buk., and S. ajanhuiri Juz. et Buk.; triploids (2N = 3X = 36) S. chaucha Juz. et Buk. and S. juzepczukii Buk.; tetraploid (2N = 4X = 48) S. tuberosum ssp. andigena Hawkes, the immediate ancestor of our present-day S. tuberosum ssp. tuberosum cultivars; and pentaploid (2N = 5X = 60) S. curtilobum Juz. et Buk.49 Many landraces of these species are selections with adaptation to particular area and specific horticultural traits. The cultivated potato, S. tuberosum ssp. tuberosum, is a vegetatively propagated autotetraploid species with tetrasomic inheritance and high heterozygosity. Its genetic variability is narrow in regard to resistance to biotic stresses. Classical breeding strategies have been based on sexual crosses between and within closely related species. However, the tetrasomic segregation of traits in S. tuberosum ssp. tuberosum makes it difficult to determine the genetics of trait inheritance. This can necessitate selection from enormous seedling populations.102 Unfortunately, detrimental traits from wild species are often introduced along with the desired trait. This necessitates backcross breeding to obtain genotypes with acceptable agronomic traits. Presence of interspecific incompatibility barriers makes incorporation of most wild genes difficult. On the other hand, the wild potatoes, of which 180 species are tuber-bearing,49 represent an enormous resource of genetic variability. Recent advances in plant breeding techniques overcome many limitations of traditional breeding techniques.5,25,70 This may make possible the exploitation of previously identified sources of resistance not possible to transfer to S. tuberosum © 1999 by CRC Press LLC

by conventional breeding methods. These advances invite screening of unexplored germplasm not previously considered of practical utility. Hougas and Peloquin57 devised a variation of conventional breeding methods based on haploid S. tuberosum ssp. tuberosum (2N = 2X = 24). This method permitted crosses with wild diploid species,67 and allowed breeders to work with disomic or monosomic segregation. Successful selection can be achieved with fewer seedlings and fewer generations than in breeding at the tetraploid level. Such crosses are also more likely to reveal undesirable traits.102 Haploids can be obtained by gynogenesis through crosses with S. phureja that induce production of unreduced gametes,10 by in vitro microspore culture (androgenesis)102 or through anther culture.115 Peloquin93 proposed the following crossing schemes for generating 4X progenies: 2X × 2X, 4X × 2X, 2X × 4X, where 2X is an interspecific haploid hybrid with unreduced gametes, and 4X is a tetraploid cultivar. These methods have been used to transfer insect resistance (L. decemlineata and M. persicae) in S. berthaultii Hawkes125 and P. operculella resistance in S. sparsipilum (Bitt.) Juz. et Buk.92 to S. tuberosum. Embryo rescue, combined with diploid breeding theory, have allowed introgression of genes from S. brevidens Phil., a non-tuber bearing species distantly related to S. tuberosum.136 In vitro culture techniques (micropropagation, regeneration) have been used in conjunction with physical or chemical agents to promote mutagenesis. This has allowed potato breeders to take advantage of both genetic and somaclonal variation, use precocious in vitro selection techniques, and overcome certain interspecific incompatibilities.117 For example, X-rays have been used to induce mutations and break genetic linkages between undesirable wild traits and trichome B presence in S. tuberosum x S. berthaultii hybrids.116 It may soon be possible to use in vitro callus from S. tuberosum to screen for insect resistance. This technique has been used to select somaclonal variants of tomatoes, Lycopersicon esculentum Mill., with resistance to P. operculella larvae.120 This strategy has the advantage of separating chemical resistance factors from morphological resistance factors and may be of use in studying newly discovered potato alkaloids, e.g., calystegines.83 Interspecific incompatibility can occur at either pre- or post-zygotic levels. Prezygotic barriers have been overcome by application of double crosses with bridge species compatible with each of the incompatible species.35,52,53 Post-zygotic incompatibility, caused by degeneration of endosperm, can be predicted using Endosperm Balance Number theory.70 Crosses that result in endosperm with a ratio other than 2:1 (female parent chromosomes: male parent chromosomes) will be unsuccessful. Post-zygotic incompatibility barriers can be overcome by in vitro culture of ovaries and embryos from immature berries harvested just days after pollination,68,113 use of triplandroids,19 or use of somatic hybridization by symmetric or asymmetric fusion of protoplasts.50,80,99,118 Somatic fusion has been used to transfer the following traits into cultivated potato lines: resistance to fungal pathogens Phytophthora infestans [Mont.] de Bary, Verticillium dahliae Kleb, and Fusarium oxysporum Schlecht; bacterial pathogens Erwinia carotovora (Jones) Bergey et al. and Pseudomonas solanacearum E. F. Smith; the nematodes Globodera rostochinensis (Wollen. 1972) Beherens 1975 and G. pallida (Stone 1972) Beherens 1975; and tolerance to frost.

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Development of isoenzymatic33 and molecular markers63,121 has greatly increased our knowledge of potato genetics. Markers allow selection of useful traits without actual observation of phenotypic expression. Therefore, marker assisted backcrossing is faster and requires screening fewer progeny than conventional backcrossing schemes. Markers also facilitate identification of potato cultivars36 and of potato hybrids that produce 2N gametes.12 Two genetic maps of potato have been developed.17,43,44,65 Quantitative Trait Loci (QTL) contributing to trichome-mediated insect resistance have been mapped for S. tuberosum x S. berthaultii hybrid progenies.16,138 Genes for resistance to G. rostochiensis, P. infestans, and potato virus X have been mapped.11,42,44,100 Molecular markers have been identified for resistance to the nematodes G. rostochiensis and G. pallida in F1 progeny of S. tuberosum x S. spegazzini Bitt., and resistance to P. infestans in F1 populations from crosses S. microdontum Bitt. x S. tuberosum.104 Transgenic cultivars, while outside the scope of this chapter, are alternatives to traditional plant breeding. Transgenic potatoes have been developed with resistance to potato pests, e.g., transgenic plants with lepidopteran- and coleopteran-active proteins from Bacillus thuringiensis Berliner.38

IV.

PEST RESISTANCE IN EXOTIC POTATOES

There are many instances of insect resistance in wild potatoes.9,39,40,97,119,125,126 For example, accessions highly resistant to M. persicae were identified within 36 potato species, to the potato aphid, Macrosiphum euphorbiae (Thomas), within 24 species, to L. decemlinata within 10 species, to potato flea beetle, Epitrix cucumeris (Harris), within 25 species, and to potato leafhopper, Empoasca fabae (Harris), within 39 species.39 Two studies of germplasm from the NRSP-6 collection84 indicate evolutionary and geographical associations with insect resistance.39,40 Insect resistance appeared to be a primitive trait in wild potatoes. Insect resistance was also characteristic of the most advanced species. Susceptibility was most common in the primitive and cultivated Tuberosa. The glycoalkaloid tomatine was associated with resistance to L. decemlinata and E. fabae while other glycoalkaloids did not cause resistance at the species level. Dense hairs and glandular trichomes have been associated with resistance to M. persicae, E. cucumeris, and E. fabae and to L. decemlinata, E. cucumeris, and E. fabae, respectively. Species from hot and arid areas were resistant to L. decemlinata, E. cucumeris, and E. fabae, while species from cool or moist areas tended to be resistant to M. euphorbiae. There is evidence potatoes from certain germplasm areas are resistant to some of our more common potato pests more frequently than would be expected.40 The Mexico-U.S. potatoes as a group exhibited resistance to all insects except L. decemlineata. Germplasm from Colombia and Ecuador were susceptible to L. decemlineata, M. euphorbiae, E. cucumeris, and E. fabae while material from Peru was resistant to M. persicae. Potatoes from Bolivia were susceptible to M. euphorbiae, but resistant to L. decemlineata, E. cucumeris, and E. fabae. Material from Argentina tended to be resistant to L. decemlineata and M. persicae, but susceptible to M. euphorbiae and E. fabae. North America potatoes collected at or below 2,500 m were resistant to M. persicae, L. decemlineata and E. cucumeris, while potatoes from South America at elevations © 1999 by CRC Press LLC

greater than 3,000 m were resistant to M. persicae and M. euphorbiae. Additionally, this group of potatoes collected at or below 3,000 m were resistant to L. decemlineata, E. cucumeris, and E. fabae. In this review, emphasis is placed on resistance in the wild potatoes to three of the most important insect pests of potato: P. operculella, M. persicae, and L. decemlineata (Table 1). Information on resistances to other insects, pathogens, and nematodes, as well as to stress tolerance, is included, particularly for insect-resistant species that have not been used in potato breeding programs. Reported resistances may refer to all screened accessions of a potato species or to a particular accession within that species, depending upon the source of the information. Furthermore, definitions of resistance and susceptibility are subjective and criteria used to discriminate may differ among researchers. Resistance ratings for P. operculella are from studies at the International Potato Center,144 unless otherwise noted. Two interpretations of “resistance” to M. persicae and L. decemlineata are provided (Table 1). The first is from the NRSP-6 database, representing a synthesis of results from trials conducted by various workers,9 while the second is based on Minnesota screening data.39 The latter interpretation is liberal and includes moderately resistant accessions. Many wild species have been identified as potential sources of insect resistance but are not presently being used for potato germplasm enhancement (Table 1). These species are: S. acroglossum Juz., S. boliviense Dun., S. brachistotrichum (Bitt.) Rydb., S. bulbocastanum Dun., S. canasense Hawkes, S. capsicibaccatum Cárdenas, S. cardiophyllum Lindl., S. chomatophilum Bitt., S. etuberosum Lindl., S. gourlayi Hawkes, S. jamesii Torr., S. lignicaule Vargas, S. marinasense Vargas, S. medians Bitt., S. papita Rydb., S. polyadenium Greenm., S. trifidum Corr., and S. verrucosum Schlechtd. with resistance to M. persicae; S. acroglossum, S. alandiae Cárd., S. blanco-galdosii Ochoa, S. bulbocastanum, S. canasense, S. capsicibaccatum, S. chomatophilum, S. circaeifolium Bitt., S. gourlayi, S. immite Dun., S. jamesii, S. lignicaule, S. marinasense, S. mochiquense Ochoa, S. neocardenasii Hawkes et Hjerting, S. neorossii Hawkes et Hjerting, S. okadae Hawkes et Hjerting, S. oplocense Hawkes, S. papita, S. pinnatisectum Dun., S. stoloniferum Schlechtd. et Bché., S. tarijense Hawkes, and S. trifidum with resistance to L. decemlineata; and S. coelestipetalum Vargas, S. medians, S. multiinterruptum Bitt., and S. pinnatisectum with resistance to P. operculella. The diversity of the wild potatoes is still being explored for insect resistance. In the NRSP-6 Collection, there are 146 potato species with 97 species screened for M. persicae resistance and 104 screened for L. decemlineata resistance, while 28 species have not been screened for resistance to M. persicae, L. decemlineata, or P. operculella. These wild potato species are S. achacachense Cárd., S. albornozii Corr., S. arnezii Cárd, S. astleyi Hawkes et Hjerting, S. buesii Vargas, S. cacetanum Ochoa, S. chilliasense Ochoa, S. contumazaense Ochoa, S. flahaultii Bitt., S. garciabarrigae Ochoa, S. hastiforme Corr., S. hoopesii Hawkes et Okada, S. hypacrarthrum Bitt., S. irosinum Ochoa, S. lobbianum Bitt., S. matehualae Hjerting et Tarn, S. minutifolium Corr., S. nayaritense (Bitt.) Rydb., S. orocense Ochoa, S. otites Dun., S. pamplonense L. López, S. paramoense Bitt., S. rechei Hawkes et Hjerting, S. solisii Hawkes, S. subpanduratum Ochoa, S. sucubunense Ochoa, © 1999 by CRC Press LLC

TABLE 1 Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa S. abancayense N=24, EBN=2, Series Tuberosa S. acaule Bitt. N=48 (363) 72 (2) 96 (1), EBN = 2, Series Acaulia

S. acroglossum N=24, EBN= ?, Series Piurana S. acroscopicum N=24, EBN= ?, Series Tuberosa

S. alandiae N=24, EBN= ?, Series Tuberosa

S. albicans (Ochoa) Ochoa N=72, EBN=4, Series Acaulia

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Resistance statusb

Resistance trait Myzus persicae Leptinotarsa decemlineata M. persicae L. decemlineata Phthorimaea operculella frost, PVX, PLRV, nematodes heat Phytophthora infestans Rhizoctonia sp. M. persicae L. decemlineata P. operculella M. persicae L. decemlineata Meloidogyne incognita (Kofoid and White, 1919) Chitwood, 1949 L. decemlineata P. operculella PVX, PVY, Tobacco rattle virus (TRV) M. persicae L. decemlineata P. operculella Empoasca fabae frost

9

Use of germplasm, comments

39

0/3, 1/1 0/2,9 0/139 0/86,9 11/7839 5/33,9 2/1039 S (Cisneros, p.c.) R8 T8 R87 R85 1/1,9 1/139 1/1,9 1/139 S (Cisneros, p.c.) 0/2,9 1/339 1/2,9 2/239 R69

5/7,9 1/339 S (Cisneros, p.c.) R55 0/6,9 0/339 0/6,9 1/339 S (Cisneros, p.c.) R39 R47

Frost resistance, and in some cases, potato virus X resistance in 39 European,51 and 17 North American cultivars.94 Breeding techniques: S. commersonii bridge (V. dahliae);8 embryo rescue (PLRV, potato spindle tuber viroid (PSTV)),133 triplandroid bridge;19 2N pollen (Rhizoctonia);85 2N eggs.24 Crossing ability with other wild species and S. tuberosum,71 e.g., transfer traits from S. etuberosum.27

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa S. andreanum N=24, EBN= ?, Series Tuberosa S. avilesii Hawkes et Hjerting N=24, EBN= ?, Series Tuberosa S. berthaultii N=24, EBN = 2, Series Tuberosa

M. persicae L. decemlineata Epitrix cucumeris L. decemlineata

1/139 0/3,9 2/239 R39 1/29

M. persicae L. decemlineata

2/13,9 3/939 27/37,9 8/8,39 9/9 (Arnone, unpubl.), R97,98 R15

P. operculella Trialeurodes vaporariorum (Westwood) E. cucumeris, E. fabae P. infestans Fusarium solani, Verticillium dahliae

S. blanco-galdosii N=24, EBN= ?, Series Piurana S. boliviense N=24, EBN=2, Series Megistacroloba S. brachistotrichum N=24, EBN=1, Series Pinnatisecta

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Resistance statusb

Resistance trait

M. persicae L. decemlineata E. cucumeris M. persicae L. decemlineata Tetranychus urticae Koch M. persicae L. decemlineata

R39 R142 R79

1/1,9 1/139 2/29 R39 1/5,9 5/639 1/5,9 0/539 S (Tingey, p.c.) 8/8,9 7/7,39 R106 0/10,9 2/839

Use of germplasm, comments

Glandular trichome-based trait of S. berthaultii incorporated into NYL 235-4, a named, advanced breeding line95 with L. decemlineata and E. fabae resistance. The development of this and closely related lines of the “hairy potato” is well described by Tingey and Yencho.125 NYL 235-4 is resistant to Ostrinia nubilalis (Hübner), Liriomyza huidobrensis (Blanchard) in Chile, and Diabrotica balteata LeConte in Brazil (Tingey, p.c.), E. cucumeris, E. fabae, potato virus X, and Globodera rostochiensis.95 S. berthaultii used for P. operculella resistance in breeding programs.82,97 Techniques: protoplast electrofusion, somaclonal variation in callus culture, and S. tuberosum haploid crosses.37,54,67,77,110

Frost resistance in S. tuberosum breeding lines,103 crossed with haploid S. tuberosum,54 crossed with S. commersonii.131

S. brachycarpum Corr. N=72, EBN=4, Series Demissa

S. brevidens N=24, EBN=1, Series Etuberosa

S. bukasovii N=24, EBN=2, Series Tuberosa S. bulbocastanum N=24, EBN=1, Series Bulbocastana

S. cajamarquense Ochoa N=24, EBN= ?, Series Tuberosa

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M. persicae L. decemlineata E. fabae PLRV PVX, PVY Meloidogyne hapla Chitwood, 1949 M. persicae L. decemlineata E. fabae PVY PLRV Andean potato mottle virus (APMV), PVX, PVA, PVM, and PVS M. persicae L. decemlineata P. operculella M. persicae L. decemlineata P. operculella E. cucumeris M. hapla Meloidogyne chitwoodi Golden, O’Bannon, Santo, and Finley, 1980 F. solani P. infestans M. persicae

0/5,9 0/539 1/7,9 0/539 R39,106 R45 R128 R59

E. fabae resistance associated with high glycoalkaloid content.106

1/2,9 2/239 0/4,9 1/339 R39 R56,129 R5,46,129 R129

Frost resistance used in breeding programs.103 Gene intogression via embryo rescue and diploid breeding theory.70 Utilization of the virus resistance in this species possible with advent of protoplast fusion techniques.5,13,46,101 High levels of tuber glycoalkaloids a potential drawback (Tingey, p.c.).

7/25,9 11/1639 2/19,9 1/1239 S (Cisneros, p.c.) 26/33,9 31/3139 3/19,9 8/1739 S (Cisneros, p.c.) R39 R59 R21

Present in pedigree of M. persicae-resistant breeding lines (Longtine, p.c.). Can be crossed with haploid S. tuberosum.139

R79 R52 2/239

Recently, fusion hybrids with S. tuberosum have been made with nematode resistance,6,21 and late blight resistance.108 Successful crosses of this species have been reported with S. acaule;71 with S. cardiophyllum using detached inflorescences;74 and with S. tuberosum using a double bridge hybrid for blight resistance.52

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa S. canasense N=24, EBN=2, Series Tuberosa S. capsicibaccatum N=24, EBN=1, Series Circaeifolia

S. cardiophyllum N=24, EBN=1, Series Pinnatisecta S. chacoense N=24, EBN=2, Series Yungasensa

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Resistance statusb

Resistance trait M. persicae L. decemlineata M. persicae L. decemlineata E. cucumeris Globodera pallida, P4A, P5A P. infestans Rhizoctonia sp. M. persicae L. decemlineata M. hapla M. persicae L. decemlineata V. dahliae Pseudomonas solanacearum Meloidogyne sp. heat

7/21,9 21/24,39 R122 6/19,9 2/1939 1/1,9 2/239 2/39,39 R39 R29 R142 R85 7/12,9 11/1639 1/10,9 2/939 R59 1/74,9 2/4539 88/99,9 24/2939 R79 R81 R81 T73

Use of germplasm, comments Frost resistance in S. tuberosum breeding lines103 can be crossed with haploid S. tuberosum.54,139 Cyst nematode resistance retained in crosses with S. tuberosum haploids.29 Can be regenerated from protoplasts, good potential for somatic hybridization.137

Field resistant, but susceptible in laboratory to L. decemlineata.126 Can be crossed with S. bulbocastanum using detached inflorescences,74 also with S. commersonii131 and S. acaule.71 One of the first species to be used in programs to breed insect resistant potatoes.126 Advanced breeding lines with L. decemlineata and E. fabae resistance have been developed.105-106 The biggest obstacle to commercialization of leptine-based L. decemlineata resistant breeding lines is that tubers tend to retain undesirable levels of traditional tuber glycoalkaloids (Tingey, p.c.). Excessive tuber glycoalkaloids has often been a limiting factor in S. chacoense based E. fabae resistant breeding lines, although low tuber glycoalkaloid lines have been developed.106 The cultivar “Lenape,”3 with S. chacoense in its pedigree was withdrawn from the market because of excessive tuber glycoalkaloids.141 In the pedigree of 17 North American potato cultivars,94 and at least 10 European cultivars.51

S. chancayense N=24, EBN=1, Series Tuberosa S. chomatophilum N=24, EBN=2, Series Conicibaccata

S. circaeifolium N=24, EBN=1, Series Circaeifolia

S. clarum Corr. N=24, EBN= ?, Series Bulbocastana S. coelestipetalum N=24, EBN= ?, Series Tuberosa S. commersonii N=24, EBN=1, Series Commersoniana

S. demissum N=72, EBN=4, Series Demissa

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M. persicae L. decemlineata E. fabae M. persicae L. decemlineata E. cucumeris

0/2,9 1/239 2/29,39 R39 7/11,9 7/1039 5/8,9 8/839 R39

Can be crossed with S. tuberosum using 2n pollen, or embryo rescue techniques.88,113

L. decemlineata P. operculella E. cucumeris PVX M. persicae L. decemlineata P. operculella

2/3,9 1/139 S (Cisneros, p.c.) R39 R55 1/19 1/39 R (Cisneros, p.c.)

P. infestans and G. pallida cyst nematode resistance used via somatic fusion.80

M. persicae L. decemlineata P. operculella frost

0/16,9 3/839 11/15,9 3/439,R126 R30,125 R103

M. persicae L. decemlineata P. operculella PVX, PVY

4/54,9 4/4939 38/92,9 2/7739} S (Cisneros, p.c.) R128

Source of L. decemlineata resistance in potato breeding lines.66 Crossed with S. capsicibaccatum and S. lignicaule, but less than 2% of clones maintained resistance to P. operculella.30 Synthetic tetraploids used as a bridge from species in the series Acaulia and Longipedicellata to S. tuberosum.8 Somatic fusion hybrids have been made.41 Frost resistance trait used in potato breeding lines.131 Resistance to P. infestans explains why S. demissum is in the pedigree of at least 335 European cultivars51 and in the pedigree of 58% of North American cultivars.94 L. decemlineata resistance transferred into advanced S. tuberosum-based breeding lines.106,126 Enhanced germplasm lines with L. decemlineata resistance have been made, with S. chacoense, S. verrucosum.126 Crossed with S. pinnatisectum using detached inflorescences,74 also can be crossed with S. commersonii.131

Frost resistance, crossing ability with other wild sp. to create enhanced germplasm, e.g., S. commersonii 131 frost resistance in S. tuberosum breeding lines.103

M. persicae resistance used using haploid/dihaploid crosses (Longtine, p.c).

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa S. doddsii Corr. N=24, EBN=2, Series Tuberosa S. x edinense Berth. N=60, EBN= ?, Series Demissa S. etuberosum N=24, EBN=1, Series Etuberosa

S. fendleri N=48, EBN=2, Series Longipedicellata

S. fernandezianum N=24, EBN=1, Series Etuberosa S. gourlayi N=24(82), 48(91), EBN=2,4, Series Tuberosa

S. hjertingii N=48, EBN=2, Series Longipedicellata

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Resistance trait L. decemlineata P. operculella

Resistance statusb

Use of germplasm, comments

0/2,9 1/239 S (Cisneros, p.c.) In pedigree of the late blight resistant W races.94

M. persicae L. decemlineata E. fabae APMV, PVX, PLRV, PVY, PVA M. persicae L. decemlineata P. operculella

2/2,9 3/339 1/3,9 2/239 R39 R129

Potato leafroll virus resistance used via S. pinnatisectum and S. acaule, bridging to S. phureja.27 PVY, PLRV, frost tolerance, can be crossed with S. pinnatisectum.53 Fusion hybrids have been made.89

1/16,9 4/1539 8/19,9 0/11,39 S126 S (Cisneros, p.c.)

In advanced breeding lines with L. decemlineata resistance (Lorenzen, p.c.). P. infestans resistance used via 2n pollen producing triploids.2 In Verticillium wilt-resistant breeding lines,8 and in pedigree of at least 9 cultivars.58,94 Other techniques: anther culture.112

L. decemlineata PVY PLRV, PVA M. persicae L. decemlineata M. incognita

2/29 R56,129 R129 1/38,9 15/2939 5/14,9 1/739 R69

G. pallida P4A, P5A resistance used via haploid S. tuberosum and 2n wild pollen.29 Years ago, this was used in breeding-S. gourlayi X S. phureja (Lauer, p.c.).

M. persicae L. decemlineata F. solani M. hapla V. dahliae

2/5,9 6/7,39 R143 0/39,39 R79 R59 R8

In the pedigree of M. persicae advanced breeding lines (Longtine, p.c.). P. infestans resistance and lack of enzymatic browning used through 2n pollen producing triploids.2 Solanum commersonii can be used as a bridge for this species.8 Potential for use via somatic hybridization.137

S. hondelmannii Hawkes et Hjerting N=24, EBN= ?, Series Tuberosa S. hougasii Corr. N=72, EBN=4, Series Demissa

S. immite N=24, EBN= ?, Series Tuberosa S. incahuasinum Ochoa N=24, EBN= ?, Series Tuberosa S. incamayoense Okada et Clausen N=24, EBN= ?, Series Tuberosa S. infundibuliforme N=24(109),48(1), EBN=2, Series Cuneoalata S. iopetalum (Bitt.) Hawkes N=72, EBN=4, Series Demissa

S. jalcae EBN= ?, Series Piurana S. jamesii N=24, EBN=1, Series Pinnatisecta

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M. persicae L. decemlineata

0/139 1/3,9 0/239

M. persicae L. decemlineata P. operculella M. hapla M. chitwoodi M. persicae L. decemlineata E. cucumeris P. operculella

1/2,9 1/439 0/39 S (Cisneros, p.c.) R59 R20 0/29,39 3/3,9 1/139 R39 S (Cisneros, p.c.)

M. persicae L. decemlineata M. persicae L. decemlineata M. hapla M. persicae L. decemlineata E. cucumeris F. solani P. infestans L. decemlineata M. persicae L. decemlineata E. cucumeris

0/19 1/6,9 0/439 10/50,9 24/3039 5/18,9 3/939 R59 0/29,39 0/2,9 1/239 R39 R79 R87 1/1,39 R125 3/9,9 8/10,39 R143 18/18,9 8/8,39 R126 R39

Crossed with S. gourlayi, S. vidaurrei, S. spegazzinii, and S. infundibuliforme.91 Crossed with haploid S. tuberosum ssp. tuberosum.54,139 Can be crossed with S. acaule.71 Used in insect-resistant breeding lines.94

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa

Resistance trait

Resistance statusb

S. kurtzianum Bitt. et Wittm. N=24, EBN=2, Series Tuberosa

M. persicae L. decemlineata E. fabae M. hapla

0/42,9 1/1839 0/41,9 1/339 R106 R59

S. leptophyes Bitt. N=24, EBN=2, Series Tuberosa

M. persicae L. decemlineata

0/4,9 3/1139 0/3,9 0/439

S. lignicaule N=24, EBN=1, Series Lignicaulia

M. persicae L. decemlineata P. operculella G. pallida, P4A, P5A M. persicae L. decemlineata M. persicae L. decemlineata E. fabae M. persicae L. decemlineata P. operculella E. cucumeris M. hapla PVY, PVX

1/3,9 2/339 2/4,9 2/339 S (Cisneros, p.c.) R29 0/29 1/29 0/139 2/29,39 R39 5/7,9 6/839 5/8,9 4/539 S (Cisneros, p.c.) R39 R59 R56

S. longiconicum Bitt. N=48, EBN= ?, Series Conicibaccata S. lycopersicoides N=24, EBN= ?, Series Juglandifolia S. marinasense N=24, EBN=2, Series Tuberosa

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Use of germplasm, comments In the pedigree of advanced hybrids (Longtine, p.c.). Rhizoctonia resistance transferred to a S. tuberosum breeding line using 2n pollen and diploid S. tuberosum germplasm.85 Can be crossed with S. acaule, then with S. tuberosum.71 Present in the pedigree of “Conestoga.” 94 M. persicae resistance used with haploid/dihaploid crosses (Longtine, p.c.), crossing ability of wild-S. acaule hybrid with S. tuberosum,71 cyst nematode (G. pallida P4A, P5A) resistance in S. tuberosum diploids via 2n pollen.29

Poor prospect for use in 2X or 4X breeding lines, doesn’t hybridize in nature, needs protoplast fusion (Hanneman, p.c.).

S. medians N=24, EBN=2, Series Tuberosa

S. megistacrolobum N=24, EBN=2, Series Megistacroloba

S. michoacanum (Bitt.) Rydb. N=24, EBN= ?, Series Pinnatisecta S. microdontum N-24, EBN=2, Series Tuberosa

S. mochiquense N=24, EBN=1, Series Tuberosa

S. morelliforme Bitt. et Muench N=24, EBN= ?, Series Morelliformia

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M. persicae L. decemlineata P. operculella E. fabae M. persicae L. decemlineata P. operculella frost M. incognita M. persicae

1/8,9 3/839 1/8,9 0/839 R (Cisneros, p.c.) R106 6/48,9 23/3939 2/27,9 2/1839 S (Cisneros, p.c.) R103 R69 R143

High glycoalkaloid content associated with E. fabae resistance.106 Used in breeding lines for tolerance to environmental stress.94

M. persicae L. decemlineata P. operculella V. dahliae Rhizoctonia spp. PLRV P. infestans P. solanacearum M. persicae L. decemlineata E. cucumeris alfalfa mosaic virus (AMV), PVX L. decemlineata P. operculella

0/47,9 2/3539 1/19,9 1/739 S (Cisneros, p.c.) R79 R85 R45 R87,142 R81 0/3,9 0/439 3/3,9 2/339 R39 R55

Produces 2n pollen, which makes it possible to cross this species with S. tuberosum.54 The species has good frost resistance, and can be crossed with S. acaule, and the resulting hybrid with S. phureja.71 Fusion hybrids have also been created.99 In the pedigree of “Conestoga”94 and two European cultivars.51

1/49 S (Cisneros, p.c.)

M. persicae-resistant trait is being used in Minnesota breeding lines (Longtine, p.c.). Crosses with S. acaule71 and S. commersonii.131 Frost resistance,103 used in potato breeding programs.

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa S. multidissectum N=24, EBN=2, Series Tuberosa S. multiinterruptum N=24, EBN=2, Series Tuberosa

S. neocardenasii N=24, EBN= ?, Series Tuberosa

S. neorossii N=24, EBN = ?, Series Tuberosa S. ochranthum N=24, EBN= ?, Series Juglandifolia S. okadae N=24, EBN= ?, Series Tuberosa

S. oplocense N=48(11), 72(41), EBN=4, Series Tuberosa

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Resistance statusb

Resistance trait M. persicae L. decemlineata M. hapla M. persicae L. decemlineata P. operculella M. hapla M. persicae L. decemlineata E. fabae PVX PVY AMV L. decemlineata AMV, PVX M. persicae L. decemlineata E. fabae M. persicae L. decemlineata P. operculella G. pallida M. persicae L. decemlineata V. dahliae PVX P. infestans

9

39

1/7, 5/6 3/8,9 0/239 R59 1/5,9 2/539 1/3,9 0/439 R (Cisneros, p.c.) R59 R75 R34,111,125 R39 R128 R55,128 R55 3/5,9 0/539 R55 0/139 0/2,9 0/1,39 R126 R (Tingey, p.c.) 0/2,9 1/439 3/49,39 S (Cisneros, p.c.) R114 0/23,9 1/2239 6/13,9 4/739 R79 R127 R87

Use of germplasm, comments Traits from this species have been incorporated into 4X S. tuberosum breeding lines: M. persicae resistance,106 nematode resistance94 and frost resistance.103

Disadvantages: produces a very poor type vine (Sanford, p.c.), 2X by 4X crosses and protoplast fusion have failed, and tuber glycoalkaloids a problem (Tingey, p.c.).

Poor prospect for use in 2X or 4X breeding lines, doesn’t hybridize in nature, needs protoplast fusion (Hanneman, p.c.). Used as a parent for cold storage chipping ability (Hoopes, p.c.).

S. orophilum Corr. N=-, EBN= ?, Series Tuberosa S. pampasense Hawkes N=24, EBN=2, Series Tuberosa

S. papita N=48, EBN=2, Series Longipedicellata S. paucijugum Bitt. N=48, EBN= ?, Series Conicibaccata S. paucissectum Ochoa N=24, EBN=2, Series Piurana S. phureja N=24, EBN=2, Series Tuberosa

S. pinnatisectum N=24, EBN=1, Series Pinnatisecta

S. piurae Bitt. N=24, EBN= ?, Series Piurana

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E. fabae

R (Tingey, p.c.)

M. persicae L. decemlineata E. fabae E. cucumeris V. dahliae P. infestans M. persicae L. decemlineata M. hapla E. fabae

0/4,9 1/439 1/3,9 0/139 R72 R39 R79 R23 0/8,9 5/1439 3/6,9 1/339 R59 R (Tingey,p.c.)

M. persicae L. decemlineata PVX, TRV, AMV M. persicae L. decemlineata PVY P. solanacearum Meloidogyne sp. heat M. persicae L. decemlineata

0/29 1/29 R55 0/66,9 18/7039 0/24,9 0/2039 R (Backlund, p.c.) R73,81 R81 T73 1/10,9 1/1139 14/15,9 10/10,39 3/3 (Arnone, unpubl.) R4,30 R39 R53 0/2,9 1/239 0/19,39 R39

P. operculella E. cucumeris frost, PVY, PLRV M. persicae L. decemlineata E. cucumeris

In pedigree of M. persicae-resistant breeding lines (Longtine, p.c.) and in P. infestans-resistant breeding lines.23

Verticillium wilt resistance can be used using S. commersonii bridge.8 Can be crossed with 2n pollen producing triploids.2

Has been used in the development of cultivated, primitive, 2X populations because of desirable traits, and suitability for the highland and lowland tropics.81,135 Selections produce as much as 80% big pollen making S. phureja a useful bridge species.2,10 Present in the pedigree of 11 North American cultivars,94 and in 27 European cultivars.51 Used as a bridge to transfer resistant germplasm from S. etuberosum.27 Also crossed with S. demissum using detached inflorescences,74 and S. pinnatisectum-S. acaule hybrids have been crossed with S. phureja.71 Fusion hybrids incorporate P. infestans resistance.108 It has been used in somatic hybrids.107 Tuber glycoalkaloids may be a problem (Tingey, p.c.).

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa S. polyadenium N=24, EBN= ?, Series Polyadenia

S. polytrichon Rydb. N=48, EBN=2, Series Longipedicellata S. raphanifolium N=24, EBN=2, Series Megistacroloba

S. x sambucinum Rydb. N=24, EBN= ?, Series Pinnatisecta S. sanctae-rosae N=24, EBN=2, Series Megistacroloba S. santolallae N=24, EBN= ?, Series Conicibaccata S. schenckii Bitt. N=72, EBN=4, Series Demissa S. sparsipilum N=24, EBN=2, Series Tuberosa

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Resistance trait

Resistance statusb 9

39

M. persicae L. decemlineata E. fabae E. cucumeris V. dahliae M. persicae L. decemlineata E. cucumeris M. persicae L. decemlineata P. operculella P. solanacearum, heat M. persicae E. fabae M. persicae L. decemlineata

2/14, 3/13 16/16,9 9/9,39 R126 R39,106 R39 R79 1/16,9 2/1139 1/14,9 1/539 R39 0/26,9 2/2339 4/10,9 0/139 R (Cisneros, p.c.) R73 0/139 R39 6/9,9 6/8,39 R106,122 0/10,9 1/839

M. persicae L. decemlineata M. persicae L. decemlineata M. persicae L. decemlineata P. operculella Meloidogyne sp. G. pallida P. solanacearum

0/1,9 1/139 0/29,39 0/139 2/5,9 1/339 0/279,39 0/12,9 0/739 R4,30 (Cisneros p.c.) R59,69,81 R28 R81

Use of germplasm, comments Resistance to L. decemlineata and E. fabae appears to be based on tomatine, a highly toxic glycoalkaloid, and tuber glycoalkaloids are likely to be a problem (Tingey, p.c.).106 This species can be regenerated from protoplasts, so there is good potential for use via somatic hybridization.137 Present in the pedigree of M. persicae-resistant breeding lines (Longtine, p.c.).106 Late blight resistance used in crosses with S. tuberosum.2 Can be crossed with S. acaule.71 Present in the pedigree of M. persicae-resistant breeding lines (Longtine, p.c.). Frost resistant trait used in potato breeding programs.103 Used as cold chipping parent in breeding lines (Hoopes, p.c.). Present in the pedigree of cultivar “Krantz.”76

Crossed with S. acaule71 and S. commersonii.131 Good frost resistance103 has been used in breeding programs. Present in the pedigree of M. persicae-resistant S. tuberosum breeding lines (Longtine, p.c.).

Crossed with S. tuberosum haploids to create P. operculella resistant breeding lines.4,30,92 Present in the pedigree of 2 European cultivars.51 Root knot nematode resistance used in potato breeding lines.81,94

S. spegazzinii N=24, EBN=2, Series Tuberosa

S. stenophyllidium Bitt. N=24, EBN= ?, Series Pinnatisecta S. stenotomum N=24, EBN=2, Series Tuberosa S. stoloniferum N=48, EBN=2, Series Longipedicellata

S. sucrense N=48, EBN=4, Series Tuberosa S. tarijense N=24, EBN=2, Series Yungasensa

S. toralapanum N=24, EBN=2, Series Megistacroloba

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M. persicae L. decemlineata P. operculella frost PLRV Rhizoctonia sp. M. hapla G. pallida, G. rostochiensis M. persicae

2/27,9 8/1739 2/25,9 1/1039 S4 R71,103 R45 R85 R59 R51 R106

Present in the pedigree of M. persicae-resistant breeding lines in Minnesota (Longtine, p.c.). Other desirable horticultural traits from this species transferred to S. tuberosum using haploid/dihaploid crosses.139 G. rostochiensis & G. pallida resistant genes from this species used in 11 European cultivars.51 Present in the pedigree of “Krantz.”76

M. persicae L. decemlineata V. dahliae M. persicae L. decemlineata PVY P. infestans V. dahliae M. hapla M. persicae L. decemlineata P. operculella M. persicae L. decemlineata P. operculella E. cucumeris M. hapla V. dahliae M. persicae L. decemlineata frost

0/15,9 4/1639 0/14,9 0/939 R79 11/40,9 32/40,39 R106 13/46,9 5/2639 R2,8,51,78 R2 R8 R59 0/3,9 0/1039 5/15,9 2/839 R,30 S (Cisneros p.c.) 17/36,9 23/3239 43/48,9 17/1939 R30,125 R39 R59 R79 2/5,9 11/1239 2/10,9 0/739 R71

Tolerance to abiotic stress and disease used in the development of enhanced, cultivated 2N populations.81,135 Present in the pedigree of “Conestoga.”94 In the pedigree of M. persicae-resistant breeding lines at the University of Minnesota (Longtine, p.c.). In the pedigree of 41 PVA and PVY resistant European cultivars.51 Complex hybrids of this species can be created with S. acaule and S. tuberosum,71 as well as with S. commersonii.8

Used in Europe as a source of PVY resistance (Lauer, p.c.).

P. operculella resistance transferred to S. tuberosum ssp. andigena lines.30,125 In the pedigree of non-S. tuberosum-based M. persicae-resistant breeding lines (Longtine, p.c.). P. operculella resistance lost in crosses with haploid S. tuberosum.30 Germplasm from this species can be transferred using S. tuberosum haploids.54

In pedigree of M. persicae-resistant breeding lines via haploid/dihaploid crosses (Longtine, p.c.). Present in pedigree of cv. “Conestoga.”94 Can be crossed with S. acaule,71 S. commersonii, and S. phureja.131

TABLE 1 (continued) Occurrence and Use of Insect-Resistant Germplasm in Potato Breeding Programs Speciesa

Resistance statusb

Resistance trait M. persicae L. decemlineata E. cucumeris PVY M. persicae L. decemlineata see comments for other traits

9/9,9 10/10,39 R,106 R143 9/11,9 10/1139 R39 R56 0/185,9 4/21139 4/39,9 0/2839

S. tuberosum ssp. tuberosum N=48, EBN=4, Series Tuberosa

M. persicae L. decemlineata

1/6,9 1/5339 0/139

S. tuquerrense Hawkes N=48, EBN=2, Series Piurana

M. persicae L. decemlineata

1/29 0/29

S. trifidum N=24, EBN=1, Series Pinnatisecta

S. tuberosum ssp. andigena N=48, EBN=4, Series Tuberosa

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Use of germplasm, comments Crossing ability with S. acaule.71

Long been cultivated in South and Central America. Hundreds, if not thousands, of locally named cultivars exist. Little inherent pest resistance, but extensive screening, particularly at CIP, has found accessions resistant to M. persicae, P. solanacearum, A. solani, PVY, PVX, PLRV, Synchytrium endobioticum (Schilb.) Perc., P. infestans, Verticillium wilt, Streptomyces scabies (Thaxt.) Waksman & Henrici, G. pallida and G. rostochiensis, and/or Meloidogyne sp.18,81,134 Resistance to G. rostochiensis used in 298 European cultivars.51 The potato of commerce. Has little inherent resistance to most pests. However, named cultivars “Ulster Tarn” and “Record” have M. persicae resistance.106 “Greta,” “Alpha,” and “Cruza 148” have P. solanacearum resistance.73 Recurrent selection of a S. tuberosum population led to moderate E. fabae resistance.106 The cultivars “Alpha (A),” “Alpha (B),” and “Alpha (E)” also have E. fabae resistance.1

S. x vallis-mexici N=36, EBN= ?, Series Longipedicellata S. vernei Bitt. N=24, EBN=2, Series Tuberosa

S. verrucosum N=24, EBN=2, Series Tuberosa

S. wittmackii Bitt. N=24, EBN= ?, Series Tuberosa

M. persicae L. decemlineata M. persicae L. decemlineata Globodera sp. V. dahliae P. infestans frost M. persicae L. decemlineata P. infestans

1/139 S126 0/16,9 0/1039 1/10,9 0/739 R18,28 R79 R87 R71 1/16,9 13/1839 0/8,9 0/239 R23

L. decemlineata

S126

Present in pedigree of S. endobioticum resistant “Pushkin,”132 “Gibridnyi 14.”86 The cyst nematode resistance trait from this species has been incorporated into “Hampton,”94 and 41 European cultivars.51 Produces 2n pollen which makes it possible to cross this species fairly easily with haploid S. tuberosum.139,140 It can also be crossed with S. chomatophilum,131 and S. acaule.71 Fusion hybrids have been created.99 One of first species to be crossed with S. etuberosum and S. pinnatisectum through use of colchicine.53 Potential use as a bridge species to haploid S. tuberosum or to S. phureja.131,140 In the pedigree of two European cultivars.51

Wild potato species (no. accessions screened) that have not shown resistance to M. persicae and/or L. decemlineata in screening trials:9,39 S. agrimonifolium Rydberg (6), S. ambosinum Ochoa (4), S. brevicaule (Bitt.) (2), S. candolleanum Berth. (3), S. chaucha (1), S. chiquidenum Ochoa (3), S. colombianum Ochoa (3), S. curtilobum Juz. et Buk. (5), S. dolichocremastrum Bitt. (1), S. gandarillasii Cárd. (3), S. guerreroense Corr. (2), S. huancabambense Ochoa (4), S. laxissimum Bitt. (1), S. lesteri Hawkes et Hjerting (1), S. maglia Schlechdt. (1), S. moscopanum Hawkes (2), S. oxycarpum Schiede (2), S. pascoense Ochoa (1), S. scabrifolium Ochoa (1), S. sitiens Johnston (1), S. sogarandinum Ochoa (2), S. venturii Hawkes et Hjerting (1), S. violaceimarmoratum Bitt. (4), S. weberbaueri Bitt. (2). Wild potato species that did not show resistance to P. operculella in screening trials (F. Cisneros, personal communication): S. ambosinum, S. candolleanum, S. huancabambense, S. huarochirense Ochoa, S. limbaniense, S. sogarandinum, S. virgultorum (Bitt.) Cárd et Hawkes. a b

N = chromosome number, EBN = Endosperm Balance Number.8,49 R = reported as resistant, S = reported as susceptible,T = reported as tolerant, x/y = no. resistant accessions/total accessions screened, p.c. = personal communication.

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FIGURE 1 Utilization of potato species in plant breeding programs. Wild = wild potato species screened for insect resistance; improved = potato species used in non-S. tuberosum breeding lines; 2X tbr br. line = potato species used in diploid S. tuberosum breeding lines; 4X tbr br. line = potato species used in tetraploid S. tuberosum breeding lines; commercial cultivar/clone = potato species used in a named cultivar or a widely released clone. R = resistant species, S = susceptible species.

S. tarnii Hawkes et Hjerting, and S. ugentii Hawkes et Hjerting. Thirty-six species have been screened at the International Potato Center for P. operculella resistance. It is hoped that plant collectors will continue to expand the world’s potato germplasm resource because not all potato species are represented in genebanks.

V.

USE OF RESISTANT GERMPLASM IN BREEDING

Despite years of breeding, there are as yet no temperate-zone potato cultivars with genes for insect resistance from exotic potatoes (Figure 1). However, at least 20 species have been, or are currently being used, as sources of M. persicae resistance in potato breeding programs (Table 1). At least 12 potato species have been used as sources of L. decemlineata resistance in breeding programs (Table 1). It is likely

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that cultivars will soon be developed that are based on the advanced insect resistant breeding line NYL-235-4 which has S. berthaultii in its pedigree.95 Advanced L. decemlineata resistant lines have been developed with S. fendleri Asa Gray in their pedigrees.145 At least six potato species have been used as sources of P. operculella resistance. Plant breeders, geneticists, entomologists, and plant pathologists (see Acknowledgments) who work with Section Petota were asked for opinions on prospects for use of particular wild potato species in breeding lines and in commercial cultivars. Potato species that have contributed to the parentage of advanced tetraploid or diploid breeding lines were given the greatest prospect of being sources of insect resistance in commercial cultivars in the near future. These potato species are S. berthaultii, S. bukasovii Juz, S. commersonii Dun., S. demissum Lindl., S. fendleri, S. hjertingii Hawkes, S. megistacrolobum Bitt., S. microdontum, S. multidissectum Hawkes, S. raphanifolium Cárd. et Hawkes, S. sanctae-roae Hawkes, S. sparsipilum, S. spegazzinii, S. stenotomum, S. stoloniferum, S. x sucrense, S. tarijense, S. toralapanum Cárd et Hawkes, and S. vernei Bitt. et Wittm. The following potato species were mentioned as good short-term (next 5 years) to mid-term (next 15 years) prospects for use in S. tuberosum-based breeding lines: S. abancayense Ochoa, S. acroglossum, S. acroscopicum Ochoa, S. alandiae, S. andreanum Baker, S. blancogaldosii, S. boliviense, S. bulbocastanum, S. canasense, S. cardiophyllum, S. chancayense Ochoa, S. chomatophilum, S. gourlayi, S. infundibuliforme, S. marinasense, S. medians, S. multiinterruptum, S. neorossii, S. okadae, S. oplocense, S. santolallae Vargas, S. trifidum, and S. x vallis-mexici Juz. Solanum lycopersicoides Dun. and S. ochranthum Dun. were rated as poor prospects for use because they are distantly related or do not hybridize easily. Wild potato species that seem to be good prospects, based on strong insect resistance, may be limited in their use because progeny contain excessive tuber glycoalkaloids, e.g., S. brevidens, S. chacoense Bitt., S. neocardenasii, S. pinnatisectum, and S. polyadenium. Plant breeders must select not only for resistance to insects but also for resistance to diseases, nematodes, and tolerance to environmental stresses. The breeding program at the International Potato Center81,134 is an excellent example. For the past 25 years, systematic germplasm enhancement at the diploid level has resulted in breeding lines with resistance to bacterial wilt Pseudomonas solanacearum, early blight Alternaria solani (Sorauer), cyst nematode Globodera spp., potato leafroll virus (PVX), potato virus X (PVX) and potato virus Y (PVY), root-knot nematode Meloidogyne spp., and soft-rot Erwinia carotovora.135 This germplasm improvement scheme also has been used to develop lines that are resistant to P. operculella and Liriomyza spp.97 These breeding lines started with cultivated potato germplasm such as S. tuberosum ssp. tuberosum, S. tuberosum ssp. andigena, S. phureja, or S. stenotomum. Resistance genes from wild species have been incorporated at the 2X level, which is much faster than at the 4X level. Breeding lines were developed in collaboration with North American universities and European research institutes to enlarge the genetic variability of cultivated potatoes and introduce resistance traits to biotic stresses.51 Enhanced breeding lines are then used to create commercial cultivars. This approach has led to release of at least 30 new cultivars with multiple pest (plant pathogen and nematode) resistance.81 © 1999 by CRC Press LLC

VI.

CONCLUSIONS

Solanum tuberosum is a tetraploid species, whereas many exotic species are diploid or hexaploid. Thus, many potential sources of insect resistance have been difficult to access using conventional breeding methods. Advances in potato breeding, such as the use of haploid S. tuberosum lines, occurrence of unreduced pollen in diploid species, use of bridge species, and somatic hybridization, have made it possible to exploit more and more wild potato species. Genes from at least 18 exotic potato species have been incorporated into North American and European potato cultivars primarily for stress tolerance or disease resistance. However, this represents less than 8% of the extraordinary potato species diversity that could potentially be exploited as resistance sources. There are still no commercial potatoes that were developed specifically for insect resistance. Advanced breeding lines with insect resistance exist. Prospects are good for the release of insect-resistant cultivars in the next five years and excellent for release within the next 15 years. It is the authors’ hope that this review will help researchers see where the greatest gaps in knowledge occur.

ACKNOWLEDGMENTS The following potato researchers generously contributed unpublished information on sources and utilization of insect-resistant potato germplasm: J. E. Backlund, University of Minnesota, St. Paul, Minnesota; C. R. Brown, USDA-ARS, Prosser, Washington; F. Cisneros, International Potato Center, Lima, Peru; R. E. Hanneman, Jr., University of Wisconsin, Madison, Wisconsin; K. G. Haynes, USDA-ARS, Beltsville, Maryland; R. W. Hoopes, Frito-Lay, Inc., Rhinelander, Wisconsin; S. H. Jansky, University of Wisconsin, Stevens Point, Wisconsin; F. I. Lauer and C. A. Longtine, University of Minnesota, St. Paul, Minnesota; J. H. Lorenzen, North Dakota State University, Fargo, North Dakota; R. L. Plaisted and W. M. Tingey, Cornell University, Ithaca, New York; L. L. Sanford, USDA-ARS, Beltsville, Maryland; R.E. Veilleux, VPI & SU, Blacksburg, Virginia; and G. C. Yencho, North Carolina State University, Plymouth, North Carolina. The authors thank J. Bamberg, M. Martin, and J. Schartner for providing information about accessions in the NRSP-6 collection.

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125. Tingey, W. M. and G. C. Yencho, Insect resistance in potato: a decade of progress, In: G. W. Zehnder, R. K. Jansson, M. Powelson, and K. V. Raman, Eds., Advances in Potato Pest Biology and Management, APS Press, St. Paul, 1992, 405. 126. Torka, M., Breeding potatoes with resistance to the Colorado beetle, Am. Potato J., 27:263-271, 1950. 127. Tozzini, A. C., M. F. Ceriani, M. V. Saladrigas, and H. E. Hopp, Extreme resistance to infection by potato virus X in genotypes of wild tuber-bearing Solanum species, Potato Res., 34:317-324, 1991. 128. Turuleva, L. M. and M. A. Balmasova, Wild potato species-sources of resistance to virus diseases, Selektsiya i biotekhnologiya kartofelya, 1990, 100. 129. Valkonen, J. P. T., G. Brigneti, L. F. Salazar, E. Pehu, and R. W. Gibson, Interactions of the Solanum spp. of the Etuberosa group and the nine potato-infecting viruses and a viroid, Ann. Appl. Biol., 120:301-313, 1992. 130. Vavilov Institute of Plant Industry Collection, on the World Wide Web (www.dainet.de/genres/vir/tuber.htm), 1997. 131. Vavilova, M. A., Use of wild frost-resistant species of the potato, S. commersonii Dun. and S. chomatophilum Bitt. in interspecific hybridization, In: A.-Ya Kameraz, Ed., Systematics, Breeding, and Seed Production of Potatoes, Oxonian Press Pvt. Ltd., New Dehli, 1985, 157 (translated from Russian). 132. Veselovskii, I., and S. Khusein, Some causes of incompatibility between potato species, Kartofel` i Ovoshchi, No. 4, 1973, 14. 133. Watanabe, K., C. Arbizu, and P. E. Schmiediche, Potato germplasm enhancement with disomic tetraploid Solanum acaule. I. Efficiency of introgression, Genome, 35:53-57, 1992. 134. Watanabe, K. N., M. Orrillo, and A. M. Golmirzaie, Potato germplasm enhancement for resistance to biotic stresses at CIP. Conventional and biotechnology-assisted approaches using a wide range of Solanum species, Euphytica, 85:457-464, 1995. 135. Watanabe, K., M. Orrillo, M. Iwanaga, R. Ortiz, R. Freyre, and S. Perez, Diploid potato germplasm derived from wild and land race genetic resources, Am. Potato J., 71:599-604, 1994. 136. Watanabe K. N., J. P. T. Valkonen, M. Orillo, and J. Pelttari, Production of sexual hybrids between tuber-bearing and non tuber-bearing potatoes using S. phureja for rescue pollination, In: 13th Triennial Conf. European Assoc. Potato Res., The European Association for Potato Research, Veldhoven, 1996, 49. 137. Xu, Y. S., E. Pehu, R. Malone, and M. G. K. Jones, Plant regeneration from protoplasts of Solanum species with potential agricultural value (S. hjertingii, S. polyadenium, S. capsicibaccatum), Plant Cell Rep., 9:520-522, 1991. 138. Yencho, G. C., M. W. Bonierbale, W. M. Tingey, R. L. Plaisted, and S. D. Tanksley, Molecular markers locate genes for resistance to the Colorado potato beetle, Leptinotarsa decemlineata, in hybrid Solanum tuberosum X S. berthaultii potato progenies, Entomol. Exp. Appl., 81:141-154, 1996. 139. Yerk, G. L. and S. J. Peloquin, Evaluation of tuber traits of 10, 2X(2EBN) wild species through haploid X wild species hybrids, Am. Potato J., 66:731-739, 1989. 140. Yerk, G. L. and S. J. Peloquin, Performance of haploid X wild species, 2X hybrids (involving five newly evaluated species) in 4X x 2X families, Am. Potato J., 67:405-417, 1990. 141. Zitnak, A. and G. R. Johnston, Glycoalkaloid content of B5121-6 potatoes, Am. Potato J., 47:256-261, 1970.

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142. Zoteeva, N. M. and L. M. Turuleva, Wild potato species-genetic sources of resistance to Phytophthora, Sbornik Nauchnykh Trudov po Prikladnoi Botanike, Genetike i Selektsii, 115:27-34, 1987. 143. Zykin, A. G. and E. D. Gerasenkova, Resistance of potato to aphid vectors of viruses, Doklady Vsesoyuznoi Ordena Lenina Akademii Sel'skokhozyaistvennykh Nauk., 1972, No. 4, 1972, 17. 144. Cisneros, F., Personal communication, 1996. 145. Lorenzen, J.H., Personal communication, 1996.

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Section V Basic Research and Biotechnology

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12

Plant Genetic Resources for the Study of Insect-Plant Interactions Sanford D. Eigenbrode and Stephen L. Clement

CONTENTS I. Introduction II. Plant Defenses Against Herbivory A. Physical Factors B. Secondary Chemicals 1. Photyhemagglutinin-arcelin-α-amylase Inhibitors in Phaseolus 2. Glandular Trichome Exudates III. Factors Mediating Host Selection IV. Plant Tolerance to Herbivory V. Multi-Trophic Interactions A. Endophytic Fungi B. Predators and Parasitoids 1. Plant Volatiles and Entomophagous Insects 2. Physical Factors 3. Surface Waxes 4. Glandular Trichomes VI. Genetics of Plant Traits Affecting Insects A. Inheritance B. Population Genetics and Geographic Variation VII. Germplasm Collections as Repositories for Living Vouchers VIII. Limitations of Genetic Resources for Research in Insect-Plant Interactions IX. Conclusions Acknowledgments References

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I.

INTRODUCTION

The development of plant resistance to insects and the study of the ecology and evolution of insect-plant interactions are connected conceptually and historically. Both fields of study are facilitated through knowledge of the mechanisms and the inheritance of defense or tolerance in plants to insect attack. Crop genotypes varying in susceptibility to insect attack have been important models in developing the field of insect-plant interactions.3,43,73 The consolidation of crop germplasm into national and international germplasm repositories facilitates its use in research in insect-plant interactions. The collections are extensive, centralized, in principle well documented and preserved, and easily accessed by researchers. Some of the nearly 6 million accessions of about 200 crop species in repositories worldwide36 are characterized for insect resistance (antibiosis, antixenosis, tolerance) or traits that can affect insect biology. More emphasis on collection of the wild and weedy relatives of crops, which are generally underrepresented in collections (Table 1), would improve genetic diversity and add value to repositories for research in insect-plant interactions. Accessions of wild species are valuable for addressing ecological and evolutionary questions because they have not been manipulated genetically by humans. An estimated 15% or about one million accessions in world collections are from wild species,36 representing thousands of plant species. For example, most of the 10,000 plant species in the U.S. National Plant Germplasm System (NPGS) are wild.49 This chapter focuses on the uses of conserved germplasm for the study of insectplant interactions. Examples presented herein show how these genetic resources have advanced our knowledge of the factors mediating insect-plant interactions. Expanded use of conserved germplasm by entomologists and ecologists has the potential to reveal more about the ecology and evolution of insect-plant interactions.

II.

PLANT DEFENSES AGAINST HERBIVORY

Increased knowledge of plant defense mechanisms will facilitate pest resistance breeding and expedite the incorporation of different defensive traits into commercial cultivars. Moreover, knowledge of defense mechanisms in natural systems provides for a better understanding of how plants are defended against insects and increased insights on the selective factors that shape plant defenses. The reader is referred to several reviews for in-depth discussions on plant characteristics that affect insect biology.7,8,97,107

A. PHYSICAL FACTORS Plant morphological traits such as tissue toughness, pubescence, and trichomes interfere with ingestion, oviposition, and colonization by insects.109 The insectdefensive value of these traits is illustrated by crop genotypes differing in leaf toughness,5 presence or densities of glandular and nonglandular simple and hooked trichomes,73,83 morphology and chemistry of surface waxes,28 and silica content.92 Crop germplasm collections are sometimes characterized for such traits, for example trichomes in accessions of Medicago (Chapter 8 in this volume), Triticum (Chapter 2 in this volume), Lycopersicon (Chapter 9 in this volume), and Cucurbita.49 © 1999 by CRC Press LLC

TABLE 1 Germplasm Holdings of Selected Crops and Their Wild Relativesa No. accessions (%)

Crop Rice (Oryza) Wheat (Triticum) Barley (Hordeum) Maize (Zea) Soybean (Glycine) Chickpea (Cicer) Cowpea (Vigna) Alfalfa (Medicago)c Tomato (Lycopersicon) Potato (Solanum) Tobacco (Nicotiana)

No. species

Repositoryb

Cultivars, breeding lines, and landraces

Wild and weedy species

Wild species and subspecies

Wild species and subspecies with >50 accessions

IRRI CIMMYT NPGS NPGS ICARDA NPGS NPGS NPGS IITA NPGS

77,869 (97) 94,356 (95) 41,580 (89) 30,046 (98) 22,422 (93) 19,604 (99) 18,587 (96) 4,444 (96) 14,816 (90) 3,347 (93)

2,777 (3) 4,549 (5) 5,020 (11) 460 (2) 1,670 (7) 243 (1) 828 (4) 178 (4) 1,651 (10) 236 (7)

20 — 24 27 — 6 14 20 — 14

7 — 13 1 — 2 4 0 — 1

1,271 1,053 1,567 5,540 249

15 15 — 218 64

7 7 — 26 1

NPGS TGSC CIP NPGS NPGS

8,771 1,885 4,690 4,643 1,841

(87) (64) (75) (46) (88)

(13) (46) (25) (54) (12)

a

Source: Refs. 49, 106, and curators (see Acknowledgments). A few values are close approximations. Additional information is in Chapters 1 through 11 in this volume. b Repositories of the NPGS (U.S. National Plant Germplasm System), TGSC (Tomato Genetic Stock Center, University of California, Davis), and centers of the Consultative Group on International Agriculture Research: CIMMYT, International Maize and Wheat Improvement Center, El Battan, Mexico; CIP, International Potato Center, Lima., Peru; ICARDA, International Center for Agricultural Research in Dry Areas, Aleppo, Syria; IITA, International Institute of tropical Agriculture, Ibadan, Nigeria. c Perennial Medicago spp.

B. SECONDARY CHEMICALS Several important examples of plant chemical defenses to insects were discovered with the aid of conserved crop germplasm (Table 2). Fraenkel43 used 6-methoxybenzoxazolinone isolated from Ostrinia nubilalis (Hübner)-resistant maize lines to support his suggestion that defense was the raison de’etre for the evolution of plant secondary chemistry. In addition, the toxicity of the sesquiterpene aldehyde dimer, gossypol, was discovered because of the susceptibility to insects of low-gossypol glandless cottons derived from conserved but obsolete cultivars such as “Acala.”12 Identification of insect-defensive chemicals in crops and crop relatives continues, exemplified by research with conserved germplasm to reveal the diversity and complexity of chemical defenses in plants.

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TABLE 2 Examples of Plant Chemical Defenses to Insects and Their Discovery in Genetic Resources Compound

Plant taxon

Genetic resources

Ref.

Cucrbitacins

Cucumis, Citrellus

14, 18

DIMBOA/MBOA

Zea mays

Leptine glycoalkaloids Methylketones Steroidal glycoalkaloids

Solanum Lycopersicon

Low cucurbitacin mutant of Citrellus vulgaris, PI274035 Wisconsin Breeding Line, W22 Iowa State University inbred line, CI 31A Several Plant Introductions PI134417 L. pimpenellifolium, LA 1335; L. esculentum var. “cerasiforme,” LA 1310 L. pennellii, LA 716

Acyl-sugars

3 68 93, 104 25 64 46, 54

1. Phytohemagglutinin-arcelin-α-amylase Inhibitors in Phaseolus Plant defenses include proteins that are toxins or digestive inhibitors.101 A group of related defensive proteins in Phaseolus vulgaris L., namely phytohemagglutinins, arcelins, and α-amylase inhibitors,81 have been studied as the basis of pest resistance in beans and as potential transgenic resistance factors for other crops (Chapters 1, 2, 4, 5, 6, 7, and 13 in this volume).86,87,112 Relying largely on the Phaseolus collection at the Centro Internacional de Agricultura Tropical (CIAT), Cali, Colombia, and Native Seed Search, Tucson, Arizona, researchers sequenced many of these proteins and measured their toxicity to the seed bruchids Zabrotes subfasciatus (Boheman), Acanthoscelides obtectus (Say), Callosobruchus maculata (F.), and C. chinensis L., and to Tenebrio molitor L.42,58,112 Toxicities are variable to the different beetle species, suggesting insect adaptation may have occurred. Some α-amylase I isoforms are toxic to Old World bruchids but not toxic or less toxic to Z. subfasciatus,57 which occurs with Phaseolus in nature. Among the New World species, Z. subasciatus is more tolerant of these inhibitors than is A. obtectus.42 Interestingly, when expressed in cultivated pea, Pisum sativum L., a bean α-amylase inhibitor confers striking resistance to the pea specialist Bruchus pisorum (L.).102 Thus, the groundwork is laid for more research involving Phaseolus germplasm to examine the relationships between bean proteins and toxicity to seed predators from an evolutionary perspective. 2. Glandular Trichome Exudates The insect defensive function of glandular trichomes was discovered largely through the association between resistance and trichome densities in crops73 and conserved crop germplasm. For example, accessions of Medicago (Chapter 8 in this volume)20 and Solanaceae (Chapter 11 in this volume)32,40,96 have improved our understanding of trichome-based defensive mechanisms. Some trichome exudates in insect-resistant

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wild Lycopersicon and Solanum contain blends of toxic or behaviorally active methyl ketones, acyl-glucoses, acyl-sucroses, and sesquiterpenes.1,24,31,45,120 In addition, insects are trapped in polyphenolic resins that form when trichome glands of some Solanum and Lycopersicon species are ruptured.26,48 Trichome exudates of Nicotiana spp. contain a variety of diterpenoids (duvatriene diols, monols, and labdanoid diterpenes), glucose and sucrose esters, and n-acylnornicotines and n-hydroxyacylnornicotines.59 The nornicotines are toxic to Manduca sexta (L.)56 and the diols, monols, and sugar esters are apparently toxic to Heliothis virescens (F.).60 Research on trichome defenses in Solanum, Lycopersicon, and Nicotiana has depended mostly on accessions from the NPGS or the Tomato Genetic Stock Center, University of California, Davis. One plant introduction (PI134417) of Lycopersicon hirsutum f. glabratum C.H. Mull., in particular, has been the subject of extensive research. However, the chemically based mechanisms of defense in this accession and other Lycopersican accessions31,120 are imperfectly known. For example, accessions of L. pennellii (Corr.) D’Arcy that vary in acylsugars103 merit study in relation to patterns of insect resistance in this germplasm. Moreover, much needs to be learned about the mechanisms of trichome-mediated defense in wild Solanum species in repositories (Table 1) (Chapter 11 in this volume).

III.

FACTORS MEDIATING HOST SELECTION

The host ranges of phytophagous insects are relatively restricted, presumably because of the combined effects of plant defenses and extrinsic ecological factors.6 Whatever the ultimate factors, plant traits affecting host selection behavior are often the most important in ecological time.8 Antixenosis in crops has provided many examples of chemical deterrents and repellents, stimulants and attractants, and physical factors that influence herbivore host selection.107 For example, trichome exudates contain oviposition stimulants,59,62 in addition to deterrents and repellents.1,45,54 Likewise, nonglandular trichomes are associated with both defense and increased acceptance by insect herbivores.83 The factors affecting host selection by phytophagous insects are complex and require understanding how insects use multiple cues to find their host plants.52 Although artificial models are helpful, it is preferable to study this process with living plants varying in multiple traits that exist in conserved germplasm. For example, breeding for reduced or increased glucosinolate content in oilseed crops has produced germplasm characterized for specific glucosinolates that may help determine the role of these compounds in mediating insect response.9

IV.

PLANT TOLERANCE TO HERBIVORY

Plants vary genetically in “tolerance” or the degree to which insect feeding reduces yield or fitness.88 First described in sorghum, Sorghum bicolor (L.) Moench., cultivars tolerant to the chinch bug, Blissus leucópterus leucópterus (Say),108 tolerance has since been documented in at least 13 crop species106 (see Chapters 1, 4, 5, 6, 7, and 8 in this volume). Most reports of intraspecific variation in tolerance to insects

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are based on work with crop genotypes and germplasm accessions. Variation in the production of dry matter67 or photosynthetic tissues,119 insensitivity to salivary toxins (Chapter 4 in this volume) or other inducers of disease-like symptoms,110 and rapid growth through vulnerable stages123 are among the tolerance mechanisms in crops. The physiological basis of these kinds of tolerance is unknown. Moreover, the inheritance of tolerance rarely has been determined because several mechanisms are involved.116 Deciphering the mechanisms can help in understanding the genetics of tolerance. Resistance to Russian wheat aphid, Diuraphis noxia (Mordvilko), in wheat, Triticum aestivum L., and barley, Hordeum vulgare L., germplasm arises from reduced plant stunting, chlorosis, and leaf rolling in response to aphid feeding.110 These separate mechanisms appear to be simply inherited and depend on a few genes (Chapters 2 and 4 in this volume). Plant tolerance to insect herbivory is receiving more attention from ecologists and evolutionary biologists,98,113 exemplified by the research of Welter and colleagues.99,119 This research demonstrated the value of conserved germplasm for studying tolerance in wild species to herbivory. More research of this type will improve our understanding of the mechanisms of tolerance and its variation in natural plant populations.

V.

MULTI-TROPHIC INTERACTIONS

Insect-plant interactions must be understood in the context of the ecological community. Predators, parasitoids, diseases of plants and insects, and plant-mycosymbiont associations affect ecological interactions and the evolution of insect herbivores and their host plants.10,53,90,105

A. ENDOPHYTIC FUNGI Although fungal endophytes occur in almost all higher plants,89 they typically are neutral with respect to herbivores. However, endophytic fungi in the genus Neotyphodium Glen, Bacon & Haulin (formerly Acremonium) frequently influence plantinsect interactions in grasses and thus, are well studied.16 Neotyphodium endophytes of cool season perennial grasses (i.e., tall fescue, Festuca arundinacea Schreb., and perennial ryegrass, Lolium perenne L.) are associated with increased resistance to at least 40 species of insects, including important pests of grasses.16 This insect resistance is caused by alkaloids characteristic of grass-fungus associations.19 Because some of these alkaloids are toxic to livestock,55 scientists have sought Neotyphodium strains that produce anti-insect alkaloids but little or no detectable levels of animal toxins.100 The discovery of different Neotyphodium strains and species in wild barley, Hordeum spp., and tall fescue germplasm15,122 foretells the probable existence of a vast endophyte resource in conserved grass germplasm. This “microbial germplasm” may provide the pool of endophytic fungi and alkaloid diversity needed to develop novel grass-endophyte combinations for insect resistance. Studies with cereal aphids, Rhopalosiphum padi (L.) and Diuraphis noxia (Mordvilko), and tall fescue, perennial ryegrass, and wild barley germplasm harboring

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different species or strains of Neotyphodium endophytes, indicate the importance of host plant species/genotype and endophyte species/strain on the expression of aphid resistance.16,17 The research also shows that both deterrent and toxic factors mediate aphid responses to endophyte infected grasses. In addition, endophytic fungi in grasses influence insect parasitoids and perhaps predators.13

B. PREDATORS

AND

PARASITOIDS

Variation in plant defense characteristics affect predators and parasitoids.10,50,90 Although these interactions have been studied most in the context of pest management, there is increasing interest in the mechanisms of tri-trophic level interactions and the ecological and evolutionary implications of these interactions for plant and insect ecology.77 1. Plant Volatiles and Entomophagous Insects Observations of entomophagous insects in the field suggest they use plant volatiles for habitat selection.117 Crop systems have provided evidence for the use of specific volatiles by parasitoids. For example, the parasitoid Campoletis sonorensis (Cameron) is attracted to specific terpenoids released from cultivars of cotton, Gossypium hirsutum L.34,35 If plants benefit from attracting entomophages, selection should favor production of the attractant volatiles specifically when plants are attacked by herbivores. Crop systems provide evidence for this kind of inducible extrinsic defense mechanism.22,114 For example, cotton plants fed upon by Spodoptera exigua (Hübner) are induced to produce a complex of volatiles that attract parasitoids.114 Moreover, crops provide evidence for genetic variability of this type of induction. The quantity and quality of parasitoid-attractive terpenoids released after feeding by larvae of S. exigua varies between a wild accession and cultivars of G. hirsutum.75 After infestation by Tetranychus urticae Koch, two cultivars of Phaseolus lunatus L. differed in their attractiveness to predaceous mites.21 The search for additional variation in inducibility could begin with conserved germplasm. 2. Physical Factors Plant surface traits and structures impede or attract entomophagous insects, as they do herbivores.115 Simple trichomes on leaves of cucumber, Cucumis sativus L., hinder the movement of the parasitoid Encarsia formosa Gahan searching for whitefly hosts. Van Lenteren and colleagues115 found the optimal densities of trichomes for parasitoid effectiveness. These densities were achieved by incorporating genes for trichome density from a hairless mutant of C. sativus (IVT 761077) in the collection of the Institute for Horticultural Plant Breeding, Wageningen, The Netherlands. Another example involves an architectural mutation in P. sativum that affects the behavior and increases predation by coccinellid beetles.65 Genes governing this and other architectural traits are conserved in the Marx pea genetic stocks collection maintained by the U.S. Department of Agriculture, Agricultural Research Service, Western Regional Plant Introduction Station, Pullman, Washington (see Chapter 7

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in this volume). Lastly, differences in leaf rolling response to feeding by D. noxia among accessions of wheat and barley affect exposure of the aphids to predators and other mortality factors.94 Chapters 2 and 3 in this volume review the genetics of wheat and barley resistance to D. noxia, respectively. 3. Surface Waxes Plants with “glossy” mutations that reduce surface waxblooms often have lower populations of some insect herbivores compared with normal-wax plants in the field.29 This phenomenon is partly caused by enhanced mobility and effectiveness of insect predators on glossy plants. Three predator species reduce populations of the diamondback moth, Plutella xylostella (L.), more effectively on glossy cabbage, Brassica oleracea var. “capitata” L., with a mutation from cauliflower (PI234599), B. oleracea var. “botrytis” L., than on plants with a waxbloom.30 These predators are more mobile and adhere better to the surface of glossy cabbage.28 A glossy mutation (wel) in P. sativum (Figure 1) also facilitates predator adhesion, mobility, and efficacy.33 This may explain reduced populations of pea aphid, Acyrthosiphon pisum (Harris), on wel/wel plants in the field. In other crops with glossy mutations, the tri-trophic component of resistance to insects has not been demonstrated, but tritrophic interactions are hypothesized for glossy wheat, sorghum, and brassicas.28 There are tradeoffs, however, because glossy Brassica and Pisum are more susceptible to coleopteran herbivores.111,121 Genes influencing waxbloom are available in accessions of Brassica, Pisum, Saccharum, and Allium, which have “reduced bloom” as a descriptor in the NPGS computer database (Genetic Resources Information Network [GRIN]).49 The inheritance of the wax traits is known for some germplasm in these collections. For example, Pisum accessions in the Marx genetic stocks collection include wax mutants at eight loci, and accessions of Brassica include mutations in at least five loci. Thus, it is possible to study the ecological implications of plant surface attributes under simple genetic control, and from these to examine the ecology of waxbloom polymorphism in natural plant populations. 4. Glandular Trichomes Glandular trichomes and their exudates can interfere with biological control.4,85 The complex mechanisms of this interference have been studied by Kennedy and colleagues working with trichome-based resistance from PI134417, L. hirsutum f. glabratum. Percent parasitism of Heliothine eggs by Trichogramma and Telenomus spp. is reduced on PI134417 and on hybrids and backcrosses to L. esculentum Miller that have 2-tridecanone in the trichome exudates.37,66 Moreover, survival of the parasitoid C. sonorensis is reduced in larvae of Helicoverpa zea (Boddie) that feed on interspecific hybrid plants producing 2-tridecanone in trichomes. Also, planidia of the parasitoid tachinid Archytas marmoratus (Townsend) are killed on the leaf surface by 2-tridecanone, while mortality of the fly larvae is increased within H. zea hosts ingesting 2-undecanone but not 2-tridecanone. In contrast, the tachinid Eucelatoria bryani (Sabroski) does not deposit planidia on the leaf surface, avoiding

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FIGURE 1 Monogenic mutations available in conserved germplasm are useful for testing the effects of specific plant characteristics on insects. Two pea genotypes vary in surface wax (wel/wel, glossy mutation with reduced waxbloom; Wel/Wel, normal-wax plants). The glossy genotype (reduced wax) is on the left in the photograph (upper panel) and in the scanning electron micrographs (×4000) (lower panels).

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TABLE 3 Use of Genetic Resources for Determination of Plant Defenses Inheritance and Other Allelochemicals Affecting Insects Compound or characteristic

Plant taxon

Genetic resourcesa

Arselins Tomatine

Phaseolus Lycopersicon

University of Wisconsin, accession UW 335 L. pimpinellifolium, LA 1335; L. esculentum var. “cerasiforme,” LA 1310 L. hirsutum f. glabratum, PI134417 L. pennellii, PI 246502; L. hirsutum, PI134417 Mentha spp. from NPGS NPGS Plant Introductions S. berthaultii, PI310927, PI283070; Unnamed NPGS Plant Introductions B. oleracea, PI234599, PI126597

Methyl ketones Glandular trichomes Terpenes Glycoalkaloids Trichomes Surface waxes a

Mentha Solanum

Brassica

Ref. 87 63 39 39, 72 74 104 44, 80 23

NPGS, U.S. National Plant Germplasm System.

direct toxicity from trichomes, but fewer larvae emerge from host larvae ingesting 2-tridecanone versus 2-undecanone.38 Finally, the predators Geocoris punctipes (Say) and Coleomegilla maculata (DeGeer) consume fewer Helicoverpa eggs in the presence of trichomes producing 2-tridecanone.2 Study of trichome exudates in other Lycopersicon germplasm and their effects on entomophages should improve our understanding of how glandular trichomes influence higher trophic levels.

VI.

GENETICS OF PLANT TRAITS AFFECTING INSECTS

A. INHERITANCE The genetic basis of plant defense and tolerance to insects was first demonstrated in crops, and crop examples still dominate (Table 3).107 Progress in understanding the genetics of defensive traits (e.g., trichomes39,69,72,80 and seed proteins58,81,87) reflects their economic importance and their availability in conserved germplasm. The development of molecular techniques for identifying quantitative trait loci (QTLs)70 for agronomic traits, including insect resistance,79 has depended on conserved germplasm. For example, the molecular maps for Zea mays L., P. vulgaris, Oryza sativa L., H. vulgare, P. sativum, Allium cepa L., L. esculentum, Solanum tuberosum L., T. aestivum and others were constructed from crosses with accessions of wild species or landraces. QTLs for insect resistance or insect-resistant traits have been identified in two wild species of Lycopersicon and in Solanum berthaultii Hawkes.11,76,82 It is possible to use these QTLs to study the distribution of quantitatively inherited defensive traits in these plant species.

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B. POPULATION GENETICS

AND

GEOGRAPHIC VARIATION

Gould47 stated that wild species in germplasm collections could be a resource for study of the population genetics of plant defenses against insects. Wild accessions approximate random samples because they usually are collected without reference to insect resistance. Using aphid infestation scores of over 200 NPGS potato accessions screened by Radcliffe et al.,91 Gould estimated among-population genetic variation for resistance to two aphid species in 14 Solanum species. This analysis illustrates the prevalence of genetic variation for insect resistance in these species. Today, a similar analysis could be conducted using the more extensive data available. Flanders et al. (Chapter 11 in this volume) present data on the extent of multi-insect species resistance in wild potato species and accessions. Among this germplasm, several species are represented by enough accessions to support deriving estimates of among-population genetic variation for insect resistance. “Passport” data on the ecogeographic origin of accessions can be used to reconstruct the geographic distribution of insect resistance in wild species. Gould47 illustrated this potential using Gibson’s44 survey of geographic variation of trichome defenses in collections of Solanum spp. Flanders et al.40,41 also investigated geographic variation in insect resistance among wild potato accessions and found “hot spots” in Latin America where greater than expected numbers of accessions of Solanum spp. are resistant to insects (Chapter 11 in this volume). The factors responsible for this pattern (Figure 2) warrant investigation. Variation in insect resistance with elevation and latitude can provide clues to possible selective factors for defense. Resistance to O. nubilalis and two pathogens among 87 inbred lines and accessions of maize from the International Maize and Wheat Improvement Center (CIMMYT), El Battan, Mexico, is related to elevation of plant origin.95 Another study found that resistance to S. exigua declined with elevation of origin among accessions of L. hirsutum.27 Among accessions of Solanum species, resistance to Colorado potato beetle, Leptinotarsa decemlineata (Say), was more prevalent in germplasm from elevations below 2500 m in North America or 3000 m in Central America.41

VII.

GERMPLASM COLLECTIONS AS REPOSITORIES FOR LIVING VOUCHERS

It is important to preserve plant germplasm used in basic and applied studies, otherwise it can be lost to science and agriculture. This happened when Brassica genotypes with glossy-wax mutations were not conserved, thus restricting more research on these mutations.124 Efforts by Waiss et al.118 to clarify the expression of insect resistance in two collections of “Zapalote Chico” maize were hampered when original collections of this “resistant variety” were not conserved in a genebank. The practice of depositing living vouchers improves genebank collections, which gain well-characterized accessions, and maximizes the value of research based on these accessions. For example, Lapointe and Tingey71 were able to research aphid

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FIGURE 2 Geographic distribution of resistance among wild potato accessions to five insect species. As compared with a null model of equal resistance over all quadrants sampled (most of Mexico and western South America), each of the quadrants shown yields a greater than expected proportion of Solanum accessions with resistance to at least one insect species. Species evaluated were Myzus persicae (Sulzer), Macrosiphum euphorbiae (Thomas), Leptinotarsa decemlineata (Say), Epitrix cucumeris (Harris), and Empoasca fabae (Harris). Resistance was determined over several years based on damage scores (L. decemlineata) or insect densities (all other species). Data from Ref. 41.

resistance in the wild potato species Solanum neocardenasii Hawkes and Hjerting (PI502642) because this germplasm was preserved in the NPGS.

VIII.

LIMITATIONS OF GENETIC RESOURCES FOR RESEARCH IN INSECT-PLANT INTERACTIONS

From 1990 to 1995, conserved germplasm was used in research in 2 to 3% of the entomology papers in four prominent journals that publish basic and applied research (Journal of Chemical Ecology, Entomologia Experimentalis et Applicata, Journal of Economic Entomology, Crop Science). This seemingly low use of germplasm by entomologists and ecologists may reflect a low knowledge or appreciation by researchers of genebanks as sources of plant materials for basic research. It also may reflect a reluctance by researchers to seek material from germplasm systems that have historically provided plant material to crop improvement programs rather than to basic research programs. There are several limitations associated with the use of conserved germplasm for insect research, which should be considered before drawing ecological or evolutionary © 1999 by CRC Press LLC

inferences from accession data. The genetics and mechanisms of defense in crop germplasm may not be relevant to natural plant populations because resistance breeding tends to favor traits conditioned by one or a few genes whose effects stand out in a genetic background of susceptibility.51 In contrast, plants in natural populations are likely to have more polygenic defensive traits. Moreover, wild accessions in genebanks may not accurately represent the original collected material. Under ex situ regeneration and storage conditions, gene frequencies can change because of inbreeding deterioration, outcrossing, inadvertent selection, and other factors.84 Modern technologies, like cryo-preservation, can minimize these problems, but older accessions could have degenerated before the advent of improved storage methods. Allogamous wild species abound in germplasm collections and some of this material may have been contaminated by genes from crop relatives and other plant species prior to their acquisition. Indeed, Jones61 questioned his data on the distribution of cyanogenesis in wild Lotus corniculatus L. because of the widespread use of cultivars of this species for forage and cover in proximity with wild populations. Lastly, ecological data for accessions are sparse or inadequate and entomological data are almost always lacking and often cannot be reconstructed because of habitat destruction. Other limitations include the problems of undocumented accessions in collections, legal and proprietary constraints, and necessary, but sometimes impeding, quarantine regulations.78 In addition, collections include only a small sample of the world’s 250,000 vascular plant species. The low number of accessions available for any one species and the way they were collected may limit their value for some research. Most wild species are represented by a few accessions (Table 2), making them inadequate for study of intraspecific variation in traits associated with insect defense. For example, Solanum rostratum Dunal, native host of Colorado potato beetle in western North America, is represented by one accession in the NPGS.49

IX.

CONCLUSIONS

Genetic variation is the essence of biology, not only as an engine of evolutionary processes, but because of its effects on the distribution, abundance, associations, and interactions of organisms, including phytophagous insects and their host plants. The mechanisms of plant defense to insects, their genetics and regulation, the ecological and metabolic costs of different defense strategies, the interactions of plant traits within the context of community, multitrophic interactions, and how these myriad factors might be shaped over evolutionary time, are not clearly understood. The conserved repositories of genetic variation in plant species are a source of material for the study of these interactions. Thus, ancillary to their primary mission of providing the raw genetic material for crop improvement, the world’s genebanks can contribute to basic research in insect-plant interactions.

ACKNOWLEDGMENTS We thank T. Hymowitz, J. Rosenthal, C. J. Simon, D. Stout, and D. Vaughan for information about germplasm holdings and L. Elberson for general assistance.

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75. Loughrin, J. H., A. Manukian, R. R. Heath, and J. H. Tumlison, Volatiles emitted by different cotton varieties damaged by feeding beet armyworm larvae, J. Chem. Ecol., 21(8):1217-1227, 1995. 76. Maliepaard, C., N. Bas, S. Van Heusden, J. Kos, G. Pet, R. Verkerk, R. Vrielink, P. Zabel, and P. Lindhout, Mapping of QTLs for glandular trichome densities and Trialeurodes vaporariorum (greenhouse whitefly) resistance in an F2 from Lycopersicon esculentum x Lycopersicon hirsutum f. glabratum, Heredity, 75:425-433, 1995. 77. Marquis, R. J., Plant morphology and recruitment of the third trophic level: subtle and little-recognized defenses?, Oikos, 75:330-334, 1996. 78. Marshall, D. R., Limitations to use of germplasm collections, In: A. H. D. Brown, O. Franke, D. R. Marshall, and J. T. Williams, Eds., Use of Plant Genetic Resources, Cambridge University Press, New York, 1989, 105. 79. McMullen, M. D. and K. D. Simcox, Genomic organization of disease and insect resistance genes in maize, Mol. Plant-Microbe Inter., 8:811-815, 1995. 80. Mehlenbacher, S. A., R. L. Plaisted, and W. M. Tingey, Inheritance of glandular trichomes in crosses with Solanum berthaultii, Am. Potato J., 60:699-708, 1983. 81. Mirkov, T. E., J. M. Wahlstrom, K. Hagiwara, F. Finardi-Filho, S. Kjemtrup, and M. J. Chrispeels, Evolutionary relationships among proteins in the phytohemagglutinin-arcelin-α-amylase inhibitor family of the common bean and its relatives, Plant Mol. Biol., 26:1103-1113, 1994. 82. Mutschler, M. A., R. W. Doerge, and J. A. Shapiro, QTL analysis of pest resistance in the wild tomato Lycopersicon pennellii: QTLs controlling acylsugar level and composition., Theor. Appl. Genet., 92:709, 1996. 83. Norris, D. M. and M. Kogan, Biochemicial and morphological bases of resistance, In: F. Maxwell and P. R. Jennings, Eds., Breeding Plants Resistant to Insects, John Wiley, New York, 1980, 23. 84. NRC (National Research Council), Managing Global Genetic Resources: Agricultural Crop Issues and Policies, National Academy Press, Washington, D.C., 1993. 85. Obrycki, J. J., The influence of foliar pubescence on entomophagous species, In: D. J. Boethel and R. D. Eikenbary, Eds., Interactions of Plant Resistance and Parasitoids and Predators of Insects, Ellis Horwood Limited, New York, 1986, 61. 86. Osborn, T. C., T. Blake, P. Gepts, and F. A. Bliss, Bean arcelin. 2. Genetic variation, inheritance and linkage relationships of a novel seed protein of Phaseolus vulgaris L., Theor. Appl. Genet., 71:847-855, 1986. 87. Osborn, T. C., T. Blake, P. Gepts, and F. A. Bliss, Insecticidal activity and lectin homology of arcelin seed protein, Science, 240:207-210, 1988. 88. Painter, R.H., Insect Resistance in Crop Plants, Macmillan, New York, 1951. 89. Petrini, O., Taxonomy of endophytic fungi of aerial plant tissues, In: N. J. Fokkema and J. van den Heuvel, Eds., Microbiology of the Phyllosphere, Cambridge University Press, Cambridge, 1986, 175. 90. Price, P. W., C. E. Bouton, P. Gross, B. A. McPheron, J. N. Thompson, and A. E. Weis, Interactions among three trophic levels: influence of plants on interactions between insect herbivores and natural enemies, Ann. Rev. Ecol. Syst., 11:41-63, 1980. 91. Radcliffe, E. B., F. I. Lauer, M.-H. Lee, and D. P. Robinson, Evaluation of the United States potato collection for resistance to green peach aphid and potato aphid, Agric. Exp. Stn. Tech. Bull. 331, University of Minnesota, St. Paul, 1981. 92. Ramachandran, R., and Z. R. Khan, Mechanisms of resistance in wild rice Oryza brachyantha to rice leaffolder Cnapholocrocis medinalis (Guenée) (Lepidopotera: Pyralidae), J. Chem. Ecol., 17:41-65, 1991.

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93. Raman, K. V., W. M. Tingey, and P. Gregory, Potato glycoalkaloids: effect on survival and feeding behavior of the potato leafhopper, J. Econ. Entomol., 72:337-341, 1979. 94. Reed, D. K., J. A. Webster, B. G. Jones, and J. D. Burd, Tritrophic relationships of Russian wheat aphid (Homoptera: Aphididae), a hymenopterous parasitoid (Diaeretiella rapae McIntosh), and resistant and susceptible small grains, Biological Control, 1:35-41, 1991. 95. Reid, L., J. T. Arnason, C. Nozzolillo, and R. Hamilton, Resistance of maize germplasm to European corn borer, Ostrinia nubilalis, as related to geographical origin, Can. J. Bot., 68:311-316, 1990. 96. Rick, C. M., C. F. Quiros, W. H. Lange, and M. A. Stevens, Monogenic control of resistance in the tomato to the tobacco flea beetle: probable repellence by foliage volatiles, Euphytica, 25:521-530, 1975. 97. Rosenthal, G. A. and M. R. Berenbaum, Herbivores: Their Interactions with Plant Secondary Metabolites, 1st Edition, Academic Press, San Diego, 1991. 98. Rosenthal, J. P. and P. M. Kotanen, Terrestrial plant tolerance to herbivory, Trends Ecol. Evol., 9:145-148, 1994. 99. Rosenthal, J. P. and S. C. Welter, Tolerance to herbivory by a stemboring caterpillar in architecturally distinct maizes and wild relatives, Oecologia, 102:146-155, 1995. 100. Rowan, D. D. and G. C. M. Latch, Utilization of endophyte-infected perennial ryegrasses for increased insect resistance, In: C. W. Bacon and J. F. White, Jr., Eds., Biotechnology of Endophytic Fungi of Grasses, CRC Press, Boca Raton, 1994, 169. 101. Ryan, C. A., Proteinase inhibitors, In: G. A. Rosenthal and D. H. Janzen, Eds., Herbivores: Their Interactions with Plant Secondary Metabolites, Academic Press, New York, 1979, 599. 102. Schroeder, H. E., S. Gollasch, A. Moore, L. M. Tabe, S. Craig, D. Hardie, M. J. Chrispeels, D. Spencer, and T. V. J. Higgins, Bean α-amylase inhibitor confers resistance to the pea weevil, (Bruchus pisorum), in transgenic peas (Pisum sativum L.), Plant Physiol., 107:1233-1239, 1995. 103. Shapiro, J. A., J. C. Steffens, and M. A. Mutschler, Acylsugars of the wild tomato Lycopersicon pennellii in relation to its geographic distribution, J. Biochem. Syst., 22:545-561, 1994. 104. Sinden, S. L., L. L. Sanford, and R. E. Webb, Genetic and environmental control of potato glycoalkaloids, Am. Potato J., 61:141-156, 1984. 105. Sinelnikov, E. A. and A. E. Tzyplenkov, Mosaic viruses influence the susceptibility of barley to net blotch and frit fly, Arch. Phytopath. Pflanz., 29:111-118, 1994. 106. SINGER (System-wide Information Network on Genetic Resources), Consultative Group on International Agricultural Research, on the World Wide Web (http://nocl. cgiar.org/cgintrus.htm), 1997. 107. Smith, C. M., Plant Resistance to Insects: A Fundamental Approach, Wiley, New York, 1989. 108. Snelling, R. O. and R. G. Dahms, Resistant varieties of sorghum and corn in relation to chinch bug control in Oklahoma, Agric. Exp. Stn. Bull. 232, Oklahoma State University, Stillwater, 1937. 109. Southwood, T. R. E., Plant surfaces and insects — an overview, In: T. R. E. Southwood and B. Juniper, Eds., Insects and the Plant Surface, Edward Arnold, London, 1986, 1. 110. Souza, E., Host plant resistance to the Russian wheat aphid (Homoptera: Aphididae) in wheat and barley, In: S. S. Quisenberry and F. B. Peairs, Eds., A Response Model for an Introduced Pest — The Russian Wheat Aphid, Entomological Society of America Thomas Say Publication, Lanham, 1997, in press.

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111. Stoner, K. A., Glossy leaf wax and host-plant resistance to insects in Brassica oleracea L. under natural infestation, Environ. Entomol., 19:730-739, 1990. 112. Suzuki, K., M. Ishimoto, K. Fumio, and K. Kitamura, Growth inhibitory effect of an α-amylase inhibitor form wild common bean resistant to the Mexican bean weevil (Zabrotes subfasciatus), Jap. J. Breed., 43:257-265, 1993. 113. Trumble, J. T., D. M. Kolodny-Hirsch, and I. P. Ting, Plant compensation for arthropod herbivory, Ann. Rev. Entomol., 38:93-119, 1993. 114. Turlings, T. C. J., J. H. Tumlinson, and W. J. Lewis, Exploitation of herbivoreinduced plant odors by host-seeking parasitic wasps, Science, 250:1251-1253, 1990. 115. van Lenteren, J. C., Biological control in a tritrophic system approach, In: D. C. Peters, J. A. Webster, and C. S. Chlouber, Eds., Aphid-Plant Interactions: Populations to Molecules, Oklahoma State University, Stillwater, 1990, 3. 116. Velusamy, R. and E. A. Heinrichs, Tolerance in crop plants to insect pests, Insect Sci. Applic., 7:689-696, 1986. 117. Vinson, S. B., Habitat location, In: D. A. Nordlund, R. L. Jones, and W. J. Lewis, Eds., Semiochemicals: Their Role in Pest Control, Wiley, New York, 1981, 51. 118. Waiss, A. C., Jr., B. G. Chan, C. A. Elliger, B. R. Wiseman, W. W. McMillian, N. W. Widstorm, M. S. Zuber, and A. J. Keaster, Maysin, a flavone glycoside from cornsilks with antibiotic activity toward corn earworm, J. Econ. Entomol., 72:256-258, 1979. 119. Welter, S. C. and J. W. Steggall, Contrasting the tolerance of wild and domesticated tomatoes to herbivory: agro-ecological implications, Ecol. Applic., 3:271-278, 1993. 120. Weston, P. A., D. A. Johnson, H. T. Burton, and J. C. Snyder, Trichome secretion composition, trichome densities, and spider mite resistance of ten accessions of Lycopersicon hirsutum, J. Am. Soc. Hort. Sci., 114:492-498, 1989. 121. White, C. W. and S. D. Eigenbrode, unpublished data, 1996-1997. 122. Wilson, A. D., S. L. Clement, and W. J. Kaiser, Endophytic fungi in a Hordeum germplasm collection, Plant Genetic Resources Newsl., 87:1-4, 1991. 123. Zwer, P. K., M. G. Mosaad, A. A. Elsidaig, and R. W. Rickman, Effect of Russian wheat aphid on wheat root and shoot development in resistant and susceptible genotypes, Crop Sci., 34:650-655, 1994. 124. Stoner, A.K., Personal communication, 1997.

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13

Biotechnological Applications of Plant Genes in the Production of Insect-Resistant Crops Angharad M. R. Gatehouse

CONTENTS I. Introduction II. Plant Genetic Resources for Insect Resistance III. Inhibitors of Proteolytic Enzymes A. Serine Protease Inhibitors B. Cysteine Protease Inhibitors C. Amylase Inhibitors IV. Enzymes V. Lectins VI. Insect-Resistant Transgenic Plants A. Protease Inhibitors B. Lectins and Lectin-Like Genes C. Chitinases VII. Pyramiding Genes for Durable Resistance VIII. Mechanisms of Insecticidal Protein Action IX. Conclusions Acknowledgments References

I.

INTRODUCTION

It is estimated that insects constitute around half of the global species diversity.38 Because a large proportion of these are phytophagous insects, the quantity and diversity of vegetation present in the world is surprising. That plants are able to tolerate predation is a result of the continual co-evolution of insects and plants, thus illustrating the fine balance which occurs in nature. What is clear is that humankind’s expectation of selected plants, i.e., those under cultivation, is very different from the requirements the plant places upon itself. In the case of the former, it is exploitation,

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and in the latter, it is survival. Thus, cultivation has upset this balance, particularly as crops have been selected on the basis of high yield, nutritional value, and adaptation to certain environmental conditions, together with low mammalian toxicity. This “selection pressure” has severely disrupted the co-evolutionary relationship between plants and insects, such that very few cultivated species have retained the resistance exhibited by their wild relatives.19 Recent figures show that the demand for food is rapidly being outstripped by supply. It is estimated that by the year 2000 there will be approximately six billion people on earth. To maintain adequate nutrition for increasing human populations (albeit suboptimal in many regions), a twofold increase in protein of plant origin and a fourfold increase in protein of animal origin are required. Because such an increase in the supply of protein from animal sources is not appropriate, particularly in the developing countries, proteins of plant origin will play a much more important and widespread role in human nutrition in the future. There is a great need for increasing agricultural production and better protecting cultivated crops. Despite an estimated annual insecticide budget of $7.5 billion (U.S.), approximately 37% of crops worldwide are lost to pests and diseases, with 13% being lost to insects alone.25 At present, crop protection in intensive agricultural systems relies almost exclusively on the use of agrochemicals, although cases exist where plant resistance and biological control have been successfully employed. Insecticide usage by pest order shows that Lepidoptera (moths and butterflies, 37% of usage) are the major target for insecticides, followed by Homoptera (sucking pests, 26%), Coleoptera (beetles, 11%) and Mites (11%).35 Although the agrochemical industry is investing in the production of safer and more environmentally friendly pesticides, much higher levels of protection are still required. Both conventional breeding and biotechnology are very important to this end. It is interesting to note that there has been a 600-fold return in research investment in varietal resistance compared to only a fivefold return in chemical pesticides.75 Therefore, the enlightened pest manager is looking toward integrated pest management, which combines practices such as the judicious use of pesticides, crop rotation, field sanitation, and the use of pest-resistant plant cultivars.58 Whichever route is embarked upon, be it via conventional plant breeding or genetic engineering, specific questions must be addressed. These include potential toxicity of the introduced gene products to intended consumers, possible environmental risks such as the effects on beneficial insects or those involved at the tritrophic level, and in the case of genetic engineering, possible gene escape. It is important that potential risks are objectively assessed.

II.

PLANT GENETIC RESOURCES FOR INSECT RESISTANCE

Plants are protected from insect and pathogen attack by many different defense mechanisms, but perhaps the most important are chemically based. Much work has been carried out to identify sources of resistance for use in crop breeding programs (Chapters 1 through 11 in this volume).10 However, in general, the mechanisms responsible for the observed resistance are poorly understood and are polygenic involving complex biosynthetic pathways. In such cases, these sources of resistance © 1999 by CRC Press LLC

are not amenable to exploitation by genetic engineering, although some research is focusing on the use of secondary metabolites via this technology.41 Endophytic fungi in some plants are responsible for resistance to at least 40 species of insects,11 but these sources of resistance are not presently amenable to genetic engineering. However, it may be possible to exploit some forms of wound-inducible resistance in plants which occur in response to insect attack. A few plant defense mechanisms are based on proteins, and because these are products of single genes, they are more amenable to genetic engineering. This chapter briefly describes these insecticidal proteins, summarizes much of the literature on their use in crop protection, and discusses the potential for improving their efficacy. The reader is referred elsewhere for detailed information on the identification of insecticidal proteins and the techniques for isolation of their encoding genes and introduction into crops by plant transformation.33,56

III.

INHIBITORS OF PROTEOLYTIC ENZYMES

By definition, protease inhibitors are substances that inhibit proteolytic enzyme activity and are specific in their interaction with proteases. Those of plant origin inhibit the proteases in animal, bacterial, and fungal fluids and secretions, but only occasionally do they inhibit plant proteases. Protease inhibitors can essentially be divided into four categories: those that specifically inhibit the serine proteases, the sulphyldryl proteases, the metallo carboxypeptidases, and the acid proteases.70 Most plant protease inhibitors fall into the first category (the serine proteases); these have received the most attention for potential use in crop protection. Attention also has focused on the use of inhibitors of sulphydryl proteases, particularly for control of coleopterans.54

A. SERINE PROTEASE INHIBITORS Protease inhibitors are widely distributed throughout the plant kingdom and are particularly abundant in seeds and storage organs where they may accumulate to approximately 1 to 10% of the total protein content. They vary in size from Mr 4,000 to 80,000, but are usually in the range of 8,000 to 20,000. This is particularly true for the legume serine protease inhibitors. Many of these inhibitors, especially those that have a molecular weight of around 10,000, are capable of simultaneously inhibiting two molecules of enzyme per inhibitor molecule. Not only can these enzymes be different from one another, as in the case of the Bowman-Birk inhibitor from soybean, Glycine max (L.) Merr., which inhibits one molecule of trypsin and one of chymotrypsin, but the enzymes inhibited may be of two different classes, as in the case of the inhibitor from ragi, Eleusine coracana (L.) Gaertner, which inhibits both trypsin and α-amylase.8 There are several Bowman-Birk iso-inhibitors present in cowpea, Vigna ungiculata (L.) Walp. (CpTI), that either inhibit trypsin only or trypsin and chymotrypsin.32 Potato, Solanum tuberosum L., and tomato, Lycopersicum esculantum Miller, plants contain two powerful inhibitors of serine proteases designated Inhibitor I (monomer Mr 8,100) and Inhibitor II (monomer Mr12,300).62 Inhibitor I is an inhibitor of chymotrypsin that only weakly inhibits trypsin at its single reactive site, while

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inhibitor II contains two reactive sites, one of which inhibits trypsin while the other inhibits chymotrypsin. Both inhibitors are synthesized as precursors and undergo post-translational modification to form the mature proteins which are sequestered into the vacuole.12 These inhibitors are particularly interesting because they are induced in leaf tissue of potato and tomato in response to wounding, even though they accumulate in potato tubers.37 Because the major digestive endoproteases in the insect gut are the serine proteases trypsin and chymotrypsin,78 these inhibitors have great potential in crop protection (especially against lepidopteran pests). Therefore, studies have evaluated the effects of these inhibitors on insects in vitro,9,51 against gut enzymes to identify those with high binding affinities, and in artificial diets to identify those that are toxic.6,7,16,31,50,76

B. CYSTEINE PROTEASE INHIBITORS Cysteine protease inhibitors have been isolated from a number of plant sources where they are thought to play an important role in regulating endogenous proteolysis. Unlike the serine protease inhibitors, a role in pest defense is less likely because they are usually present at very low levels. However, many insects, particularly Coleoptera,26 rely on cysteine proteases for major digestive proteolysis. It would therefore be logical to determine if inhibitors of these particular enzymes would effectively control coleopterans. To this end, in vitro proteolysis studies have demonstrated that the inhibitor from rice, Oryza sativa L., oryzacystatin (OC-1), inhibits the digestive proteases of several different coleopterans.17,55 This group of inhibitors has been effective in vitro and their insecticidal properties have been demonstrated in artificial diet studies.17

C. AMYLASE INHIBITORS Protein inhibitors of mammalian α-amylases, abundant in cereal grains, are also present in legume and other seeds.69 There are several distinct sequence families, with some being homologous to the Bowman-Birk type protease inhibitors. Generally, these inhibitors have relatively low molecular weights and although some may inhibit both mammalian and insect α-amylase activity, others are far more specific, being effective against either one or the other. Although induced synthesis of amylase inhibitors by insect attack has not been reported, those purified from wheat, Triticum aestivum L., and common bean, Phaseolus vulgaris L., are insecticidal toward coleopterans when incorporated into artificial diets.30,46,59 Different types of α-amylase inhibitor in wheat endosperm are differentially active against different lepidopterans. When tested in artificial diet against a range of phytophagous Lepidoptera and Coleoptera, the effects of wheat α-amylase inhibitor in vivo ranged from little or no effect, to significant effects on development and mortality.39

IV.

ENZYMES

Several plant enzymes have potential in crop protection. These include peroxidases (POD) which oxidize an array of substrates using H2O2 as the oxidizing agent,

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lipoxygenases (LOX) which catalyze the oxidation of certain polyunsaturated fatty acids, and polyphenol oxidases (PPO) which catalyze the oxidation of mono-, di-, and polyhydric phenols to quinones. All three classes have been implicated in conferring resistance to insects. Soybean LOX is toxic to rice brown plant hopper, Nilaparvata lugens (Stal.), when fed an artificial diet,63,64 while other lypoxygenases and peroxidases are induced as a result of herbivory.20,21,23 Felton and Gatehouse22 review the role of these enzymes in crop protection.

V.

LECTINS

Lectins, a large diverse group of proteins, can be identified by a common property of binding to carbohydrate residues, either as free sugars or as part of oligo- or polysaccharides. Most lectin molecules contain multiple binding sites and thus can cross-link oligo- or polysaccharides. The first plant lectin was described over a century ago when Stillmark77 demonstrated that extracts from castor bean, Ricinus communis L., agglutinated red blood cells. Many belong to a homologous family of proteins based on an amino acid chain of approximately 220 residues, although totally different sequence types have been shown to have similar functional properties. Pusztai’s 65 review is a general reference for plant lectins. A limited number of insecticidal lectins have been identified by their protective role within the seed, as typified by the bean lectins and winged bean, Psophocarpus tetragonolobus (L.) DC, lectins.28,34,48 Many studies, however, also have involved the screening of purified lectins against economically important insect pests in an attempt to identify insecticidal proteins,61 and hence isolate their encoding genes for subsequent plant transformation. Such screening has identified lectins toxic to several insect pests (Table 1).35 As mentioned earlier, before transforming crops encoding antimetabolic/toxic proteins to confer protection against pests, consideration must be given to their potential mammalian toxicity. This is particularly true for lectins, many of which are highly toxic to mammals and birds.1

VI.

INSECT-RESISTANT TRANSGENIC PLANTS

Production of insect-resistant plants by genetic engineering became possible almost as soon as the technology became available, with the first report describing the use of genes encoding the insecticidal toxins from Bacillus thuringiensis Berliner (Bt).81 Bt has an established track record in agriculture as sprays and dusts. An alternative strategy is to exploit defensive proteins found in plants. High expression of foreign proteins is relatively straightforward because the gene transfer is from one plant species to another, and the transcription and translation systems are similar. The drawback is that insect predators of plants have been exposed to the defensive proteins being introduced and thus may have mechanisms to counteract their effects. Fortunately, the diversity of defensive proteins found in the plant kingdom, and the differing specificities shown by proteins of the same type from different plant sources, means that transfer of such proteins from one plant species to another can give effective levels of resistance.

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TABLE 1 Insecticidal Plant Lectinsa Insect order

Insect

Lepidoptera

Ostrinia nubilalis

Coleoptera

Lacanobia oleracea Callosobruchus maculatus

Homoptera

Zabrotes subfaciatus Diabrotica undecimpunctata Nilaparvata lugens

Diptera

Acyrthosiphon pisum Myzus persicae Aulacorthum solani Empoasca fabae Lucilia cuprina

a

Lectin

Sugar specificity

Ref.

Castor bean Camel’s foot tree Wheatgerm (WGA) Snowdrop (GNA) Bean (PHA) Winged bean Griffonia Various sources Rice Stinging nettle (UDA) Snowdrop (GNA) Elderberry (SNA-1) Bean (Arcelin) Various sources Snowdrop (GNA) Snowdrop (GNA) Wheatgerm (WGA) Jackbean (Con A) Snowdrop (GNA) Snowdrop (GNA) Various sources Wheatgerm (WGA) Jackbean (Con A)

GalNAc GalNAc GlcNAc Mannose Complex carbohydrates GlcNAc GlcNAc GalNAc & GlcNAc GlcNAc GlcNAc Mannose 2,6-neuraminyl-gal/GalNAc — GalNAc & GlcNAc Mannose Mannose GlcNAc Glucose/Mannose Mannose Mannose GalNAc & GlcNAc GlcNAc Glucose/Mannose

13 13 13 24 28,48 34 83 61 45 45 35 35 59 13 17 63,64 63 67,68 71 14 40 18 18

Artificial diet studies.

A. PROTEASE INHIBITORS The first example of a foreign plant gene conferring resistance to insects was the transfer of a chimaeric cowpea trypsin inhibitor (CpTI) gene, under the control of the constitutively expressed CaMV 35S promoter, into tobacco, Nicotiana tabacum L.43 Plants expressing the inhibitor at approximately 1% of total soluble protein exhibited significantly enhanced levels of resistance to the tobacco budworm, Heliothus virescens (F.) (Figure 1), and later to other lepidopteran pests, including H. zea (Boddie), Spodoptera littoralis (Boisd.), and Manduca sexta (L.). Although initially expressed in tobacco, CpTI also has been transferred into many other crops where it is effective against insects (Table 2). Despite CpTI being an effective antimetabolite against a wide spectrum of insect pests, mammalian feeding trials have failed to demonstrate acute toxicity.66 This is an important consideration for the use of this gene in crop plants. After these initial studies with CpTI, many other plant serine protease inhibitors have been expressed in plants where they confer enhanced levels of protection to insects (Table 2).

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FIGURE 1 Bioassay of control and CpTI expressing tobacco plants against larvae of Heliothis virescens (Tobacco/Cotton budworm). Control plant (left) showing high levels of leaf damage; transgenic plant (right) showing minimal damage.

Although cysteine (thiol) protease inhibitors have been suggested for use in transgenic plants for control of coleopterans, little work has been done on the expression of genes encoding these inhibitors. One example, however, is the constitutive expression of a gene encoding the cysteine protease inhibitor from rice (oryzacystatin or OC-1) in transgenic poplar trees, where it confers resistance to the beetle Chrysomela tremulae F.54

B. LECTINS

AND

LECTIN-LIKE GENES

Lectins are plant derived genes of interest for insect control, particularly for control of Homoptera (Table 3). However, as previously mentioned, caution must be exercised in the selection of appropriate lectin encoding genes since many of these proteins are toxic to mammals. The first lectin expressed in transgenic plants was that encoding the pea lectin, where expression at about 1% of total soluble protein in tobacco resulted in enhanced levels of resistance to H. virescens.3 More recently, the gene encoding snowdrop lectin (GNA) has been expressed constitutively in potato plants where it confers significant levels of protection toward aphids and Lepidoptera (Figure 2). Against the tomato moth, Lacanobia oleracea (L.), there were significant reductions in larval biomass (>50%) and leaf damage (>70%) under laboratory conditions.27 Similar results were obtained in large-scale glasshouse trials.24 GNA expressing potato plants also significantly reduced the development and fecundity of two aphid species, Myzus persicae (Sulz.) and Aulacorthum solani (Kalt.).14,29 These results were reproducible under glasshouse conditions.14 Phloem-specific GNA expression in transgenic tobacco and rice plants has been achieved, using a gene construct containing the GNA coding sequence driven by the promoter from the rice sucrose synthase gene, Rss1.74 Additionally, research

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TABLE 2 Transgenic Plants Expressing Plant Protease Inhibitor Genes Genesa

Insect

Ref.b

CpTI Pot PI II CpTI + p-lec Na PI CpTI Pot PI I

Heliothis virescens Lepidoptera Heliothis virescens Helicoverpa punctigera Lacanobia oleracea Helicoverpa armigera Teleogryllus commodus Helicoverpa armigera Teleogryllus commodus

43 49, 53, 57 3 42 Durham Univ./Pestax* Ryan & Markwick* Ryan & Burgess* Ryan & Markwick* Ryan & Burgess* Pestax* 15 15 82 82 36 Axis & Van Der Haave* Appleby & Burgess* Pestax* Pestax* INRA INRA INRA 79+ 80+ HRI* 54 INRA Von Weissenberg*

Plant Tobacco

Potato Tomato

Pot PI II

Rice

CpTI Pot PI II CpTI

Strawberry Lettuce Sweet potato Oilseed rape

Cotton Alfalfa Apple Poplar Birch

CpTI CpTI Pot PI II CpTI CpTI OC-I CII M.S PI M.S PI CpTI OC-1 CII Pot PI II

Sesamia inferens Chilo suppressalis Sesamia inferens Chilo suppressalis Otiorhynchus sulcatus Teleogryllus commodus

Coleoptera Lepidoptera Diptera Bemisia tabaci Thrips Cydia pomenella Chrysomela tremulae Lepidoptera

a

CpTI = cowpea trypsin inhibitor; CII= double headed serine protease inhibitor from soybean; Na PI = Nicotiana alata protease inhibitor; OC-1 = oryzacystatin; Pot PI II= Potato proteinase inhibitor II; Pot PI I = Potato proteinase inhibitor I; p-lec = pea lectin. b * = unpublished; + = Proteinase inhibitor from Manduca sexta; INRA = Institute National de la Recherche, France.

demonstrates that expression of GNA in the phloem of transgenic rice plants confers protection against the brown plant hopper.84 The α-amylase inhibitor from bean is encoded by a lectin-like gene, whose product is referred to as LLP.60 The seeds from transgenic pea plants, which expressed the α-amylase inhibitor at about 0.3% total protein, were highly resistant to attack by the bruchids Bruchus pisorum (L.) (Chapter 7 in this volume) and Callosobruchus maculatus (F.).72,73 Expression of this gene in Adzuki bean, Vigna angularis (Willd.) Ohwi and Ohashi, also confers resistance to C. chinesis (L.).47

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TABLE 3 Transgenic Crops Expressing Plant Lectin and Lectin-Like Genes Genesa

Plant Tobacco

GNA

Potato

p-lec CpTI + p-lec GNA

GNA + BCH Tomato Rice Sweet potato Oilseed rape Pea

GNA GNA GNA GNA a-AI

Azuki bean

a-AI

Insect Heliothis virescens Myzus persicae Heliothis virescens Lacanobia oleracea Myzus persicae Aulacorthum solani Myzus persicae Aulacorthum solani Lacanobia oleracea Nilaparvata lugens

Zabrotes subfaciatus Bruchus pisorum Callosobruchus chinensis

Ref.b Durham Univ./Pestax* 44 3 3 27 29 14 29 Down* Pestax/CSL* Durham Univ./Hodges* Pestax* Pestax* 73 72 47

a

GNA = snowdrop lectin; p-lec = pea lectin; CpTI + p-lec = cowpea trypsin inhibitor + pea lectin; GNA + BCH = snowdrop lectin + bean chitinase; a-AI = bean α-amylase inhibitor. b * = unpublished.

C. CHITINASES Beyond their potential in control of fungal pathogens, chitinases may protect crops against certain insects, especially Homoptera. Transgenic potato plants, expressing a gene encoding bean chitinase (BCH) under control of the constitutive CaMV 35S promoter, reduced the fecundity of the glasshouse potato aphid, A. solani, although this reduction was not statistically significant. However, nymphs produced on these BCH expressing plants were significantly smaller compared to those on control (untransformed) plants.85

VII.

PYRAMIDING GENES FOR DURABLE RESISTANCE

One of the goals of the plant breeder is to “pyramid” genes expressing agriculturally desirable characteristics. This strategy also has been adopted by the biotechnologist. It is envisaged that “packages” of different genes will be introduced into crops to increase the protective efficacy, spectrum of activity, and durability of resistance. The components of such packages should each act on different targets within the insect and thus, mimic the multi-mechanistic resistance which occurs in nature. Protease inhibitors should be particularly valuable in this respect since, apart from their

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FIGURE 2 Bioassay of control and GNA expressing potato plants against larvae of Lacanobia oleracea (Tomato moth). Control plant (left) showing severe levels of damage; transgenic plant (right) showing significantly reduced levels of damage.

inherent insecticidal effects, they would protect other introduced gene products from premature digestion in the insect gut. The first demonstration of gene pyramiding was the introduction of both CpTI and pea lectin (P-Lec) into tobacco. These plants were obtained by cross-breeding plants derived from the two primary transformed lines.3 The insecticidal effects of the two genes were additive, not synergistic, with insect biomass on the double expressors reduced by nearly 90% compared with those from control plants, and 50% compared with those from plants expressing either CpTI or P-Lec alone. Also, leaf damage was the least on the double expressing plants. More recently, potato plants expressing both GNA and bean chitinase (BCH) have been obtained. When tested for enhanced aphid resistance, fecundity was reduced by approximately 95% compared with control plants. Fecundity on plants expressing GNA alone was reduced by about 70% while the reduction in fecundity on those expressing BCH © 1999 by CRC Press LLC

alone was small and not statistically significant. These findings clearly demonstrate that these two genes have a synergistic effect against aphids.14 Interestingly, this was not observed in trials where these double expressors were infested with larvae of the tomato moth. In this instance, larval biomass and leaf damage were significantly reduced compared with control plants, but were higher on the plants expressing GNA alone.27 This occurred because larvae performed better and caused more damage on the BCH expressing plants. These results emphasize the care required in selecting appropriate genes for controlling specific pests.

VIII.

MECHANISMS OF INSECTICIDAL PROTEIN ACTION

Although there are many examples of different plant species, including trees, which have been transformed with insecticidal genes of plant origin, and which show significantly enhanced levels of resistance to insect pests (Tables 2 and 3), it is questionable if these levels of protection are sufficient to satisfy commercial criteria. To fully exploit this technology, detailed studies are being conducted on the modes of action of selected insecticidal proteins at the biochemical and molecular level with a view to optimizing their efficacy in the field. Some authors have suggested that selection of protease inhibitors for use in crop protection should be based upon the binding affinity of the inhibitor to pest insect proteases, although there is little published evidence of this approach being followed. One approach of great potential that is under investigation is the use of a phage display system to select inhibitors with desired properties.86 However, the assumption that the higher the affinity of an inhibitor for a target protease in vitro, the more effective that inhibitor will be in vivo, is open to question. Interestingly, although the soybean Kunitz trypsin inhibitor (SKTI) has been shown to be a potent inhibitor of gut proteases of several major lepidopteran pests in vitro, there do not appear to be any reports of transgenic plants expressing SKTI which exhibit significantly enhanced levels of insect resistance. In experiments by the author and colleagues, transgenic tobacco plants expressing high levels of SKTI were not protected against larvae of H. virescens, despite the fact that this inhibitor had been shown to inhibit gut protease enzymes in vitro. One explanation for this observation may be the induction of “inhibitor insensitive” proteases, recently reported for some Lepidoptera5,52 and Coleoptera.2 Further work on characterizing insect digestive proteases, both natural and induced, and on the mechanisms that control their expression, will be necessary before rational prediction of inhibitor effectiveness can be made. Recent results have shown that lepidopteran larvae contain and express at least 20 genes encoding digestive proteases, and that ingestion of protease inhibitors up-regulates expression of some of these genes and down-regulates expression of others.4 The mode of action of insecticidal lectins is less understood than that of protease inhibitors. Although lectins have been observed to bind to the gut surface in insects28 and to the peritrophic membrane,18 it is not clear why such binding should lead to toxic effects, or indeed whether the toxicity is dependent on the observed binding. Studies in progress in the U.K. are attempting to relate structure to function in the insecticidal lectin GNA through in vitro mutagenesis. This research should provide © 1999 by CRC Press LLC

greater understanding of the modes of activity of lectins, and should enable researchers to target them more precisely and to increase their efficacy.

IX.

CONCLUSIONS

There have been several clear examples that certain proteins in plants, e.g., enzyme inhibitors and lectins, have a protective role, particularly within storage tissues such as seeds. Others have been shown to be insecticidal to important insect pests when tested in artificial diets and, in a few limited examples, when their encoding genes have been expressed, in transgenic plants. Engineering transgenic plants to express foreign insecticidal proteins is a means of producing crops with enhanced levels of insect resistance, which, if adopted, could complement other forms of crop protection. The technology has the potential to move farming closer to ecologically sustainable practices, both in developed and developing countries. A strategy that has been suggested to maximize the utility of this technology is to go beyond the use of single genes, and use gene combinations whose products are targeted to different biochemical and physiological processes within the insect. In this way, it is hoped to provide a multi-mechanistic form of resistance which can be tailored to the different crops and prevailing insect pests at a given time. However, proposals to use insecticidal proteins as resistance factors in transgenic plants must consider the possible effects on beneficial insects, e.g., insect parasitoids and predators, and insect pollinators. Several laboratories are actively addressing this aspect. Genetic engineering to enhance resistance of plants to insect attack has been developed from a theoretical possibility to a commercial reality, and will come to play an increasingly important role in crop protection. However, it is not an exclusive technology and should not be thought of as a replacement for naturally occurring resistance genes which may be present in the gene pool of crop species. Engineered and natural resistance can be seen as complementary technologies to combat pests, and are likely to act in concert with each other. Many natural resistance mechanisms involve alterations in secondary metabolism, or in the physical structures of plant tissues, which are not achievable at present through genetic engineering. As well, it is possible to envisage crops in which protection against a specific pest is conferred by both natural and engineered resistance genes, and where the protective genes rely on different mechanisms of toxicity or deterrence. Developing transformation technologies that allow efficient engineering of resistant crop cultivars is a priority. Multiple mechanisms of resistance in a single crop would be highly advantageous in the field, resulting in more durable resistance compared with that conferred by single genes. Although genetic engineering will assist in protecting crops, global plant genetic resources will remain vital to plant breeding in the future.

ACKNOWLEDGMENTS The author acknowledges the Rockefeller Foundation, the Department of Trade and Industry, Axis Genetics Ltd (Pestax), the Biotechnology and Biological Sciences Research Council (BBSRC), the Ministry of Agriculture, Food and Fisheries

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(MAFF), the European Union, and The Scottish Office for financial support. I thank the following for permission to report on unpublished data or current research: M. Jongsma, E. P. J. Burgess, C. A. Ryan, T. K. Hodges, N. Markwick, K. Von Weissenberg, L. Jouanin, and scientists at Institut National de la Recherche (INRA; France), Central Science Laboratory (CSL), and Horticulture Research International (HRI; UK), together with my colleagues J. A. Gatehouse, M. Bharathi, R. E. Down, and R. Raemakers at Durham University, U.K.

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