Recent Advances in Entomological Research: From Molecular Biology to Pest Management

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Tongxian Liu Le Kang

Recent Advances in Entomological Research From Molecular Biology to Pest Management

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Tongxian Liu Le Kang

Recent Advances in Entomological Research From Molecular Biology to Pest Management

With 87 figures, 3 of them in color

Editors Tongxian Liu Key Laboratory of Applied Entomology Northwest A & F University Yangling, Shaanxi, 712100, China; Email: [email protected]

Le Kang State Key Laboratory of Integrated Management of Pest Insects and Rodents

Institute of Zoology Chinese Academy of Sciences Beijing, 100101, China Email: [email protected]

ISBN 978-7-04-028988-6 Higher Education Press, Beijing ISBN 978-3-642-17814-6 Springer Heidelberg Dordrecht London New York

e-ISBN 978-3-642-17815-3

Library of Congress Control Number: 2011920986 © Higher Education Press, Beijing and Springer-Verlag Berlin Heidelberg 2011 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Printed on acid-free paper Springer is part of Springer Science + Business Media (www.springer.com)

Preface

Insects represent the most diverse group of species at planet earth, accounting for over 50% of known organisms. Their close interaction with human and other life forms has significant impacts on human health, environments, agriculture, biosafety, etc. Thus, entomology has been a hot research topic for worldwide scientists for long time. The development of modern biology such as molecular biology, cell biology, genetics, integrates new elements and concepts into the classical entomology. Now, over ten insect genomes have been sequenced. These data, plus the novel tools and thoughts, provide tremendous amounts of information for entomological researchers to deeply and systematically study insects. We, as entomologists, were fascinated, and then were inspired to edit a book to present such rapid advances and progresses in entomological research. This motivation was realized as a result of our opportunity in interacting with numerous entomologists during academic research. We invited more than forty scientists with research specialties ranging from molecular biology to pest management to contribute chapters with a most comprehensive overview to date to include most, if not all, recent advances in their field of specialties. This book contains 25 chapters, ranging from molecular biology to applied pest management, authored by 49 scientists. The first section, Insect-Plant Interactions, include five chapters, covering deciphering the plant-insect phenotypic arms race, inducible plant defense against insect herbivores, host marking and host discrimination in phytophagous insects, and plant’s defense modulated by minerals. The second section, Molecular Biology, Physiology, Behavior and Ecology, comprises seven chapters, including recent advances in virus infection in honey bee, biological function of insect yellow gene family, the function of bursicon, a neuropeptide hormone, chemical ecology of bark beetle, inforchemical tritrophical interactions in soybean aphids-host plants-natural enemies, the response of insects to global warming, and the biology and reproductive strategies of the subterranean termites.

The third section, Insect Toxicology and Insecticide Resistance Management, consists of seven chapters, including the roles of P450s in insecticide resistance and interactions with bioactive agents, metamorphosis of innate insect resistance in host plants research, new discoveries in genetically modified crops and natural enemies, and the molecular mechanism of insecticide resistance in mosquitoes and other insect pests of field crops. The fourth section, Emerging Pest Management Strategies and Technologies, contains six chapters with a broad range from RNAi technologies, anti-tick vaccine, veterinary pests, biological and integrated management strategies of various field crops, invasive imported red fire ants, urban pest management, and an emerging area of entomological science that utilizes lignocellulose-feeding insects for viable biofuels. We think that each chapter is sufficiently thought-provoking that it is expected to find its way onto the bookshelves of scientists, post-graduate students and advanced undergraduate students who are interested in insect molecular biology, insect-plant interactions, insecticide toxicology and resistance management, integrated pest management, and agriculture and urban entomology. We would like to acknowledge the numerous referees that read and commented critically on each chapter. They are acknowledged in each chapter. Also, we want to acknowledge Miss. Dan Yu of Institute of Zoology, Chinese Academy of Sciences, Miss. Chao Pan of Higher Education Press, and people in Springer for their great assistance during the publication process. Tongxian Liu Le Kang December 24, 2010

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Contents

Section 1: Insect-Plant Interactions 1. Deciphering the Plant-Insect Phenotypic Arms Race · · · · · · · · Xianchun Li, Xinzhi Ni

3

2. Insect Herbivory-Inducible Proteins Confer Post-Ingestive Plant Defenses · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · ·Keyan Zhu-Salzman, Tongxian Liu

34

3. Inducible Direct Defense of Plants Against Insects · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Mingshun Chen, Junxiang Wu, Guohui Zhang

49

4. Host Marking and Host Discrimination in Phytophagous Insects · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Siwei Liu, Baige Zhao, Edmond Bonjour

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5. Nitrogen Modulation on Plant Direct and Indirect Defenses · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Yigen Chen, Xinzhi Ni

86

Section 2: Molecular Biology, Physiology, Behavior and Ecology 6. Viruses and Viral Diseases of the Honey Bee, Apis mellifera · · · · · · ·Yanping Chen 105 7. Biological Function of Insect Yellow Gene Family · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Jianyong Li, Bruce M. Christensen

121

8. Bursicon, a Neuropeptide Hormone That Controls Cuticle Tanning and Beyond · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Qisheng Song, Shiheng An

132

9. Chemical Ecology of Bark Beetles in Regard to Search and Selection of Host Trees · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · John A. Byers, Qinghe Zhang

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10. Infochemical-tritrophic Interactions of Soybean Aphids-host Plants-natural Enemies and Their Practical Applications in Pest Management · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · ·Junwei J. Zhu

191

11. The Responses of Insects to Global Warming · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Kun Guo, Osbert Jianxin Sun, Le Kang

201

12. Biology and Reproductive Strategies in the Subterranean Termites (Isoptera: Rhinotermitidae) · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Xingping Hu

213

Section 3: Insect Toxicology and Insecticide Resistance Management 13. P450–mediated Insecticide Detoxification and Its Implication in Insecticide Efficacy · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Zhimou Wen, Xing Zhang, Yalin Zhang

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14. House Fly Ctyochrome P450s: Their Role in Insecticide Resistance and Strategies in the Isolation and Characterization · · · · · · · · · · · · · · · · · · · · ·Nannan Liu, Fang Zhu

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15. Metamorphosis of Cisgenic Insect Resistance Research in the Transgenic Crop Era · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Xinzhi Ni, Xianchun Li, Yigen Chen, Fuzhen Guo, Jinian Feng, Huiyan Zhao

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16. Time for a New Look at the Relationship Between Bt Plants and Insect Natural Enemies · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Mao Chen, Anthony M. Shelton

280

17. The Development of Pyrethroid Resistance in the Mosquito Culex quinquefasciatus · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Qiang Xu, Nannan Liu

295

18. Resistance to Transgenic Bacillus thuringiensis Crops in Target Insect Pests: Current Status and Prospect · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Fangneng Huang

306

19. Potential Use of Proteinase Inhibitors, Avidin, and other Bio-reagents for Synergizing Bt Performance and Delaying Resistance Development to Bt · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · ·Yucheng Zhu, Mingshun Chen, Craig A. Abel

330

Section 4: Emerging Pest Management Strategies and Technologies 20. Advances and Prospects of RNAi Technologies in Insect Pest Management · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · ·Xin Zhang, Jianzhen Zhang, Kunyan Zhu

347

21. Anti-tick Vaccine Development: Status and Perspectives · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Quentin Q. Fang, Oscar J. Pung

359

22. Hymenopteran Parasitoids and Their Role in Biological Control of Vegetable Liriomyza Leafminers · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Tongxian Liu, Le Kang, Zhongren Lei, Ricardo Hernandez 376

23. Development of an Integrated Greenhouse Whitefly Management Program on Strawberries · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Jianlong Bi

404

24. Advances in Research on the Venom Chemistry of Imported Fire Ants · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · · Jian Chen, Hanwu Shang

417

25. Utilization of Lignocellulose-feeding Insects for Viable Biofuels: an Emerging and Promising Area of Entomological Science · · · · · Jianzhong Sun, Xuguo Joe Zhou

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434

Contributing Authors

The editors would like to extend our special and sincere thanks to all 49 people who contribute their research and time to this book. Craig A. Abel

Research Entomologist, Southern Insect Management Research Unit, United States Department of Agriculture, Agriculture Research Service, Stoneville, MS, 38776, USA ([email protected]).

Shiheng An

Division of Plant Sciences, University of Missouri, Columbus, MO, 65211, USA.

John A. Byers

Research Entomologist, Arid-Land Agricultural Research Center, USDA-ARS, 21881 North Cardon Lane, Maricopa, Arizona, 85238, USA ([email protected]).

Jianlong Bi

Extension Specialist, University of California Cooperative Extension, Salinas, CA, 93901, USA ([email protected]).

Edmond Bonjour

Department of Entomology and Plant Pathology, Oklahoma State University, Stillwater, OK, 74078, USA ([email protected]).

Jian Chen

Research Entomologist, Biological Control of Pests Research Unit, Agriculture Research Service, United States Department of Agriculture, Stoneville, MS, 38776, USA ([email protected]).

Mao Chen

Department of Entomology, Cornell University, NYSAES, Geneva, New York, 14456, USA ([email protected]).

Mingshun Chen

Research Entomologist, Department of Entomology, Kansas State University, Manhattan, USDA-ARS, KS, 66506, USA ([email protected]).

Yanping Chen

Research Entomologist, Bee Research Laboratory, U.S. Department of Agriculture, Agricultural Research Service, Beltsville, MD, 20705, USA ([email protected]).

Yigen Chen

Department of Entomology, Michigan State University,

East Lansing, MI, 48824, USA ([email protected]). Bruce M. Christensen Department of Pathobiological Sciences, University of Wisconsin-Madison, 1655 Linden Drive, Madison, WI, 53706, USA. Qingquan Fang

Department of Biology, Georgia Southern University, Statesboro, GA, 30460, USA (qfang@georgiasouthern. edu).

Jinian Feng

College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China ([email protected]. cn).

Fuzhen Guo

College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China.

Kun Guo

State Key Laboratory of Integrated Management of Pest Insects and Rodents, Institute of Zoology, Chinese Academy of Sciences, Beijing, 100101, China.

Le Kang

State Key Laboratory of Integrated Management of Pest Insects and Rodents, Institute of Zoology, Chinese Academy of Sciences, Beijing, 100101, China ([email protected]).

Ricardo Hernandez

Texas AgriLife Research, Texas A&M University System, Weslaco, Texas, 78596, USA.

Xingping Hu

Department of Entomology and Plant Pathology, Auburn University, Auburn, Alabama, 36849, USA ([email protected]).

Fangneng Huang

Department of Entomology, Louisiana State University Agricultural Center, Baton Rouge, Louisiana, 70803, USA ([email protected]).

Jianyong Li

Department of Biochemistry, Virginia Tech, 111 Engle Hall, Blacksburg, VA, USA ([email protected]).

Xianchun Li

Department of Entomology, University of Arizona, Tucson, AZ, 85721, USA ([email protected]).

Zhongren Lei

Institute of Plant Protection, Chinese Academy of Agricultural Sciences, Beijing, 100081, China ([email protected]).

Nannan Liu

Department of Entomology and Plant Pathology, Auburn University, Auburn, Alabama, 36849, USA (liunann@au-

x

burn.edu). Siwei Liu

Department of Biological Sciences, The University of Texas at El Paso, El Paso, Texas, 79968, USA; Department of Entomology and Plant Pathology, Oklahoma State University, Stillwater, OK, 74078, USA ([email protected]).

Tongxian Liu

Key Laboratory of Applied Entomology, Northwest A&F University, Yangling, Shaanxi, 712100, China; and Texas AgriLife Research, Texas A&M University System, Weslaco, Texas, 78596, USA.

Xinzhi Ni

Research Entomologist, Crop Genetics and Breeding Research Unit, USDA–ARS, Tifton, Georgia, 31793, USA ([email protected]).

Oscar J. Pung,

Department of Biology, Georgia Southern University, Statesboro, GA, 30460, USA.

Keyan Zhu-Salzman Department of Entomology, Texas A&M University, College Station, TX 77843, USA ([email protected]). Hanwu Shang

College of Life Sciences, China Jiliang University, Hangzhou, Zhejiang, 310018, China (hwshang@cjlu. edu.cn).

Anthony M. Shelton Department of Entomology, Cornell University, NYSAES, Geneva, New York, 14456, USA ([email protected]). Qisheng Song

Division of Plant Sciences, University of Missouri, Columbia, MO, USA ([email protected]).

Jianzhong Sun

School of the environment, Jiangsu University, 301 Xuefu Rd. Zhenjiang, Jiangsu, 212013, China (current working affiliation) ([email protected])

Osbert Jianxin Sun

Key Laboratory for Silviculture and Conservation of Ministry of Education, College of Forest Science, Beijing Forestry University, Beijing, 100083, China.

Zhimou Wen

Department of Entomology, University of Georgia, Athens, GA, 30602, USA (zwen0920@gmail. com).

Junxiang Wu

Department of Entomology, College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China ([email protected]).

Qiang Xu

Department of Biology, Abilene Christian University, Abilene, Texas 79699, USA ([email protected]). xi

Guohui Zhang

Department of Entomology, College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China.

Jianzhen Zhang

College of Life Science, Shanxi University, Taiyuan, Shanxi, 030006, China.

Qinghe Zhang

Sterling International, Inc., 3808 N. Sullivan Rd, Bldg 16P, Spokane, WA 99216-1630, USA (qing-he@rescue. com).

Xin Zhang

Department of Entomology, Kansas State University, Manhattan, KS, 66506, USA.

Xing Zhang


Yalin Zhang


Baige Zhao

Department of Plant and Soil Sciences, Oklahoma State University, Stillwater, OK, 74078, USA.

Huiyan Zhao


Xuguo Joe Zhou

Insect Integrative Genomics, Department of Entomology, University of Kentucky, Lexington, Kentucky, KY, 40546, USA ([email protected]).

Fang Zhu

Department of Entomology , S-225 Agricultural Science Center North, University of Kentucky , Lexington, KY, 40546-0091, USA.

Kun Yan Zhu

Department of Entomology, Kansas State University, Manhattan, KS, 66506, USA ([email protected]).

Junwei Jerry Zhu

Research Entomologist, Agroecosystem Management Research Unit, United States Department of Agriculture, Agriculture Research Service, Room 5, Entomology Hall, University of Nebraska, East Campus, Lincoln, NE 68583-0938, USA ([email protected]).

Yucheng Zhu

Research Entomologist, Southern Insect Management Research Unit, United States Department of Agriculture, Agriculture Research Service, Stoneville, MS 38776, USA ([email protected]).

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Section 1: Insect-Plant Interactions

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CHAPTER 1

Deciphering the Plant-Insect Phenotypic Arms Race Xianchun Li and Xinzhi Ni

Abstract Plants and herbivorous insects interact with each other on three different time scales. On the ecological time scale, the interacting species, both plants and insects, exhibit a back-and-forth attack-defense-counterdefense cycle, resulting in a phenotypic arms race. Such short-term changes in defenses and counterdefenses of individual plants and insects are mediated by reciprocal elicitation and regulation of gene expressions. All reciprocal regulation of gene expression, in turn, are stimulated by chemical or physical signals, from the environment, the organism itself, or the interaction partner. All signals, no matter internal or external, must be received and processed at the level of individual cells. A number of signals that trigger the reciprocal regulation of plant defense or insect counterdefense genes have been characterized. A growing number of microarray studies have been conducted to define the plant defense and insect counterdefense transcriptomes, i.e., genes whose transcription rate is altered by defense-counterdefense interactions. In this chapter, we reviewed the reciprocal signaling and transcriptome dynamic that underlie the plant-insect phenotypic arms race. Keywords herbivore, plant-insect interaction, defense, counterdefense, macroevolutionary time scale, microevolutionary time scale, reciprocal signaling, transcriptome dynamic

Xianchun Li Department of Entomology and BIO5 Institute, University of Arizona, Forbes 410, PO Box 210036, Tucson, AZ, 85721, USA. E-mail: [email protected] Xinzhi Ni Crop Genetics and Breeding Research Unit, USDA-ARS, Tifton, Georgia, 31793, USA. E-mail: [email protected]

T. Liu et al. (eds.), Recent Advances in Entomological Research © Higher Education Press, Beijing and Springer-Verlag Berlin Heidelberg 2011

4

Xianchun Li and Xinzhi Ni

1.1 Introduction Plants and insects have interacted with each other for millions of years (Gatehouse 2002). The interactions between the two taxa can be mutually beneficial (known as mutualism), with insects pollinating (e.g. honeybee) or protecting plants (e.g. Pseudomyrmex ants) from herbivores and plants in return providing foods and/or shelter for insects. The majority of interactions between the two groups, however, are antagonistic, involving insect herbivory of plants and plant defense against the herbivorous insects. Given its importance in ecology, evolutionary biology, and agriculture, the antagonistic plant-insect interaction has been investigated at different biological organization levels, from species (i.e. genomic arms race) to population (genotypic arms race) and individual (phenotypic arms race), and on different time scales, from macroevolution (diversification of plant and insect species) to micro-evolutionary (evolution of novel plant defense and insect counterdefense gene alleles) and ecological time scale (induction of plant defense and insect counterdefense genes). This chapter presents the current understanding of the phenotypic arms race between the interacting plant and insect individuals at the ecological time scale, with emphasis on the plant-insect signaling interactions and the plant defense and insect counterdefense transcriptomes and proteomes.

1.2 Plant-insect phenotypic arms race Plant-insect interactions have been studied at both the evolutionary and ecological time scales. Evolutionarily, the rapid diversification of plant-insect interactions may have been a consequence of reciprocal selection pressures whereby plants evolve biosynthetically novel defensive compounds and proteins, and insects (and other herbivores) overcome erstwhile toxins with novel detoxification pathways (Engler et al. 2000; Zangerl & Berenbaum 2003). As sedentary organisms that form the base of most food webs, plants are subject to intense selection pressure from herbivores, and have few options other than chemical or morphological defense for reducing the impact of herbivores (Berenbaum 1995). Ecologically, the interacting species, both plants and insects, exhibit a back-and-forth attack-defense- counterdefense cycle (Agrawal 2001). The initial herbivore feeding damage provides plants with mechanical signals (wounding) and chemical/enzymatic signals such as volicitin (Weissbecker et al. 1999; Frey et al. 2000), similar fatty acid-amino acid conjugates (FAC) (Halitschke et al. 2001; Tumlinson & Lait 2005), bruchins (Doss et al. 2000), inceptin (Schmelz et al. 2006, 2007) and glucose oxidase (GOX) (Orozco-Cardenas et al. 2001; Musser et al. 2002, 2005) present in the oral secretions (OS) or oviposition fluids (OF) of insects (Fig. 1.1). These signals in turn lead to immediate synthesis and accumulation of plant defense-signaling chemicals including jasmonic acid (JA), salicylic acid (SA), and/or ethylene (ET) (Mello & Silva-Filho 2002; Kessler &

Deciphering the Plant-Insect Phenotypic Arms Race

Fig. 1.1

Structures of six classes of herbivore-associated molecular patterns (HAMP).

5

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Fig. 1.2 Structures of plant defense signaling phytohormones (A) and small peptide hormones (B).Systemin and HypSys are aligned with their conserved central proline (P)- or hydroxyproline (O)-rich motif boxed. Sl = tomato (Solanum lycopersicum), Nt = tobacco (Nicotiana tabacum), ph = petunia (Petunia hybrid), Ib = sweet potato (Ipomoea batatas). Tomato, tobacco, and petunia are members of the Solanaceae family, whereas sweet potato belongs to the Convolvulaceae family.

Baldwin 2002; Schmelz et al. 2003a, 2003b) (Fig. 1.2). The plant defense signaling compounds then switch on the expression of an array of defense proteins involved in the production of defense end products, including allelochemicals, protease inhibitors, indigestible proteins, and volatile organic compounds (VOC) (Paré & Tumlinson 1999; Schmelz et al. 2003a, 2003b; Felton 2005). The volatile semiochemicals may serve to repel herbivores (De Moraes et al. 2001), or to recruit natural enemies as indirect defenses (De Moraes et al. 1998; Thaler 1999; Heil 2008; Dick 2009).


7

In response to plant defenses, herbivores may perceive plant defense signaling molecules (Li et al. 2002a), allelochemicals (Gatehouse 2002; Li et al. 2002b, 2007), and protease inhibitors (Giri et al. 1998; Zhu-Salzman et al. 2003; Moon et al. 2004), to up-regulate their digestive enzymes (De Leo et al. 1998; Cloutier et al. 2000; Mazumdar-Leighton & Broadway 2001), detoxification enzymes including cytochrome P450 monooxygenases (P450s) (Snyder et al. 1995a; Schuler 1996; Danielson 1997; Stevens et al. 2000; Li et al. 2002a, 2002b, 2007), esterases and glutathione S-transferases (GSTs) (Yu & Hsu 1985; Snyder et al. 1995b; Yu 1996, 1999; Ni & Quisenberry 2003), or to recruit more individuals of the same species by releasing aggregation pheromones (e.g. pine beetle) for counterdefense. Thus, individuals of the two species continually adjust their defenses or counterdefenses in response to their interaction partners in a reciprocal fashion that escalates over ecological time (Agrawal 2001). Clearly, signal detection and responses are fundamental to this on-going phenotypic arms race. All reciprocal responses are stimulated by a chemical or physical signal, from the interaction partner, the organism itself, or the environment; and all signals, internal or external, must be received and processed at the level of individual cells. The genetic machinery responsible for perceiving and responding to cues constitutes the defense (in the case of a plant) or counterdefense (in the case of an insect) signaling pathways that channel extracellular information to the genome. The extracellular information then specifically transcribes defense- or counterdefense-related genes into transcripts (transcriptome) to be translated into proteins (proteome). Collectively, the composition and content of the transcriptome and proteome determine the defense or counterdefense phenotypes in the interacting species.

1.3 Signal perceiving and transduction in plants 1.3.1

Herbivore-derived signals

When attacked by herbivorous insects, plants perceive at least two types of signals–mechanical wounding/injury (specific patterns of wounding) and chemical and enzymatic cues present in insect oral secretions (OS) or oviposition fluid (OF), the two fluids from chewing herbivores that commonly come into contact with the wounded plant tissue. Mechanical wounding appears to be a general signal common to all chewing insects, whereas chemical and enzymatic cues are herbivore-specific elicitors (Gatehouse 2002). In parallel with the termpathogen associated molecular pattern (PAMP)-used for pathogen-derived signals or elicitors in plant-pathogen interaction, the chemical and enzymatic signals identified from herbivore OS and OF are also denoted by herbivoreassociated molecular patterns (HAMPs) (Felton and Tumlinson 2008; Mithöfer and Boland 2008). Phloem sap-sucking insects such as aphids and whiteflies have a feeding habit that minimizes tissue damage, and thus reduces or avoids the wounding-elicited response in plants. Instead, they often elicit a plant defense

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response typical of pathogen-induced responses (Walling 2000; Zarate et al. 2007). This makes sense since whiteflies and aphids often act as vectors for plant pathogens. For these homopterans, chemical and enzymatic cues in their OF and OS are probably the only source of signals. Whether the chemical and enzymatic signals that plants perceive from these sucking herbivores are derived from the herbivores themselves, pathogens they vector, or both, remains unknown because no signals have been identified yet from sucking insects. In contrast, six classes of chemical signals (Fig. 1.1) and a few enzymatic signals have been characterized from a number of chewing herbivores. Among the enzymatic or protein HAMPs identified so far are β-glucosidase from the OS of Pieris brassicae larvae (Mattiacci et al. 1995) and glucose oxidase from the OS of Helicoverpa zea larvae (Musser et al. 2002, 2005). β-glucosidase triggers the release of volatiles from cabbage (Brassica capitata) leaves (Mattiacci et al. 1995), whereas glucose oxidase suppresses the wound-induced accumulation of nicotine in tobacco (Nicotiana tabacum) and of trypsin inhibitor in tomato (Lycopersicon esculentum) (Musser et al. 2002, 2005). Meanwhile, glucose oxidase induces the production of the SA-mediated pathogenesis-related protein 1a (PR-1a) in tobacco (Musser et al. 2005). The most well-known class of chemical HAMP are fatty acid (FA)-amino acid conjugates (FACs), represented by volicitin (Fig. 1.1) from Spodoptera exuiga regurgitate (Alborn et al. 1997; Weissbecker et al. 1999; Frey et al. 2000; Shen et al. 2000). FACs are synthesized in the insect gut by conjugation of host-derived FAs to amino acids (Spiteller et al. 2000; Gaquerel et al. 2009) and found in the OS of many lepidopteran larvae, including Manduca sexta (Halitschke et al. 2001), Heliothis virescens and Helicoverpa zea (Mori et al. 2001), Spodoptera littoralis (Maffei et al. 2004) and other lepidopteran larvae (Pohenert et al. 1999; Spiteller & Boland 2003). FACs have been also isolated from three nonlepidopteran species including two closely related cricket species Teleogryllus taiwanemma and Teleogryllus emma and the fruit fly (Drosophila melanogaster) (Yoshinaga et al. 2007). While the amino acid component of FACs is always Gln (Glu in M. sexta), the FA moiety of FACs varies and can be linolenic acid (C18∶3), linoleic acid (18∶2), or derivatives thereof, depending on the food plant (Alborn et al. 1997; Paré et al. 1998; Pohnert et al. 1999; Spiteller and Boland 2003; Spiteller et al. 2004; Mithöfer and Boland 2008). FACs elicit emission of volatiles from some plants such as maize (Alborn et al. 1997) and N. attenuata (Halitschke et al. 2001; Gaquerel et al. 2009), but not lima bean and cowpea (Vigna unguiculata) (Spiteller et al. 2001). FACs are also responsible for eliciting a large portion of the hundreds of genes regulated during the plant-herbivore interaction (Halitschke et al. 2003; Roda et al. 2004) as well as the reconfiguration of the proteome (Giri et al. 2006). The remaining five classes of chemical HAMP are inceptin, caeliferins, 2hydroxyoctadecatrienoic acid, bruchins, and benzyl cyanide (Fig. 1.1). Inceptin is an 11 amino acid peptide resulted from the proteolytic digestion of the cowpea chloroplastic ATP synthase γ-subunit (cATPC) in the midgut of the fall


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armyworm (Spodoptera frugiperda) (Schmelz et al. 2006, 2007). It triggers the production and release of VOC production (Schmelz et al. 2006; Carroll et al. 2008). Caeliferins are saturated and monounsaturated sulfated α-hydroxy fatty acids of 15–20 carbons with their ω-carbon functionalized with either a sulfated hydroxyl or a carboxyl conjugated to glycine via an amide bond (Fig. 1.1) (Alborn et al. 2007; Mithöfer and Boland 2008). Caeliferins were isolated from the OS of the American bird grasshopper (Schistocerca americana) and can trigger VOC emission in maize (Alborn et al. 2007). 2-Hydroxyoctadecatrienoic acid (2-HOT) is a newly identified HMAP from the OS of M. sexta (Gaquerel et al. 2009). It is derived from linolenic acid through the action of the tobacco’s αdioxygenase (α-DOX) in the M. sexta midgut (Gaquerel et al. 2009). It allows tobacco to monitor the progression of the caterpillar's attack and to sustain its production of JA (Gaquerel et al. 2009), the central hormone that coordinates antiherbivore defenses. Benzyl cyanide is a HAMP recently characterized from Pieris brassicae OF (Fatouros et al. 2008). But it is a male-derived antiaphrodisaic pheromone that is transferred to females during mating. In addition to its anti-aphrodisaic role, benzyl cyanide also acts as an elicitor for Brussels sprouts’ indirect defense, and a kairomone for the egg parasitoid Trichogramma brassicae by attracting it to mated P. brassicae females (Fatouros et al. 2005, 2008). Bruchins are long chain α,ω-diols mono- and diesterified with 3hydroxypropanoic acid (Fig. 1.1; Doss et al. 2000; Mithöfer and Boland 2008). They were isolated from the pea weevil (Bruchus pisorum) and cowpea weevil (Callosobruchus maculatus) OF. They can initiate neoplastic growth on pods of certain pea (Pisum sativum) genotypes at the site of egg attachment. This growth lifts the eggs above the oviposition site and thus prevents larval entry into the pod tissue and exposes the neonates to enemies and desiccation (Doss et al. 2000; Mithöfer and Boland 2008). Bruchins can also induce the expression of CYP93C18, a putative isoflavone synthase gene, and the accumulation of the isoflavonoid phytoalexin pisatin (Cooper et al. 2005). 1.3.2

Endogenous plant defense signals

While herbivore wounding and HAMPs described above are the initial signals triggering the escalation of plant defense phenotype, the ultimate activation of plant defense genes and the increased production of the defensive end products (e.g. allelochemicals, toxic proteins, and VOC) are mediated proximally by endogenous plant defense signals produced within seconds to minutes after recognition of herbivore-derived signals. Among the earliest signals detectable are ion fluxes, changes in plasma transmembrane potential (Vm), followed by changes in the intracellular Ca2+ concentration and the generation of hydrogen peroxide (H2O2) and nitric oxide (NO) (Maffei et al. 2007; Wu & Baldwin 2009; Howe and Jander 2008). More proximal signals are plant defense signaling molecules (Fig. 1.2) that are produced within minutes after the onset of insect herbivory (Maffei et al. 2007).

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There are two groups of plant defense signaling molecules (Fig. 1.2). One group are plant peptide hormones (Fig. 1.2B) including proline-rich systemin (Pearce et al. 1991), hydroxyproline-rich glycopeptides (HypSys peptide) (Narváez-Vásquez et al. 2007; Pearce et al. 2001, 2007; Pearce and Ryan 2003) from solanaceous plants and plants outside the Solanaceae family (Chen et al. 2008), and AtPep1 peptide from Arabidopsis (Huffaker et al. 2006; Huffaker and Ryan 2007); all of which are 15–23 amino acids in length and processed from their precursor proteins (Bari and Jones 2009). But HypSys pepetides are unique from Systemin, AtPepe1 and other plant peptide signals, being processed from polyprotein precursors: 2 from a tobacco precursor, 3 from a tomato precursor, and 6 from a sweet potato precursor (Chen et al. 2008). Besides, HypSys peptides are often glycopeptides containing a carbohydrate moiety (Fig. 1.2B). Systemin and HypSys pepetides are included in a functionally-defined Systemin family because they share a common proline or hydroxyproline-rich central core motif (boxed in Fig 2B) and have similar functional roles in defense signaling (Narváez-Vásquez et al. 2007). Systemin and HypSys peptide are found mainly in the Solanaceae family, whereas AtPep1 and its paraglogs in Arabidopsis have orthologs throughout the plant kingdom (Huffaker et al. 2006; Huffaker and Ryan 2007). Another group of plant defense signaling molecules are the well-characterized plant defense signaling hormones JA, ET and SA (Fig. 1.2A). Wounding, HAMPs, necrotrophic pathogens, and feeding by chewing herbivores often result in rapid local and systemic accumulation of JA and ET (Fig. 1.3; Caroline et al. 2007; Glazebrook 2005; Howe and Jander 2008; Maffei et al. 2007; Schmelz et al. 2009; Wu & Baldwin 2009). SA burst, on the other hand, is often induced by biotrophic and semi-biotrophic pathogens as well as phloem-sucking herbivores (Fig. 1.3; Bari and Jones 2009; Glazebrook 2005; Smith et al. 2009; Zarate et al. 2007). But there are some exceptions (Smith et al. 2009), including SA burst elicited by chewing herbivores (e.g. Helicoverpa zea, Bi et al. 1997) and HAMPs (e.g. inceptin, Schmelz et al. 2009) as well as JA burst induced by biotrophic pathogens (Thaler et al. 2004). Recent studies suggest that other phytohormones such as abscisic acid (ABA), auxin, gibberellic acid (GA), cytokinin (CK), and brassinosteroids (BR) are also implicated in plant defense signaling pathways (Bari and Jones 2009; Pieterse et al. 2009). The biosynthesis pathways of the plant defense signaling hormones JA, ET, and SA have been elucidated (Catinot et al. 2008; Ogawa et al. 2006; Wasternack 2007; Wang et al. 2002; Wu and Baldwin 2009). JA is synthesized from chloroplast membrane-derived α- linolenic acid via the octadecanoid pathway that coverts α-linolenic acid to 12-oxo-phytodienoic acid (OPDA) through the chloroplastidial enzymes lipoxygenase (LOX), allene oxide synthase (AOS), and allene oxide cyclase (AOC) in the chloroplast (the filled green circle in Fig. 1.3; Browse and Howe, 2008; Wasternack 2007; Wu and Baldwin 2009). The OPDA is further transformed to JA by OPDA reductase 3 (OPR3) and three steps of βoxidation in the peroxisome (Fig. 1.3; Wasternack 2007; Wu and Baldwin 2009).


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ET is synthesized from S-adenosyl-L-Met through two steps: conversion of Sadenosyl-L-Met to 1-aminocyclopropane-1-carboxylic acid (ACC) by ACC synthase (ACS) and oxidation of ACC to form ET by ACC oxidase (ACO) (Fig. 1.3; Wang et al. 2002; Wu and Baldwin 2009). SA can be synthesized via both the isochorismate pathway and phenylalanine pathway (Fig. 1.3; Catinot et al. 2008; Ogawa et al. 2006). The isochorismate pathway is comprised of the rate-limiting isochorismate synthase (ICS) that converts chorismate to isochorismate, and the isochorismate pyruvate lyase (IPL) that forms SA from isochorismate. The phenylalanine pathway proceeds from phenylalanine via trans-cinnamic acid and benzoic acid (BA) to SA, with phenylalanine ammonia lyase (PAL) catalyzing the conversion of phenylalanine to trans-cinnamic acid, and benzoic acid 2hydroxylase (BA2H) converting benzoic acid to SA (Fig. 1.3; Catinot et al. 2008; Ogawa et al. 2006). The relative importance of the two SA synthesis pathways is still in debate and thus merits further studies. 1.3.3

Plant defense signaling transduction pathway

The signal transduction pathway that connects herbivore-derived signals, wounding and HAMPs (Fig. 1.1), to rapid bursts of plant defense signaling molecules (Fig. 1.2) and the ultimate activation of defense genes/phenotype has being gradually elucidated from intensive research in the model solanaceous (tomato, tobacco) and brassicaceous (Arabidopsis) plants in the last decade. Based on recent excellent reviews (Howe and Jander 2008; Koornneef and Pieterse 2008; Bari and Jones 2009; Pieterse et al. 2009; Wu and Baldwin 2009) and other available models and data (Alonso and Stepanova 2004; Boter et al. 2004; Dong 2004; Du et al. 2009; Durrant and Dong 2004; Fobert and Després 2005; Guo & Ecker 2004; Lorenzo et al. 2004; Pieterse and Van-Loon 2004; Ryan and Pearce 2003; Scheer and Ryan 2002; Xiao et al. 2004; Xie et al. 1998; Yaeno and Iba 2008; Yoo et al. 2008), a comprehensive schematic model of plant defense transduction pathway is proposed here (Fig. 1.3). The whole defense signaling network is comprised of three parallel yet interconnecting phytohormone signaling pathways, namely JA (the left one in Fig. 1.3), ET (the central one in Fig. 1.3) and SA (the right one in Fig. 1.3) pathways. The JA signaling pathway can be turned on by the common mechanical wounding caused by chewing pests and elicitors derived from chewing herbivores (HAMPs) or necotrophic pathogens (PAMPs) (Fig. 1.3; Caroline et al. 2007; Glazebrook 2005; Howe and Jander 2008; Maffei et al. 2007; Schmelz et al. 2009; Wu & Baldwin 2009). Exactly how plants perceive mechanical wounding and elicitors remains elusive. In the model solanaceous plants (tomato & tobacco), wounding inevitably disrupts cellular compartments (e.g., the vacuole). As a result, prosystemin (tomato), proHypSys (tobacco or other solanaceous plants), or PROPE (Arabidopsis), a precursor polypeptide constitutively present at low levels in the cytoplasm (Narvaez-Vasquez and Ryan 2004) or cell wall (Narvaez-Vasquez et al. 2005) , is exposed to proteinases,

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Fig. 1.3 A proposed model showing the activation of the JA, ethylene, and SA signaling pathways in response to wounding, herbivore and pathogen attack. biosynthesis and signaling pathways of JA, SA, and ET) plant defense signaling pathways.Attack by herbivore or pathogen often results in rapid synthesis and accumulation of jasmonic acid (JA), ethylene (ET), or/and salicylic acid (SA), which in turn activate the corresponding JA (the left one), ET (the central one), or SA (the right one) signaling pathways. SA signaling pathway generally activates plant defense responses against biotrophic and hemibiotrophic pathogens. By contrast, JA and ET signaling pathways are usually associated with defense against necrotrophic pathogens and herbivorous insects. Arrows indicate activation or positive interaction, whereas blocked lines indicate repression or negative interaction. Loops that are comprised of green pathway components and green line/curves represent positive signaling amplification loops, whereas loops of red pathway components and red lines/curves represent negative feedback loops. LRR-RLK receptor, leucine-rich repeat receptor-like kinases receptor; MAPKs, mitogen-activated protein kinases; SIPK, salicylic acid-induced protein kinase; LOX, lipoxygenase; LOX-Pi, phosphorylated LOX; AOS, allene oxide synthase; AOC, allene oxide cyclase; OPDA, 12-oxo-phytodienoic acid; OPR3, OPDA reductase 3; JMT, JA carboxyl methyltransferase; MeJA, methyl JA; JAR1, JASMONATE RESISTANT 1; JA-Ile, jasmonoyl-isoleucine; SCF, Skp, Cullin, F-box;COI1, CORONATINE INSENSITIVE 1; JAZ, jasmonate ZIM-domain; Ub/26S, the ubiquitin/26S proteasome; PG, polygalacturonase; PI, protease inhibitors; PDF1.2, PLANT DEFENSIN1.2; ACC, aminocyclopropane-1-carboxylic acid; ACS, ACC synthase; ACO, ACC oxidase; ER, endoplasmic reticulum; ETR1, Ethylene receptor 1; ETR2, Ethylene receptor 2; ESR1, Ethylene response sensor 1; ESR2, Ethylene response sensor 2; EIN4, Ethylene insensitive 4; CTR1, constitutive triple response1; EIN2, Ethylene insensitive 2; EIN5, Ethylene insensitive 5; EIN6, Ethylene insensitive 6; EIN3, Ethylene insensitive 3; EIL1, EIN3-like 1; EBF1/EBF2, EIN3-binding Fbox protein 1 and 2; PERE, primary ethylene response element; ERF1, ETHYLENE RESPONSE FACTOR 1; BAH1/NLA, benzoic acid hypersensitive1 / nitrogen limitation adaptation; CaM, Calmodulin; AtSR1, Arabidopsis thaliana signal-responsive gene 1; EDS1, enhanced disease susceptibility 1; PAD4, phytoalexin–deficient 4; ICS1, isochorismate synthase 1; IPL, isochorismate pyruvate lyase; Phe, phenylalanine; trans-CA, trans-cinnamic acid; BA, benzoic acid; PAL, phenylalanine ammonia lyase; BA2H, benzoic acid 2hydroxylase; NPR1, NONEXPRESSOR OF PATHOGENESIS-RELATED (PR) GENES 1; Why1, whirly 1; AS-1 element, activation sequence 1 element.

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either from the disrupted plant cells or from the insect OS or OF. This leads to proteolytic release of the active plant defense peptide hormones (e.g. Systemin in tomato, HypSys in tobacco, or AtPep1 in Arabidopsis) (Ryan & Pearce 1998; Ryan 2000; Ryan & Pearce 2003; Huffaker et al. 2006). Peptide hormones (e.g. Systemin) then bind to their membrane-bound LRR-RLK (leucine-rich repeat receptor- like kinases) receptors (Scheer & Ryan 2002; Yamaguchi et al. 2006), leading to phospholipase A2-mediated release of α-linolenic acid from chloroplast membrane lipids via a mitogen-activated protein kinase (MAPK) cascade. α-Linolenic acid is converted into JA via the octadecanoid pathway composed of LOX, AOS, and AOC, followed by OPR3 reduction and three βoxidation reactions in peroxisome (Fig. 1.3). Herbivore- or necrotrophic pathogen- derived elicitors further amplify the wounding-elicited JA burst by activating the phospholipase A2-mediated release of α-linolenic acid (JA precursor) from chloroplast membrane or phosphorylating the JA biosynthesis enzymes LOX and AOS through their receptor-activated MAPK cascade (Fig. 1.3). The produced JA is converted to methyl jasmonate (MeJA) by jasmonic acid carboxyl methyltransferase (JMT) for plant-plant communication (Farmer and Ryan, 1990) and/or to the bioactive JA molecule JA-isoleucine (JAIle) by JAR1 (jasmonate resistant 1), which encodes a JA amino acid synthetase (Staswick and Tiryaki 2004; Thines et al. 2007). JA-Ile specifically binds to its putative receptor, the F-box protein coronatine insensitive 1(COI1; Xie et al. 1998; Katsir et al. 2008), leading to the activation of the E3 ubiquitin ligase SCFCOI1 (SKP1/cullin/F-box protein; COI1 is the F-box protein), which targets jasmonate ZIM-domain (JAZ) proteins for ubiquitination and subsequent degradation by the 26S proteasome (1.3; Chini et al. 2007; Thines et al. 2007). JAZ proteins are negative regulators that constitutively repress a set of transcription factors such as the basic helix-loop-helix (bHLH) MYC2, and the ethylene-response-factor1 (ERF1) and ORA59 and inhibit the expression of JAresponsive genes (Lorenzo et al. 2003, 2004; Boter et al. 2004; Chini et al. 2007; Thines et al. 2007). The degradation of JAZ proteins allows the above transcription factors to antagonistically regulate the expression of two groups of JA-induced genes, i.e., the T/G box-containing early-expressed JA signaling (e. g., JAZ, LOX, MYC2, etc.) and late-expressed herbivore defense/ woundresponsive genes by MYC2, and the GCC box-containing necrotrophic pathogen defense genes (resistance to necrotrophic pathogen) by ERF1 and ORA59, leading to elevated defenses against herbivores and pathogens (Boter et al. 2004; Lorenzo et al. 2004; Lorenzo and Solano 2005; Dombrecht et al. 2007; Chini et al. 2007; Thines et al. 2007; Bari and Jones 2009). Elicitors from chewing herbivores and necrotrophic pathogens also activate the ET signaling pathway by phosphorylating and stabilizing the ET biosynthetic enzymes aminocyclopropane-1-carboxylic acid (ACC) synthase (ACS) via their receptor-activated MAPK cascade (Fig. 1.3; Liu & Zhang 2004). If not phosphorylated, ACS will be degraded by the 26S proteasome. The accumulated ET then binds to the sensor domains of a family of 5 endoplasmic reticulum


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(ER)–localized ET receptors [ETR1, ETR2, EIN4 (ethylene insensitive4), ERS1 (ethylene response sensor1), and ERS2], which are constitutively active with their histine kinase domain interacting with the N-terminal domain of the negative regulator constitutive triple response1 (CTR1), a Raf-like serinethreonine kinase (Guo & Ecker 2004; Alonso & Stepanova 2004). Association of CTR1 with the ER-localized ET Receptors is required for the repression of the downstream positive regulators EIN2/EIN5/EIN6, MAPK cascade (MKK9MPK3/MPK6), and the transcription factors EIN3/EIL1 (EIN3-like 1) (Fig. 1.3; Guo & Ecker 2004; Alonso & Stepanova 2004; Yoo et al. 2008). CTR1 can phosphorylate the positive transcription factor EIN3 at Thr 592, promoting the degradation of EIN3 by the E3 ubiquitin ligase SCFEBF1/EBF2 (SKP1/cullin/F-box protein; EBF1 and EBF2 are the F-box proteins) and the 26S proteasome (Yoo et al. 2008). It can also repress EIN3/EIL by inhibiting the positive regulators EIN2/ EIN5/EIN6 (Guo & Ecker 2004; Alonso & Stepanova 2004). Furthermore, it can repress ET signaling by inactivating the MAPK cascade comprising MKK9 and MKP3/MPK6 whose function is to stabilize EIN3 and promote its nuclear translocation by phosphorylating EIN3 at Thr 174 (Yoo et al. 2008). Binding of ET to ER-associated ET receptors inactivates ET receptors, presumably by inducing a conformational change. This in turn leads to the dissociation of the immediate downstream negative regulator CTR1 from the ET receptors and thus the inactivation of CTR1. As a result, the CTR1-mediated repression of the downstream positive regulators including the EIN2- EIN5-EIN6 cascade, the MKK9-MPK3/MPK6 cascade and the transcription factor EIN3 is relieved, leading to the stabilization and nuclear translocation/accumulation of the transcription factors EIN3/EIL1. EIN3/EIL1, probably together with JA-induced transcription factors such as MYC2, promote the expression of the immediate early ET-response genes such as the transcription factor ERF1 by binding to the primary ethylene response element (PERE) in their promoter regions. ERF1 in turn induces the expression of the GCC box-containing necrotrophic pathogen defense genes (Guo & Ecker 2004; Alonso & Stepanova 2004; Yoo et al. 2008). The SA signaling pathway is believed to be activated by the elicitors from biotrophic pathogens and phloem sap-sucking herbivores (Fig. 1.3). Based on two recent studies (Du et al. 2009; Yaeno and Iba 2008), we propose that binding of the herbivore- or biotrophic pathogen-derived elicitors to their receptors activates BAH1/ NLA (benzoic acid hypersensitive1 / nitrogen limitation adaptation), a RING-type ubiquitin E3 ligase (Yaeno and Iba 2008). This in turn leads to the 26S proteasome-mediated degradation of the transcription repressor complex Ca2+/Calmodulin(CaM) /AtSR1 that constitutively represses the expression of the downstream transcription factor enhanced disease susceptibility1 (EDS1) by competing for the CGCG box with unidentified transcription activators in the EDS1promoter (Du et al. 2009). Consequently, EDS1 is expressed, which in turn triggers the EDS1-PAD4 (phytoalexin-deficient4; another positive transcription factor) positive feedback loop whereby EDS1 and PAD4 reciprocally drive the expression of each other (Fig. 1.3; Du et al. 2009).

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EDS1 and PAD4 then promote the expression of the SA biosynthetic enzymes in both the isochorismate (ICS1, IPL) and phenylalanine (PAL, BA2H) pathways, resulting in an SA burst (Fig. 1.3; Du et al. 2009; Durrant and Dong, 2004). SA burst induces the glutaredoxin GRX480-meidated oxidoreduction (redox) change, which reduces the conserved key cysteine residues in the TGA transcription factors and their redox-sensitive co-activator nonexpressor of pathogenesis-related (PR) genes 1 (NPR1) (Mou et al. 2003; Durrant and Dong 2004; Dong 2004; Fobert and Després 2005; Ndamukong et al. 2007; Tada et al. 2008). NPR1 is constitutively present in the cytoplasm as an oligomer formed through intermolecular disulfide bonds. GRX480-mediated reduction of Cys 82 and Cys 216 in NPR1 leads to its monomerization and nucleocytoplasmic localization, which is required for the activation of pathogenesis-related (PR) genes (NPR1 monomer in nucleus) and SA repression of the JA signaling (NPR1 monomer in cytoplasm). Reduction of conserved cysteines in TGA transcription factors enables their interaction with monomeric NPR1, leading to repression of ICS1 and expression of the activation sequence 1 element (AS-1 element)containing genes, including PR genes and SA signaling genes (Fig. 1.3; Wildermuth et al. 2001; Ogawa et al. 2007; Mou et al. 2003; Durrant and Dong 2004; Dong 2004; Fobert and Després 2005). In addition, SA burst can also directly activate the whirly (why) family transcription factors such as why1 and thus induce expression of some PR genes in a NPR1-independent manner (Durrant and Dong 2004). 1.3.4

The features of the plant defense signaling pathway

While the plant defense signaling pathway is described as three (JA, ET and SA) parallel linear pathways (Fig. 1.3), the three pathways actually cross-talk at multiple nodes, forming a signaling network that fine-tunes plant growth and defense in response to plant attackers (Bari and Jone 2009; Koornneef and Pieterse 2008). JA and ET signaling pathways can be triggered by the same signals such as HAMP and elicitors from necrotrophic pathogens and may act synergistically (when both pathways are activated) or antagonistically (when only JA pathway is activated) to modulate plant defense against necrotrophic pathogens and herbivorous insects. The JA-ET interactions are largely mediated by the positive transcription factors ERF1 and MYC2. ERF1 integrates signals from JA and ET via the JA-activated MYC2 (necessary for ERF1’s full expression) and the ET-activated EIN3/EIL1 (always required for ERF1’s expression) respectively (see Fig. 1.3) and functions as a positive regulator of JA and ET signaling for pathogen defense genes (Lorenzo et al. 2003; Bari and Jone 2009; Koornneef and Pieterse 2008). MYC2, on the other hand, induces JA mediated expression of the G box or T/G box-containing wound/herbivore response genes but represses the expression of the GCC box-containing necrotrophic pathogen defense genes (Fig. 1.3; Lorenzo and Solano 2005;


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Dombrecht et al. 2007). When both JA and ET pathway are activated, MYC2 also induces the full expression of ERF1, which in turn promotes expression of the JA-mediated pathogen defense genes. SA pathway is usually associated with plant defense against biotrophic and hemi-biotrophic pathogens and cross talks antagonistically with JA pathway. The key signaling node of the JA-SA antagonism is the redox-sensitive co-activator NPR1, whose redox-mediated conformational transition (oligomer vs. monomer) and nucleocytoplasmic translocation determine which pathway is to be activated. Oligomeric NPR1 negatively regulates SA production during herbivore attack and thus suppress SA/JA cross talk to allow induction of JA-mediated defenses against herbivores (Koornneef and Pieterse 2008). Monomeric NPR1, on the other hand, is required for SA repression of JA signaling (NPR1 monomer in cytoplasm) and the activation of SA-mediated defense against pathogens (NPR1 monomer in nucleus) (Dong 2004; Pieterse and Van Loon 2004; Spoel et al. 2003; 2007; Koornneef and Pieterse 2008; Yuan et al. 2007; Koornneef et al. 2008). But ET burst can render SA repression of JA signaling NPR1 independent (Leon-Reyes et al. 2009). Other signaling nodes modulating the JA-SA antagonism include MYC2, which acts as a negative regulator of SA signaling (Laurie-Berry et al. 2006), and GRX480 and WRKY transcription factors WRKY 62 and 70, all of which are involved in the SA-mediated suppression of JA signaling (Li et al. 2004, 2006; Mao et al. 2007; Ndamukong et al. 2007). Besides, the plant defense signaling network also has the following four features: signaling redundancy, signal amplification loops (positive feedback loop; denoted by green text linked with green arrowed lines/curves in Fig. 3), negative feedback loops (denoted by red text linked with red arrowed lines/curves in Fig. 1.3), and destruction of negative (JAZ in JA signaling and Ca2+/CaM/ AtSR1 in SA signaling) or positive (EIN3/EIL1 in ET signaling) regulators via the 26S proteasome (Fig. 1.3; Ballaré 2009). Signaling redundancy can occur both at the levels of signal perception (different environmental signals trigger similar plant responses) and signaling circuits (the same signal activates parallel response channels) (Ballaré 2009). An example for the former would be the activation of the JA signaling pathway by wounding, elicitors from different chewing herbivores and necrotophic pathogens. An example for the latter would be the activation of the JA, ET, and SA signaling pathway by the same signal inceptin (Schmelz et al. 2009). Targeted destruction of negative or positive regulators via the 26S proteasome allows plants to rapidly escalate their defense against attackers by de-repressing temporally inactivated but otherwise fully functional signaling circuits (Ballaré 2009). Co-existence of both positive and negative feedback loops enables plants to mount a defense response that is commensurate with the intensity and duration of the attack (Howe and Jander 2008). This provides host plants with a mechanism to allocate resources between growth/reproduction and defense against herbivores and pathogens.

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1.4 Signal perception and transduction in herbivorous insects When feeding on plants, herbivorous insects not only provide cues for plants to gear up plant defenses, but also obtain signals from plants to activate their own counterdefense. A great deal of evidence indicates that plant defense signaling hormones (Li et al. 2002a) and plant defense compounds such as toxic allelochemicals and protease inhibitors are signals that insects detect and use to upregulate their counterdefense genes (Li et al. 2002b; Moon et al. 2004; ZhuSalzman et al. 2003). Toxic allelochemicals often induce a number of detoxification enzymes including cytochrome P450 monooxygenases (P450s) (Yu 1982; Schuler 1996; Snyder et al. 1995; Stevens et al. 2000; Danielson 1997; Li et al. 2002b, 2007), glutathione-S-transferases (GSTs) (Yu 1996, 1999; Snyder et al. 1995; Li 2009) and esterases (Yu & Hsu 1985), which metabolize and detoxify these toxic allelochemicals. Protease inhibitors,on the other hand, elicit the overproduction of existing digestive enzymes (De Leo et al. 1998) and expression of protease inhibitor-insensitive digestive enzymes (Cloutier et al. 2000; Mazumdar-Leighton & Broadway, 2001), hydrolyzing enzymes that fragment the inhibitors (Giri et al. 1998; Zhu-Salzman et al. 2003), and even P450s (Moon et al. 2004). Little, however, is known about how insects perceive and transduce these signals into elevated counterdefense phenotypes. Perhaps, the only allelochemical transduction cascade that has been extensively studied in insects is the xanthotoxin response cascade mediating the upregulation of CYP6B1 in Papilio polyxenes, a specialist, and CYP6B4 in P. glaucus, a generalist (Brown et al. 2005; McDonnell et al. 2004; Petersen et al. 2003). Despite divergence in their coding sequences and furanocoumarinmetabolizing capabilities, the promoter sequences of the CYP6B4 and CYP6B1 genes are highly conserved in a number of sequences identified as response elements in other invertebrates and vertebrates. For example, both of the CYP6B1 and CYP6B4 promoter sequences contain an overlapping EcRE/ARE/XRE-xan element (Petersen et al. 2003; Brown et al. 2005; McDonnell et al. 2004). They also contain putative XRE-AhR elements similar to those found in mammalian P450 promoters that are activated by binding to activated aryl hydrocarbon receptor (AhR)–ARNT complexes. Both the overlapping EcRE/ARE/XRE-xan element and the XRE-AhR element are necessary for basal and xanthotoxininducible expression of the CYP6B1 (Petersen et al. 2003; Brown et al. 2005). In comparison, the EcRE/ARE/XRE-xan element is necessary for CYP6B4 induction by xanthotoxin but not for its minimal basal expression (McDonnell 2004). Recently, Brown et al. (2005) showed that Spineless (Ss) and tango (Tgo) proteins, the Drosophila melanogaster homologues of mammalian AhR and ARNT, enhanced basal expression of the CYP6B1 promoter but not the magnitude of its xanthotoxin and benzo[a]pyrene induction. Other components of the xanthotoxin transduction cascade, including transcription factors remains unknown. The only protease inhibitor transduction cascade that has been studied in


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insects is the soybean cysteine protease inhibitor soyacystatin N (scN) response cascade mediating the activation of the scN-insensitive cathepsin B-like cysteine protease called CmCatB in the cowpea bruchid (Callosobruchus maculatus) (Ahn et al. 2007). In the absence of the scN signal, CmCatB expression is negatively regulated by the C.maculatus nuclear receptor Seven-up (CmSvp) through its binding to the two tandem chicken ovalbumin upstream promoter (COUP) elements in the CmCatB promoter. In response to scN-containing diets, the protein level of CmSvp is significantly reduced, leading to the de-repression of the expression of CmCatB (Ahn et al. 2007). More experiments are needed to reveal the scN transduction pathway, including how the protein level of CmSvp is reduced and what directly triggers the process.

1.5 Herbivore-induced plant defense transcriptomes and proteomes Phenotypic changes in defenses and counterdefenses of individual plants and insects are mediated by regulation of gene expression in both organisms. Although some genes (e.g. LOX, VSP, PDF) have been known to be associated with herbivore attack, wounding, JA or SA treatment for an extended period, characterization of a broader transcriptional change in response to herbivore attack in plants has only recently been made feasible by the development of genomic transcript profiling methods. The first microarray study of plant-insect interactions compared the expression of 150 pre-selected defense-related genes in Arabidopsis plants mechanically wounded or challenged with caterpillars of the crucifer specialist Pieris rapae. This revealed a difference between insectattacked or wounded plants, particularly in the expression of dehydrationinducible genes (Reymond et al. 2000). The use of a similar microarray showed that the feeding of the green peach aphid (Myzus persicae) on Arabidopsis leaves (Col-0 ecotype) up-regulated or down-regulated genes involved in oxidative stress (GSTs, superoxide dismutases), calcium-dependent signaling, and pathogenesis-related responses (BGL2, PR-1, hevein-like protein), ethylene biosynthesis genes (ACC oxidase 1), aromatic biosynthesis genes (PAL2, chalcone synthase, tyrosine decarboxylase), and tryptophan biosynthetic pathway genes (anthranilate synthase, tryptophan synthase) (Moran et al. 2002). A study using a cDNA microarray consisting of 2,375 Arabidopsis thaliana genes revealed that JA treatment altered the expression of 371 genes (Schenk et al. 2000). Using a large-scale microarray covering 25%–30% of the Arabidopsis genome (7,200 unique genes), Reymond et al. (2004) compared the Arabidopsis defense transcriptomes in response to a specialist caterpillar, Pieris rapae, and a generalist caterpillar, Spodoptera littoralis. Although there are reported differences between the two species in salivary components, nearly identical transcript profiles were observed. One hundred fourteen genes potentially involved in defense were either induced (111 genes for P. rapae, 112 genes for S. littoralis) or repressed (3 genes for P. rapae, 2 genes for S. littoralis) in response

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to feeding by these two species, including genes involved in pathogenesis, oxylipin metabolism, indole glucosinolate metabolism, detoxification, cell survival, signal transduction and abiotic stress responses (such as dehydrins, touch genes, salt- and senescence-related genes). Moreover, 67%–84% of these caterpillar-responsive genes were controlled, totally or in part, by the JA signaling pathway. Using an oligonucleotide microarrays containing 2,158 unique 70-mer oligos selected from the 3’ end of Arabidopsis cDNAs, Kuśnierczyk et al. (2007) compared transcriptional profiles of three ecotypes (Cvi, Ler, and Ws) of Arabidopsis thaliana plants in response to infestation with a generalist aphid, Myzus persicae, and a cruciferous plant specialist, Brevicoryne brassicae. As in the case of the generalist and specialist chewing caterpillars (Reymond et al. 2004), the overall Arabidopsis defense transcriptomes triggered by this pair of generalist and specialist phloem sap-sucking aphids were similar; they both induced general stress-responsive genes (including pathogenesis related proteins) and genes belonging to octadecanoid and indole glucosinolate synthesis pathways but repressed myrosinases, enzymes hydrolysing glucosinolates in the three ecotypes. Nonetheless, the specialist and generalist aphids caused statistically significant differential regulation of 60 genes in Ws and 21 in Cvi, but none in Ler. The differentially-regulated genes were those encoding jasmonic acid and tryptophan synthesis pathway enzymes, and pathogenesis related protein 1 (PR1). Except for PR-1, which was down-regulated by the generalist M. persicae but upregulated by the specialist B. brassicae in the Cvi ecotype, all other differentiallyregulated genes were either induced or repressed by both aphids, but significantly differed in fold changes (Kuśnierczyk et al. 2007). Taken together, generalist and specialist herbivores appear to elicit more or less similar plant defense transcriptomes as long as their modes of attack or feeding styles are similar. Mode of attack or feeding style is the key factor determining plant defense transcriptome, as evidenced by several whole-genome microarray studies of Arabidopsis defense transcriptomes elicited by pathogens and herbivores that have different modes of attack (De Vos et al. 2005; Kempema et al. 2007; Ehlting et al. 2008). Using the Affymetrix ATH1 whole-genome GeneChips representing approximately 23,750 Arabidopsis genes, De Vos et al. (2005) compared the transcriptional changes of Arabidopsis (Col-01 ecotype) upon attack by a pathogenic leaf bacterium (Pseudomonas syringae pv. tomato), a pathogenic leaf fungus (Alternaria brassicicola), tissue-chewing caterpillars (Pieris rapae), cellcontent-feeding thrips (Frankliniella occidentalis), or phloem-feeding aphids (M. persicae). Although aphid feeding caused virtually no visual symptoms compared with the extensive damage caused by the other four attackers, M. persicae induced the largest number of changes (total 2,181, up 832, down 1,349) (De Vos et al. 2005). P. syringae infection resulted in a similar number of consistent changes (total 2,034, up 1,304, down 730), whereas the number of consistent changes in the other Arabidopsis-attacker combinations was much lower: 186 for P. rapae (up128, down 58), 199 for F. occidentalis (up171, down28), and 151 for A. brassicicola (up 120, down 31). Most of the


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differentially expressed genes were unique for the respective Arabidopsisattacker combinations as the overlap between each pair of attackers was relatively small. With the same microarray platform, Kempema et al. (2007) found that infestation by the B biotype whitefly (Bemisia tabaci), another phloem sapfeeding herbivore, only affected 1,256 genes (700 up-regulated and 556 downregulated), much less than the number of genes regulated by M. persicae (2,181 genes; De Vos et al. 2005). Furthermore, there was only 17% overlap between the genes up- or down-regulated by the two phloem sap-sucking herbivores (Kempema et al. 2007). It turns out that many defenses such as glucosinolate metabolism, HR, and H2O2 that are induced by aphids are not elicited by whiteflies. This is probably because B. tabaci performs fewer cellular punctures that activate JA responses or introduces effectors that suppress JA-dependent defenses (Zarate et al. 2007; Kempema et al. 2007). Likewise, using a 70-mer oligonucleotide microarray covering 26,090 gene-specific elements, Ehlting et al (2008) revealed that feeding by the diamond back moth (DBM; Plutella xylostella), a chewing specialist caterpillar, regulated 2,881 genes (up 1,854, down 1,007), 15 times more than the number of genes affected by P. rapae, which is as DBM a chewing caterpillar specialized to the Brassicacea. Many of the DBM-induced genes fall into ontology groups annotated as stress response, secondary metabolism and signaling. Many genes associated with JA and abscisic acid biosynthesis, signaling, or response were significantly up-regulated in at least one time point, whereas genes associated with gibberellic acid, SA and ET were either predominantly down-regulated (gibberellic acid) or not changed (SA and ET) (Ehlting et al. 2008). A general pattern emerging from these studies is that herbivory often re-configures plant transcriptomes by activating defense related processes while down-regulating primary metabolic processes. Herbivore-induced plant defense transcriptomes have also been studied in genetically less well-characterized plants. Using the native tobacco, Nicotiana attenuata, Ian Baldwin’s laboratory has performed a series of expression studies aimed at charactering and comparing the defense transcriptomes elicited by Manduca sexta, a lepidopteran leaf chewer (Hermsmeier et al. 2001; Halitschke et al. 2003; Hui et al. 2003; Izaguirre et al. 2003), Tupiocoris notatus, a mesophyll and cell content-sucking insect (Voelckel & Baldwin 2004), and M. nicotianae, a phloem-sucking aphid (Voelckel et al. 2004). Due to differences in their feeding habits and in the chemical composition of their oral secretions, these three types of feeders elicited species-specific defense transcriptomes, with M. sexta- and T. notatus-induced transcriptomes being more similar to each other than to M. nicotianae-induced transcriptome. The difference between M. sexta- and T. notatus-induced defense transcriptomes occurs largely in signaling genes and primary metabolism genes. Genes specifically up-regulated by T. notatus include asparagine synthase (tryptophan biosynthesis), α-amylase (hydrolytic starch degradation), flavanone-3-hydroxylase (quercetin synthesis), lignin-forming peroxidase, a WRKY-type transcriptional factor, and NPR1. Genes specifically down-regulated by T. notatus include the large subunit of RUBISCO, a lipid

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transfer protein, MEK2, metallothionins, ubiquitin, and an adenine nucleotide carrier. Compared with the above two herbivores, the phloem-feeding M. nicotianae elicited only weak responses. Aphid-specific changes included the upregulation of glutamate synthase and the down-regulation of a germin-like protein. Although different, these three defensive transcriptomes do share several commonalities. First, a range of signaling pathways are used in response to either attack including the SA-, ethylene-, cytokinin-, and oxylipin (JA) pathways. Second, there is clearly a complex reorganization of gene expression that not only involves genes associated directly with defense, but also those associated with key primary metabolic pathways, photosynthesis, cell wall, carbon and nitrogen metabolism, stress, wounding, and invasion of pathogens. Third, in general, transcripts involved in photosynthesis and/or primary metabolism were downregulated, whereas those playing a role in signaling and secondary metabolism, responding to stress, wounding, and pathogens, and involved in shifting carbon and nitrogen to defense, were up-regulated. These alterations indicate an insectinduced switch from a growth-oriented transcriptional phenotype to one that is defense-oriented. In Sorghum bicolor, another genetically less-characterized plant, suppression subtractive hybridization (SSH) and cDNA microarray techniques were combined to detect transcriptional changes in response to JA, SA, and the phloem-sucking greenbug, Schizaphis graminum (Zhu-Salzman et al. 2004). Of the 672 cDNAs, 82 showed transcriptional changes in response to greenbug feeding, methyl jasmonate (MeJA), or SA application. Proteins encoded by these altered genes function in direct defense, defense signaling, oxidative burst, secondary metabolism, abiotic stress, cell maintenance, and photosynthesis, as well as many having unknown function. Overall, genes involved in direct defense, defense signaling, secondary metabolism, and abiotic stress were upregulated, whereas genes involved in photosynthesis and cell maintenance were down-regulated, consistent with the results obtained from the tobacco-herbivore system (Voelckel et al. 2004; Voelckel & Baldwin 2004) and the coniferherbivore system (Ralph et al. 2006). This study also demonstrated that the SA signaling pathway was highly activated by the greenbug feeding, whereas the JA pathway was only slightly activated. SSH and macroarray approaches were also employed to characterize the chickpea transcriptome in response to the chewing generalist Helicoverpa armigera (Singh et al. 2008). This study revealed that most of the H. armigera-induced transcripts were MeJA and ET regulated. The plant defense phenotype triggered by herbivore attack not only depends on the herbivore-induced changes in the transcriptome, but also on the herbivoreinduced changes in the proteome. Yet our understanding of plant defense proteome is very limiting, as there have been only a few reported proteomic studies (Chen et al. 2005, 2007; Giri et al. 2006; Lippert et al. 2007). Feeding by M. sexta or treatment with M. sexta OS resulted in changes in 90 out of approximately 600 total leaf protein spots, but did not change the expression of about 100 nuclear protein spots in tobacco (N. attenuate) (Giri et al. 2006).


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Among the 90 regulated proteins, in general, proteins that increased were involved in primary metabolism, defense, and transcriptional and translational regulation; those that decreased were involved in photosynthesis (Giri et al. 2006). Feeding on conifer (Sitka spruce) bark tissues by white pine weevils or mechanical wounding induced a series of related small heat shock proteins, other stress response proteins, proteins involved in secondary metabolism, oxidoreductases, and a novel spruce protein (Lippert et al. 2007). Proteomic analysis of the midgut content (Chen et al. 2005) and frass (Chen et al. 2007) of M. sexta larvae fed on tomato (Solanum lycopersicum) plants revealed accumulation of host proteins including JA-induced defense-related PIs, arginase, threonine deaminase (TD), Leu aminopeptidase, and Germin-like protein (oxalate oxidase), Subtilisin-like proteases and other pathogenesis-related proteins, as well as proteins of unknown function (Chen et al. 2005, 2007).

1.6 Plant defense-induced insect counterdefense transcriptomes and proteomes In stark contrast, herbivore counterdefense has been almost exclusively characterized through a traditional single gene approach, focusing almost exclusively on detoxification and digestion enzymes. This is largely due to the lack of microarray resources for insect herbivores. So far, there has been only one microarray study that monitored the transcriptional changes in the cowpea bruchid larvae (Callosobruchus maculates) fed on a diet containing the soybean cysteine protease inhibitor soyacystatin N (scN) (Moon et al. 2004). Of a collection of 1920 cDNAs obtained by SSH with mRNAs prepared from scNadapted and scN-unadapted larvae, ninety four transcript species were responsive to dietary scN. Sixty-three of them were up-regulated by scN, including genes encoding protein and carbohydrate digestive enzymes, detoxification enzymes, and antimicrobial peptides. Among thirty-one down-regulated genes were cytochrome C oxidase subunit, heat shock protein, mucin, serine protease inhibitor, thymidylate synthase and COP9 signalosome complex subunit genes. There has also been only one proteomic study that found significant expressional changes in 20 aphid protein spots (only 14 were functionally identified) when the generalist aphid M. persicae switched host plants from broad beans (Vicia fabaL.) to rapeseed (Brassica napus L, one Brassicaceae) or potato (Solanum tuberosum L, one Solanaceae) (Francis et al. 2006). Ten of the 14 identified proteins were down-regulated (proteins involved in glycolysis, TCA cycle, protein and lipid biosynthesis) while the other four were overexpressed (mainly related to the cytoskeleton).

1.7 Future directions The recent years have seen considerable progress in deciphering the plant-insect

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phenotypic arms race, particularly on the plant side of the plant-insect interactions. These include identification of herbivore-derived signals perceived by plants, elucidation of plant defense signaling network, and definition of plant defense transcriptome and protenome. In stark contrast, much remains unknown on the insect side of the interaction. For example, do herbivores perceive any other plant-derived signals in addition to plant defense signaling hormones, plant allelochemicals, and plant protease inhibitors? How do herbivores perceive and transduce these signals? How do insects respond to plant defense and plantderived signals in terms of their transcriptomes and protenomes? These all are essential questions that need to be addressed to achieve a more complete understanding of the plant-insect phenotypic arms race. Even on the plant side, our understanding of how plants perceive, transduce, and respond to herbivore attacks remains in a relatively primitive state. To date, only a very small number of HAMPs have been identified from chewing herbivores and none from phloem- and cell-content-feeding insects. There is also a crucial need for characterization of HAMP receptors because there are as yet no known receptor-HAMP ligand interactions or explicit direct links between HAMPs and the immediate downstream effectors. Along the same line, the order of those early signaling events such as ion fluxes, Vm changes, changes in the intracellular Ca2+ concentration, phosphorylation cascades and generation of H2O2 and NO that occur before bursts of plant defense signaling hormones remains unknown. JA, ET, SA and other phytohormone signaling pathways clearly involve a complex mesh of interactions to fine-tune plant growth and defense in response to herbivore and pathogen attacks, yet there have been only a few cross-talking nodes identified. While a catalog of plant genes being up- or down-regulated by herbivore attack is identified, the roles of many of these differently-regulated genes in plant defense are still not clear. Further research emphasis should be placed on these topics. Acknowledgements We are grateful to Dr. Richard, O. Musser and Dr. Al Fournier for their helpful comments. We thank Min Zhang for formatting the references. Dr. Li’s research was supported by USDA SCA 58-6602-8-122.

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CHAPTER 2

Insect Herbivory-Inducible Proteins Confer Post-Ingestive Plant Defenses Keyan Zhu-Salzman and Tongxian Liu

Abstract Plants can accumulate a wide variety of compounds in their tissues, constitutively and/or after induction, that confer resistance to herbivorous insects. Applications of microarray and proteomic technologies show a broad array of proteins are involved in plant defense against herbivores. These insect herbivoryinducible proteins may be regulated by multiple signaling pathways through the action of plant hormones such as jasmonic acid, salicylic acid and ethylene, which are tailored to specific insect feeding morphologies and physiologies. Best studied is a group of jasmonate- and feeding-regulated proteins that target and interfere with digestive and absorptive processes of the insect digestive canal, thus playing a critical role in post-ingestive plant defense. This review will highlight recent work and discuss the function of plant proteins in recognition and response to insect herbivory. Keywords hormones

herbivore, coevolution, inducible protein, plant defense, plant

2.1 Insect digestive system Herbivorous insects need to obtain 10 essential amino acids from their host plants to successfully grow and reproduce. The release of peptides and amino acids from dietary proteins is fulfilled by insect digestive proteases, which can be broadly Keyan Zhu-Salzman Department of Entomology, Texas A&M University, College Station, Texas, 77843, USA Vegetable & Fruit Improvement Center, Texas A&M University, College Station, Texas, 77843, USA E-mail: [email protected] Tongxian Liu Key Laboratory of Applied Entomology, Northwest A&F University, Yangling, Shaanxi, 712100 China Texas AgriLife Research, Texas A&M University System, Weslaco, Texas, 78596, USA


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classified into serine, cysteine, aspartic and metalloproteases (Terra and Ferreira 1994). The most abundant leaf soluble protein RuBisCO, one of the major sources of amino acids for herbivorous insects, can be efficiently degraded during passage through insect digestive tracts (Chen et al. 2007). The digestive activity occurs mainly in the midgut where digestive enzymes are released and nutrients are absorbed. The midgut of many insects is lined with a protective layer called the peritrophic matrix (PM). The PM is composed of a chitin and protein matrix and protects the midgut epithelium against food abrasion, toxins, oxidative stress and microorganisms and maintains compartmentalization of digestive enzymes. Insect midguts vary among species in their physico-chemical properties of pH, redox potentials, surfactantcy, oxygen levels, etc. (Johnson and Felton 1996), impacting digestive efficacy. The midgut pH can range from a low of 5.0 to as high as 12.0, and in part, determines the type of protease that may predominate in the digestive tract. Most lepidopteran and dipteran insects have alkaline midguts and use serine proteases to degrade food proteins, while many coleopteran insects have midguts that are slightly acidic and utilize cysteine proteases as their major digestive enzymes. The midgut lumen of some caterpillar species may be close to anaerobic (Johnson and Felton 2000), and O2-dependent oxidases could have minimal function in the guts of these insects.

2.2 Involvement of a broad array of genes in defense against insects In nature, plants battle with herbivorous insects through the coordinated action of multiple defense compounds targeting various vulnerabilities in insects. One common strategy is to produce, constitutively or through induction, defense compounds that negatively affect insects they confront. In plant seeds and various other plant tissues, there are typically protease inhibitors, α-amylase inhibitors, lectins and other proteins. These defense proteins are often regulated through phytohormones in response to wounding, pathogen and herbivore damage as well as other environmental stresses. Jasmonic acid (JA)-regulated proteins play a critical role in plant defense against insects (Felton 2005; Zhu-Salzman et al. 2008). Ethylene (Et) can stimulate as well as block JA signaling (ODonnell et al. 1996; Zhu-Salzman et al. 1998; Stotz et al. 2000). Salicylic acid has been shown to be antagonistic to JA/Et function (Doares et al. 1995; Koornneef et al. 2008), while abscisic acid could synergize the wounding signal (Chao et al. 1999). Coordinated action of plant hormones forms a complex network to regulate a myriad of defense responses (Reymond and Farmer 1998; Koornneef and Pieterse 2008). Advances in functional genomics and bioinformatics have shed some light on the scale and complexity of the interlinked gene sets that are responsible for plant defense against insect pests and pathogens (Schwachtje and Baldwin 2008). DNA microarray and proteomic studies have provided clues to gene networks in response to insect herbivory and other environmental stimuli (Reymond et al.

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Keyan Zhu-Salzman and Tongxian Liu

2000; Moran et al. 2002; Reymond et al. 2004; Voelckel and Baldwin 2004; ZhuSalzman et al. 2004; Chen et al. 2005, 2007; Kempema et al. 2007).

2.3 JA- and feeding-inducible anti-nutritional proteins One important defense signaling pathway involves the defense hormone JA. JA controls the expression of many genes in response to plant damage, but only a few protein products of these genes are known to suppress insect growth and development. Some of these JA-regulated proteins play a critical role in postingestive plant defense against insects (Felton 2005). One key mechanism of these proteins is to target the insect digestive canal and impair its digestive and absorptive processes. However, their anti-insect activities, especially through enzymatic action, are often greatly impacted by the environment in the insect digestive tract. 2.3.1

Protease inhibitors and lectins

Protease inhibitors are often found in high concentrations in plant storage organs or tissues, such as seeds and tubers. They are categorized according to the proteases they inhibit, and inhibitors of all the above mentioned protease classes have been identified in plants (Ryan 1990). Some argue that this accumulation is for defense, as protection of the seed is crucial for success of the species as a whole. More convincing evidence was furnished by the seminal work of Green and Ryan (Green and Ryan 1972) several decades ago, where they demonstrated local as well as systemic induction of an endogenous plant protease inhibitor by insect herbivory in vegetative tissues. A great interest in naturally occurring protease inhibitors was provoked by this ground-breaking work. A large number of feeding experiments directly supported their defensive role against invading microorganisms and herbivorous insects. Protease inhibitor expression is a dynamic process elicited by environmental factors. Their protective function is attributed to the formation of stable complexes between inhibitors and the catalytic clefts of specific proteases. The durable association of these complexes blocks enzymatic action on food proteins, resulting in retardation of growth and development of many insect species. Given the substantial evidence for their post-ingestive anti-insect function, plant protease inhibitors received serious consideration as candidates for biotechnological approaches to confer insect resistance in transgenic plants. However, attempts to use plant protease inhibitors to enhance insect resistance have met with very limited success due to rapid insect adaptation to the transgenic products. Employing non-host inhibitors was thought to be a better strategy, because their nonhost origin would not have imposed previous selection pressure on the target insect (Harsulkar et al. 1999). Direct in vitro molecular evolution has also been proposed to generate highly potent and specific inhibitor molecules (Koiwa et al. 1998). Debilitating insect proteolysis through protease inhibitors in


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host plants and comprehension of their modes of action is certainly a viable means to decrease insects’ access to essential nutrients. Plant lectins comprise another important group of anti-nutritional proteins. They are defined as proteins that possess at least one non-catalytic domain that binds reversibly to specific carbohydrates (Van Damme et al. 1998). While some plant lectins can account for as much as 10% or more of the total soluble protein in seeds and vegetative tissues, others accumulate in response to JA, wounding or other stress treatments (Chen et al. 2002). Lectins that bind N-acetylglucosamine and are resistant to proteolysis often have anti-insect activity, as they may readily bind the chitin components of the PM and/or the glycoproteins that line the digestive tracts, and disrupt their morphology (Chrispeels and Raikhel 1991; Fitches and Gatehouse 1998; Van Damme et al. 1998). Structure disruption may interfere with normal digestive and absorptive functions and predispose the insect to pathogens and toxins. Site-directed mutagenesis that removed carbohydrate binding also caused loss of anti-insect activity (Zhu-Salzman et al. 1998). Thus recognition and binding to sugar moieties of herbivores plays a role in plant defense. 2.3.2

Plant enzymes that serve defense functions

Plants battle with herbivorous insects through the coordinated action of multiple defense compounds targeting various vulnerabilities in insects. Even a single biological strategy aimed at impairing insects’ ability to acquire amino acid nutrients involves a number of proteins working together. This is best illustrated by tomato (Solanum lycopersicum) defense proteins. Although protease inhibitors constitute part of the endogenous defense machinery via their ability to target digestive enzymes, the JA-induced tomato proteins arginase and threonine deaminase (TD2) also disrupt insect digestion (Chen et al. 2005). The two enzymes act catabolically in Manduca sexta midgut, thus depleting the essential amino acids Arg and Thr respectively, providing a synergistic protease inhibitor, anti-nutrient activity. Transgenic plant experiments confirm the antiinsect function. Interestingly, plants deploy two different Thr deaminase forms to meet the needs of Ile biosynthesis (the housekeeping isoform TD1) and postingestive defense (Thr-degradation in the insect midgut by TD2). TD2 (but not TD1) is induced by wounding treatment and its activity is not inhibited by the presence of Ile (Chen et al. 2007). This insensitivity to Ile is due to the herbivoreinduced removal of a C-terminal regulatory domain that confers negative feedback regulation by Ile. The exopeptidase Leu aminopeptidase A may directly interfere with insect gut metabolism and physiology. It is also induced by wounding and exogenous JA (Chao et al. 1999), and remains stable in alkaline environment of the lepidopteran gut (Chen et al. 2005). Its ability to release N-terminal Arg residues from polypeptides may potentiate arginase function to degrade polypeptide-derived Arg in insect guts, impacting insect growth. In addition, some evidence indicates a role in regulation of JA signaling (Walling 2006).

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Polyphenol oxidases (PPOs) exert their anti-nutritive function through catalyzing oxidation of o-diphenols to o-diquinones. Reactive o-diquinones rapidly polymerize and alkylate cellular constituents, leading to cross-linking and covalent modification of proteins, browning of tissues and extracts as well as reduced nutritional quality (Felton et al. 1992). PPOs are widespread in plants and are inducible by wounding, herbivory and JA (Constabel and Ryan 1998). Transgenic plants overexpressing PPO exhibited negative effects on growth and survival of insects (Wang and Constabel 2004). Latent PPO precursors require activation via proteolysis, which occurs within the insect midgut. Activated PPO remains intact and active in the herbivore’s digestive system (Wang and Constabel 2004). A 33-kD maize (Zea maize) insect resistance 1-cysteine protease (Mir1-CP) gene confers resistance to numerous lepidopteran pests. Mir1-CP rapidly and dramatically accumulates at the site of feeding (Pechan et al. 2000). Electron microscopy showed that Mir1-CP makes cracks and holes in the peritrophic matrix (PM), a structure that protects the midgut from toxins and assists in digestion (Pechan et al. 2002). In vitro studies indicated that Mir1-CP rendered the PM permeable in a concentration-dependent manner, which was prevented by E64, indicating that Mir1-CP directly degrades some PM proteins (Mohan et al. 2006). Increasing PM permeability compromises caterpillar defense because digestion of the PM proteins weakens the protective barrier and allows easier access of toxins and microbes to the midgut microvilli, resulting in enhanced toxicity. The glucosinolate-myrosinase complex, the so-called “mustard oil bomb”, is involved in a range of biological activities affecting insect feeding (Bones and Rossiter 1996). In undamaged tissue, plant myrosinase, a β-thioglucosidase, is stored separate from glucosinolates. When plant tissue is damaged, glucosinolates come into contact with plant myrosinase. The enzyme can remove the βglucose from glucosinolates, resulting in the formation of an unstable intermediate product. This product then breaks down to a variety of toxic products, which deter herbivore feeding. The cell- and tissue-specific expression of the myrosinase and the compartmentalization of the components of the glucosinolate-myrosinase system represent a unique mechanism of plant defense. Vegetative storage proteins (VSP) are another defense protein class with enzymatic activity. Although known as temporary reservoirs for amino acids in vegetative tissues in plants (Staswick 1994), expression of Arabidopsis VSP (AtVSP) positively correlated with plant resistance (McConn et al. 1997; Stotz et al. 2000; Ellis and Turner 2001). Also AtVSPs are induced by JA application and insect feeding (Berger et al. 1995; Stotz et al. 2000; Reymond et al. 2004). The anti-insect activity of AtVSP was confirmed by its potent activity against several insect species (Liu et al. 2005). A short sequence motif search prompted the evaluation of AtVSP2 for acid phosphatase activity. Site-directed mutagenesis indicated that indeed phosphatase activity is the basis for its anti-insect function (Liu et al. 2005). Presumably, AtVSP2 interferes with phosphate metabolism in herbivorous insects.


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2.4 Insect adaptation to plant defense As plants evolve defense mechanisms, their predators evolve means to evade them. Besides simply moving to avoid locally induced defenses (Paschold et al. 2007), insects can also avoid activating some defenses by certain salivary components. This has been elegantly demonstrated in Helicoverpa zea (Musser et al. 2002). When the secretory spinneret structure was ablated, salivation was prevented, including secretion of the salivary enzymes. Plants fed on by ablated caterpillars exhibited higher defense activation than those fed on by caterpillars with intact spinnerets. Glucose oxidase, one of the principal components of H. zea saliva, is responsible for the repression of induced defenses in plants. Insects may even “eavesdrop” on the hormones JA or salicylic acid by upregulating their detoxification systems in advance of induced defenses (Li et al. 2002). The plasticity and wide diversity of insect digestive proteases has gradually gained appreciation: they not only break down dietary proteins to meet nutritional requirements, but also play a role in insect counter-defense. Over-consumption and/or adjusting of digestive enzyme complements are alternative strategies insect herbivores use to counter plant defense due to endogenous plant proteases inhibitors (Jongsma et al. 1995; Bown et al. 1997; Jongsma and Bolter 1997; De Leo et al. 1998; Brunelle et al. 1999; Cloutier et al. 2000; Mazumdar-Leighton and Broadway 2001; Brunelle et al. 2004). Modulation of transcripts and protein products of major digestive protease isoforms have been demonstrated in cowpea bruchids Callosobruchus maculatus. Thirty different cDNAs encoding major digestive cathepsin L-like cysteine proteases (CmCPs) have been cloned (ZhuSalzman et al. 2003). Clustering analysis further separated these into two groups, CmCPA and CmCPB, based on sequence similarity. The bruchids selectively expressed CmCPB over CmCPA when fed on diet containing the soybean cysteine protease inhibitor scN. Further study indicated CmCPBs have several advantages over CmCPAs: (i) higher intrinsic proteolytic activity; (ii) more efficient autocatalytic conversion from the latent proenzyme to its active mature protease form; and (iii) exclusive scN-degrading activity, which did not exist in CmCPA (Ahn et al. 2004). The superiority of CmCPB is dependent on efficient propeptide autoprocessing and degradation. Efficiency of these post-translational events directly correlates with varied proteolytic activity of different isoforms (Ahn et al. 2007). Cowpea bruchids apparently make use of this diversity and selectively express CmCPB to cope with suboptimal diets. Promoter analyses have shed some light on transcriptional regulation of insect counter-defense genes (Ahn et al. 2007). However, the mechanisms of sensing dietary challenges and activation of signal transduction for effector gene expression is largely unknown. Insect midguts contain not only numerous isoforms of major digestive proteases, but enzymes that hydrolyze proteins using different catalytic mechanisms. The quantity of the latter is generally small compared to major digestive enzymes under normal conditions (Thie and Houseman 1990; Silva and Xavier-Filho 1991; Liu et al. 2004; Srinivasan et al. 2005; Vinokurov et al. 2006).

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The fact that many insect species have midgut pH gradients, from acidic through neutral to alkaline in separate regions of the gut, facilitates compartmentalized enzymatic function of a particular class of proteases (Oppert et al. 1993; Terra and Ferreira 1994; Edmonds et al. 1996; Vinokurov et al. 2006). The presence of more than one mechanistic class of proteases probably serves to broaden the spectrum of digestible proteins from their host plants. Functional redundancy resulting from multiple digestive enzymes could be a necessity to ensure amino acid supplies. Further, coordination between different classes of digestive enzymes is also required for effective fragmentation of plant defense proteins (Brunelle et al. 1999; Zhu-Salzman et al. 2003). Thus, these normally minor digestive enzymes can help insects, indirectly and directly, to cope with dietary toxins and anti-nutritional compounds they may encounter. However, herbivory-induced, post-translational fragmentation of plant proteins may occasionally backfire on insect protease-based defense. As mentioned earlier, proteolysis within the insect midgut led to the loss of feedback inhibition of TD2 and mediated maturation of latent PPO, resulting in activation of toxicity to insects in both cases (Wang and Constabel 2004; Chen et al. 2007). Another plant defense response triggered by insects’ “innocent mistake” has been elegantly illustrated in the cowpea/fall armyworm interaction (Schmelz et al. 2006). An essential leaf protein, chloroplastic ATP synthase was fragmented by fall armyworm digestive proteases after being ingested. Among the oligopeptides generated, inceptin was perceived by plants as an inappropriately proteolyzed protein fragment, serving as an indicator molecule that signals plants of insect attack. This “guard-based” perception system is responsible for initiation of the specific response in cowpea.

2.5 Phloem feeders interacting with plants Phloem-feeding insects such as aphids and whiteflies impose additional challenges to plants by introducing chemicals that alter plant development and defense and vectoring diseases. These phloem feeders use highly modified mouthparts to navigate the plant tissues and reach phloem sieve tubes with minimal tissue damage. Their unique feeding style could avoid or deter plant defense, even deceiving their host plants by secretion of salivary chemicals and proteins to manipulate plant defense signaling pathways. The silverleaf whitefly (Bermisia tabaci) represses JA- and Et-regulated Arabidopsis genes, the induction of which has been proven important in deterring whitefly development (Kempema et al. 2007; Zarate et al. 2007). Rather, the insect activates the plant’s SA defense pathway, which antagonizes the JA pathway. Suppressing effective defenses and increasing plant susceptibility through manipulation of molecular communication networks has also been observed in aphids and pathogens (Moran et al. 2002; Zhu-Salzman et al. 2004). For instance, greenbug (Schizaphis graminum), the aphid pest on sorghum (Sorghum bicolor) and other small grains,


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drastically induces SA-dependent pathogenesis-related genes. In contrast, only weak induction of MeJA-regulated defense genes was observed after greenbug treatment. Bioassays confirmed that JA-regulated pathways were effective in plant defense against greenbugs (Zhu-Salzman et al. 2004). While this seems to be a common phenomenon among some aphids (Zhu-Salzman et al. 2005; Thompson and Goggin 2006; Walling 2008), exceptions to this evasive strategy have also been observed. Both SA and JA apparently are important for tomato (Solanum lycopersicum) immunity to potato aphids (Macrosiphum euphorbiae) (Li et al. 2006). To date, only one R gene, Mi-1.2 has been molecularly characterized. It mediates resistance to certain species/biotypes of potato aphids, whiteflies, psyllids and nematodes. The mechanisms for resistance are unique for each taxon, but the biochemical basis for resistance and the pest effectors in these incompatible interactions are still unknown (Walling 2008). Further studies are needed to identify plant defense proteins and/or metabolites that confer resistance to phloem feeders.

2.6 Gene stacking The current biotechnology-based insect pest management strategy is to transfer selected resistance genes from one species to another. Constitutive expression of single defensive molecules imposes increased selection pressure, and as a result, can lead to rapid development of new insect biotypes that are insensitive to the transgenic products (Cloutier et al. 2000; Murdock and Shade 2002; Mehlo et al. 2005). These adaptive responses have become the obstacles that hinder efforts in biotechnology-based insect control. Stacking or pyramiding two or more resistance genes offers the potential for providing durable multitoxin resistance to insect pests (Zhu-Salzman et al. 2005; Gatehouse 2008). While some combinations of anti-insect proteins resulted in additive suppression of survival and development of insects, others showed markedly higher activity than the sum of the effect of the individual proteins. For instance, the effect of Bt toxins towards numerous insects was synergistically augmented by a serine protease inhibitor, an endochitinase, a lectin, or by a different Bt toxin (Macintosh et al. 1990; Regev et al. 1996; Maqbool et al. 2001; Cao et al. 2002; Mehlo et al. 2005). Recent studies demonstrated that Mir1-CP synergized Bt-CryIIA activity against several caterpillar species (Mohan et al. 2008). Mir1-CP alone had LC50 values ranging from 0.6 to 8 ppm against these species, the same order of magnitude as those of Bt-CryIIA. The combination of 60 ppb Mir1-CP and 0.5 ppm CryIIA, which showed no effect when used alone, greatly reduced insect growth and increased mortality. The complementary toxicity mechanisms could explain the synergistic effect; Mir1-CP facilitates Bt movement though the PM allowing better access to the epithelium. Thus they are a good combination to be used in transgenic plants.

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Synergistic anti-insect activity has also been demonstrated experimentally by combining various plant anti-insect proteins (Urwin et al. 1998; Inanaga et al. 2001; Oppert et al. 2003; Amirhusin et al. 2004; Outchkourov et al. 2004; Abdeen et al. 2005; Brunelle et al. 2005). One of the mechanisms for synergism was due to the prevention of the degradation of the insecticidal yet biochemically unstable proteins. Computer simulations indicate that, compared to using single genes, the strategy of using two anti-insect genes delays resistance development in insects and reduces the amount of refuge plants required (Roush 1998). Presumably it is more difficult for insects to adapt when they are challenged by two or more proteins with different modes of action than when challenged by only one. Thus, once technical constraints are fully addressed (Halpin 2005), pyramiding multiple defense genes could be very promising in reducing selection pressure on more virulent insect biotypes, and prolonging the useful life of each anti-insect gene.

2.7 Conclusion Through millions of years of co-evolution with herbivorous insects, plants have developed an arsenal of defenses to ward off insect attack, but insect herbivores have evolved means to evade them. Both plant defense and insect counterdefense are multimechanistic. Clearly, the impact of a plant defensive protein is not only determined by its potency toward an insect vulnerable system, but also by how insects respond to such a challenge. A good understanding of the natural defense regulation mechanisms as well as complex insect adaptation mechanisms will no doubt facilitate pest control in modern agriculture. Technology development is helping advance our knowledge base on factors underpinning these complicated interactions, from which new options may be discovered. To successfully use transgenic techniques to protect crops from insect pests, resistance engineered into crop plants must be durable against insect constitutive and induced digestive enzymes, and should involve multiple modes of action. Acknowledgements We thank Drs. Marvin Harris and Ron Salzman for their critical reviews of this manuscript. This work was partly supported by the USDA National Research Initiative grant# 2005-3560415438, USDA Cooperative State Research, Education and Extension Service, grant# 2008-34402-19195.

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Zhu-Salzman K, Salzman R A, Ahn J E, et al. Transcriptional regulation of sorghum defense determinants against a phloemfeeding aphid. Plant Physiology, 2004, 134: 420–431. Zhu-Salzman K, Salzman R A, Koiwa H, et al. Ethylene negatively regulates local expression of plant defense lectin genes. Physiologia Plantarum, 1998, 104: 365–372. Zhu-Salzman K, Shade R E, Koiwa H, et al. Carbohydrate binding and resistance to proteolysis control insecticidal activity of Griffonia simplicifolia lectin II. Proceedings of the National Academy of Sciences of the United States of America, 1998, 95: 15123–15128.

CHAPTER 3

Inducible Direct Defense of Plants Against Insects Mingshun Chen, Junxiang Wu and Guohui Zhang

Abstract Plant defenses against insects can be either direct or indirect. Direct defenses include all plant traits that affect susceptibility of host plants by themselves, whereas indirect defenses include those that attract natural enemies of herbivorous insects, and therefore suppress insect population. Major direct plant defenses include limiting food supply, reducing nutrient value, reducing preference, disrupting physical structures, and inhibiting physiological pathways of the attacking insect. Major known defense chemicals include allelochemicals (se-condary metabolites), inhibitors of insect digestive enzymes, cysteine proteases, lectins, amino acid deaminases and oxidases. Multiple factors with additive or even synergistic impact are usually involved in defense against a specific insect species, and factors of major importance to one insect species may only be of secondary importance or not effective at all against another insect species. High throughput analyses of gene expression, protein abundance and enzymatic activity, and allelochemical concentration will accelerate our understanding of the defense mechanisms, and accelerate identification of specific targets for plant defense enhancement in agriculture. New technologies such as RNA interference will enhance the effectiveness and durability of host plant resistance in crops for insect pest control. Keywords chemical defense, proteases, lectins, protein inhibitors, plant secondary metabolites

Plants are exposed to attacks from numerous herbivores. In response to those attacks, plants have acquired various forms of defenses, some of them are Mingshun Chen Department of Entomology, Kansas State University, USDA-ARS, Manhattan, KS 66506, USA E-mail: [email protected] Junxiang Wu, Guohui Zhang College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China


50

Mingshun Chen, Junxiang Wu and Guohui Zhang

pre-formed (constitutive) while others are induced upon an attack. Constitutive defenses include physical and chemical barriers that exist before an insect attack, whereas inducible defenses include defensive mechanisms that become activated upon attack. Since they incur less metabolic costs to host plants and are more species-specific, inducible defenses have been a focus of research for the last few decades. Three major steps are involved in inducible defense: attack detection, signal transduction, and the launch of specific chemical defenses (Dangl and McDowell 2006; Ferry et al. 2004; Kessler and Baldwin 2002; Walling 2000). Plants detect attacks of herbivores through so-called surveillance system, which can either recognize signals such as effector proteins from herbivores directly, or recognize the perturbation in the plant itself resulted from the herbivore attack. The detected signals are then transduced through a network of signal transduction pathways including receptor kinases, transcription factors, and plant hormones, which eventually lead to the launch of chemical defenses, namely the synthesis and release of chemicals toxic to the attacking herbivore (Smith and Boyka 2006; Kaloshian 2004; Hahlbrock et al. 2003; Wan et al. 2002). Two types of inducible defenses have been observed in plants: direct defenses and indirect defenses. Direct defenses include any plant traits that by themselves affect the susceptibility of host plants to herbivorous insect attacks (Kessler and Baldwin 2002). Indirect defenses, on the other hand, include plant traits that by themselves do not affect the susceptibility of host plants, but can serve as attractants to natural enemies of the attacking insect. Natural enemies suppress the population of the insect, and as a result, reduce plant damage caused by the insect. This chapter will deal only with direct inducible defense in plants against insect herbivores.

3.1 Categories of direct plant defense against insect herbivores Direct plant defenses can be broadly categorized as reducing preference, antinutrition, and toxicity (Fig. 3.1). Reducing preference can be achieved through physical or chemical means such as reducing attractants or releasing repellents. Antinutrition can occur as both pre-ingestion to limit food supply, and post-ingestion to reduce nutrient value to the attacking insect. Major characterized antinutrition defenses include physical barriers at the plant surface or strengthened cell wall, hypersensitive reaction, anti-plant manipulation, removal of essential nutrients, and inhibition of digestive enzymes. Toxicity includes physical damages such as disrupting peritrophic membrane of the insect gut by degradation of its protein components, and chemical disruptions such as inhibition of growth pathways of the attacking insect. The following sections will discuss a few relatively well-characterized categories of plant direct defense. Reducing preference through releasing repellents. Some plants respond to insect herbivory by synthesizing and releasing complex blends of volatile organic compounds (VOCs) that attract natural enemies of herbivores in indirect defenses (Kessler and Boldwin 2001; Kost and Heil 2006). However, some VOCs can also

Inducible Direct Defense of Plants Against Insects

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serve as repellents to the attacking insect as direct defenses. For example, tobacco plants emit herbivore-induced volatile blends that exhibit systematic temporal variation, and some VOCs are released exclusively at night. These nocturnal VOCs repel tobacco budworm (Heliothis virescens) female moths from oviposition on previously damaged plants or nearby undamaged plants (DeMorae et al. 2001). Some VOCs emitted after insect feeding can serve as repellents to the attacking insect itself as a direct defense, as well as attractants to the natural enemies of the attacking insect as indirect defenses (Kessler and Baldwin 2001). The production of VOCs is activated by elicitors from oral secretions of the attacking insect herbivore (Truitt et al. 2004; Schmetz et al. 2006).

Fig. 3.1

Categories of direct defense in plants against insect herbivores.

Direct plant defenses can be broadly divided into antinutrition, repellent, and toxicity. Antinutrition can be mechanisms of limiting food supply that occur before food ingestion, and mechanisms of reducing nutrient value that occur after food ingestion. Toxicity can be caused by either disrupting physical insect structures such as peritrophic membrane or inhibiting physiological pathways that affect insect growth. Limiting food supply via physical barriers. In the pre-ingestion phase, host plants can limit food supplies to insects via a range of mechanisms. A few examples of these mechanisms include induced physical barriers on the surface of plants or via cell wall fortification. A good example of physical barriers is plant trichomes. Many plants have trichomes on plant surfaces as a pre-formed defense. However, the density of trichomes can be increased following mechanical

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damage or insect attack (Traw and Dawson 2002; Traw and Bergelson 2003; Shanower 2004; Gonzales et al. 2007). Plants can limit food supplies to insects by fortifying cell walls to enhance mechanical barriers to insect probing and feeding, especially for insects with sucking mouthparts (Goussain et al. 2005; Vilaine et al. 2007). Plant trichomes can prevent herbivores from reaching the surface or reduce insect movement. Another type of physical barriers is fortified cell walls. Often cell wall of plants is strengthened following insect attack. Fortified cell wall makes it more difficult for insects to access cell content or penetrate into phloem (Goggin 2007). Fortified cell wall can also increase the indigestibility of plant tissues to insects with chewing mouthparts (Clissold et al. 2004). Many genes involved in cell wall fortification are up-regulated by insect attack, including pectin esterases, expansins, cellulose synthases, and xyloglucan endotransglycosylases/hydrolases (XTH) (Goggin 2007; Liu et al. 2007). The importance of cell wall-fortification in plant defense can be seen from the fact that a mutation in an XTH gene in Arabidopsis (xth33) renders plants more susceptible to the green peach aphid Myzus persicae (Vilaine et al. 2007). Limiting food supply via hypersensitive response. Hypersensitive response (HR) is a major form of resistance in plants against pathogens (Goodman and Novacky 1994). HR involves the rapid death of cells at the infection site, which prevents further spread of the pathogen, causing it to starve to death (Heath 2000). HR is also a type of defense mechanisms of plants against some insect herbivores, especially for insects with a fixed feeding site such as galling insects (Fernandes and Negreiros 2001; Cornelissen et al. 2002; Larsson and Wingsle 2005; Moloi and van der Westhuizen 2006). For example, HR was observed in many willow tree varieties (Salix viminalis) with resistance to the gall midge Dasineura marginemtorquens (Hoglund et al. 2005; Larsson and Wingsle 2005). HR is also associated with wheat resistance to the Hessian fly Mayetiola destructor (Grover 1995), rice resistance to the Asian rice gall midge Orseolia oryzae (Bentur and Kalode 1996), peach resistance to aphids (Sauge et al. 1998), and bean resistance to the bean-pod weevil Apion godmani (Garza et al. 2001). HR is caused by a burst of reactive oxygen species (ROS) produced locally in plants attacked by pathogens or insects. In addition to causing HR that limits food supply to herbivores, ROS is also extremely toxic to herbivores directly (Liu et al. 2009). Preventing manipulation of plants by insects. Anti-manipulation of host plants by insectsis an important mechanism of antinutrition for galling insects as well as for other herbivores that modify local environments to obtain nutrition (Agrios 1997; Danks 2002; Rohfritsch 2005). Galling insects attack susceptible plants by creating a zone of “metabolic habitat modification” within a plant tissue (commonly referred to as a gall) (Goethals et al. 2001). In this zone (gall), the insect experiences a selective advantage because of enhanced nutrition (due to the formation of a nutrient sink) and reduced plant defense (Stone and Schönrogge 2003; Harris et al. 2006; Zhu et al. 2008). Resistant plants can somehow prevent


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the formation of a gall, and therefore deprive the attacking insect of nutrition (Harris et al. 2006; Zhu et al. 2007). Reducing nutrient value. In the post-ingestion phase, antinutrition is to reduce nutrient value by removing essential nutrients. Many plant proteins are very stable in the insect gut after they are ingested by insect herbivores (Chen et al. 2007). Some plant enzymes remain active in the insect gut and destroy nutrients that could otherwise be used by the insect. Like humans, some of the amino acids such as arginine, lysine, methionine, and threonine, cannot be synthesized by insects, and therefore must be obtained from the diet. If these essential amino acids are destroyed by plant enzymes in the insect gut, the development of the insect will be impaired. Many amino acid degradation enzymes are induced by insect feeding, and remain stable and active after they are ingested into the insect gut. For example, arginase and threonine deaminase, which degrade arginine and threonine respectively, are strongly induced by insect feeding (Chen et al. 2004; Chen et al. 2005). These two enzymes remain active in the insect gut and are thought to be antinutritional by removing essential amino acids arginine and threonine (Felton 2005; Chen et al. 2007). Asparaginase, another amino acid degradation enzyme, is up-regulated specifically in resistant plants during insect attack (Liu et al. 2007). Inhibition of the herbivore’s digestive enzymes is another way to reduce nutrient value. Upon attack by insect herbivores, plants produce various protein inhibitors that can inhibit insect digestive enzymes such as proteases and amylases (Green et al. 1972; Koiwa et al. 1997). In addition, many plant secondary metabolites also have effects on food ingestion and assimilation, potentially via inhibition of insect digestive enzymes (Duffey and Stout 1996; Akhtar and Isman 2004). Inhibition of insect digestive enzymes reduces the value of ingested nutrients, and therefore impairs insect growth and development. Disrupting physical structures. In addition to antinutrition, plants can also defend themselves by producing chemicals that causes physical damage to the insect. For example, plants can over-produce proteases upon insect attack, which can digest insect structural proteins after they are ingested into the insect gut, causing physical damage to the herbivore. A good example is a 33-kDa cysteine protease (a papain-like protease) in maize (Pechan et al. 2000). This protease accumulates rapidly at the insect feeding site. After the protease is ingested into the insect gut, it digests gut proteins and damages the peritrophic matrix of caterpillars, resulting in inhibition of insect growth and development. Inhibiting physiological pathways. Similarly, defense chemicals can also cause chemical damage to insect herbivores. Insect growth and development are regulated by various pathways involving kinases, phosphatases, and regulatory proteases. Some plant chemicals are inhibitors of these regulatory enzymes. For example, some secondary metabolites can inhibit kinase or phosphatase activity of insects after they are ingested (Salunke et al. 2005; Hertiani 2007). The inhibition of these regulatory enzymes will cause chemical damage, resulting in inhibition of insect growth and development or death of insects.

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3.2 Major characterized chemicals toxic to insect herbivores Research on plant defense to herbivores has been more focused on attack detection and signal transduction. Due to the complexity of the composition of defense chemicals that cause direct toxicity to insects, not many specific chemicals have been well characterized and confirmed. Different plant species often use different chemicals as direct defense in response to attacks from different herbivores. Even for the same plant species, different tissues or even the same tissue at different developmental stages face attacks from different insects with different mechanisms of attack. To defend themselves effectively against different insects, plants have to develop corresponding mechanisms against insects with different mechanisms of attack. Considering the vast numbers of plant and insect species, it is not hard to imagine the complexity and the diversity of specific defense mechanisms associated with different plant-insect systems. This section will summarize major defensive chemicals that have been characterized in various plant-insect systems, including allelochemicals, inhibitors of insect digestive enzymes, proteases, lectins, amino acid deaminases, polyphenol oxidases, and other defensive proteins. In many cases, however, supporting evidence is still very preliminary. Allelochemicals. Most allelochemicals are plant secondary metabolites (PSMs) that are not essential for normal plant growth and development, and are often produced as by-products during the synthesis of primary metabolic products (Herbert 1989). Some PSMs perform physiological functions within the plant such as serving as a tool for transport and storage of nitrogen, as UVprotectants, and as attractants for pollinating and seed-dispersing animals (Wink 1999). The main function of PSMs, however, is thought to serve as chemical defenses against pathogens and herbivores (Bennett and Wallsgrove 1994; Theis and Lerdau 2003; Mao et al. 2007). A great deal of PSMs is toxic to herbivores unless detoxification mechanisms are developed. Even for those herbivores that have developed detoxification mechanisms, the resources invested in detoxification may in turn incur growth and development costs. The total number of PSMs for which structures have been elucidated is around 50,000, and this is likely only a small fraction of all PSMs existing in nature (De Luca and St Pierre 2000). Because of the cost to produce PSMs, each plant species produces only a small, but unique combination of these compounds. For example, cyclic hydroxamic acids are almost exclusively produced in cereal crops (Gramineae) (Niemeyer 1988), whereas saponins (glycosylated triterprenes and steroids) are widespread in dicotyledonous plant species, but deficient in most grass species (Hostettmann and Marston 1995). In addition, plants have also developed mechanisms to produce certain defense PSMs only upon herbivore attacks or after receiving warnings from neighboring plants (Baldwin et al. 2006). Despite the vast number of PSMs and extensive literature on potential defense roles of many PSMs through bioassays, definitive proof of the defense function of


55

a particular PSM remains to be determined in most cases. Table 3.1 provides a list of PSMs that have some concrete evidence on their defense roles against insect herbivores. The most convincing case is from transgenic plants with dhurrin, a cyanogenic glucoside (Tattersall et al. 2001). Dhurrin is not normally found in any brassicaceous plants such as Arabidopsis thaliana. When dhurrin was produced in A. thaliana through genetic engineering, transgenic plants with dhurrin gained compelete resistance to the flea beetle Phyllotreta nemorum. Table 3.1 Major known defensive plant secondary metabolites (PSMs). Name (Chemical Class)

Mode of Action

Plant/Insect System

Main Reference

Tannins (polyphenol)

?

Oak/Multiple insects

Forkner et al. 2004

Maysin and apimaysin (flavones)

?

Corn/Corn earworm (Helicoverpa zea)

Lee et al. 1998

Isoorientin (flavonoid)

Toxicity

Corn/Corn earworm (Helicoverpa zea)

Widstrom et al. 1998

Various flavonoids

Toxicity (Ca2C-ATPase inhibitor?)

Roostertree/Bean weevil (Callosobruchus chinensis)

Salunke et al. 2005

Glyceollin (isoflavonoid)

?

Soybean/Coleopterans

Fischer et al. 1990

Lignin

Antinutrition?

Aspen/Gypsy moth (Lymantria dispar)

Brodeur-Campbell et al. 2006

Sinalbin (thioglucosides)

Repellent

Crucifers/Multiple insects

Borek et al. 1998

Sinigrin (thioglucosides)

Repellent

Crucifers/Multiple insects

Giamoustaris et al. 1995

Dhurrin (cyanogenic glucoside)

Toxicity (hydrogen cyanide, HCN)

Arabidopsis/Flea beetle (Phyllotreta nemorum)

Tattersall et al. 2001

DIMBOA (hydroxamic acid)

Toxicity and Repellent

Corn/Multiple insects

Niemeyer et al. 1988

DIBOA (hydroxamic acid)

Toxicity and Repellent

Rye/Multiple insects

Gianoli et al. 1999

Monoterpenes and diterpenes

Toxicity

Spruce/White pine weevil (Pissodes strobi)

Duffey and Stout 1996

Saponin Tomatine (glycosylated triterpenes)

Toxicity (Disrupting cell membrane)

Tomato/Multiple insects

Duffey and Stout 1996

Sesquiterpenes

Repellent

Tobacco/Multiple insects

DeMoraes et al. 2001

Indolyl glucosinolates

?

Arabidopsis/Aphids

Kusnierczyk et al. 2008

camalexin

?

Arabidopsis/Aphids

Kusnierczyk et al. 2008

Defense PSMs belong to three categories: phenolics (such as tannins, flavones, isoorientin, flavonoids, glyceollin, and lignin), nitrogen compounds (such as

56


sinalbin, sinigrin, dhurrin, DIMBOA, and DIBOA), and terpenoid (such as Saponin). Different PSMs suppress herbivore’s growth and development via different mechanisms. Some flavonoids are inhibitors of regulatory enzymes such as calcium-dependent ATPase (Salunke et al. 2005). Lignin and other phenolics can strengthen cell walls and therefore can be antinutritional (Brodeur-Campbell et al. 2006; Schroeder et al. 2006). Some phenolics and sesquiterpenes along with other volatiles can repel herbivores from oviposition on host plants (Henzell and Hall 1974; DeMoraes et al. 2001; Huber et al. 2006). Inhibitors of Insect Digestive Enzymes. Inhibitors of insect digestive enzymes are the most extensively studied group of anti-insect chemicals. There are two major kinds of digestive enzymes in insect digestive systems: proteases and amylases (Terra et al. 1996). Inhibitors of both proteases and amylases are widely distributed in plant tissues, especially in seeds (Horiguchi and Kitagishik 1971; Feng et al. 1991; Zavala et al. 2004). Digestive enzyme inhibitors as a defense mechanism were first discovered by Ryan and his associate (Green and Ryan 1972). They found that the expression of protein proteinase inhibitors was rapidly induced in potato and tomato leaves in response to insect attacks, and the induction was mediated by a systemic signal, which could also be activated by mechanical wounding. Since then, the roles of digestive enzyme inhibitors in plant defense have been extensively studied (reviewed by Ryan 1990; Ussuf et al. 2001; Rawlings et al. 2005). Transgenic plants with a range of protease or amylase inhibitors have been developed, and in most cases resulted in detrimental effects on insect growth and development (Table 3.2).

Table 3.2 Proteinaceous inhibitors with anti-insect activity in transgenic plants. Source and Name of Inhibitor Transgenic Plants

Targeted Herbivore

Reference

Cowpea trypsin inhibitor

Tobacco

Tomato hornworm, Manduca sexta

Hilder et al. 1987

Tobacco trypsin protease inhibitor

Tobacco


Zavala et al. 2004

Potato proteinase inhibitor 1

Tobacco


Johnson et al. 1989


Tobacco

Cluster caterpillar, Spodoptera litura

Sane et al. 1997

Soybean Kunits inhibitor

Tobacco


McManus et al. 1999

Sweet potato trypsin inhibitor Tobacco


Yeh et al. 1997


Tobacco

Egyptian cotton worm, Spodoptera littoralis

Marchetti et al. 2000

Potato trypsin inhibitor

Tobacco

Green looper, Chrysodeixis eriosoma

McManus et al. 1994


57 (Continued)

Source and Name of Inhibitor Transgenic Plants

Targeted Herbivore

Reference


Tobacco

American boll worm, Helicoverpa armigera

Charity et al. 1999


Potato

Tomato moth Lacanobia oleracea

Gatehouse et al. 1997


Potato

Egyptian cotton worm, Spodoptera littoralis

Marchetti et al. 2000


Cotton


Li et al. 1998

Manduca (insect) protease inhibitor

Cotton

Sweetpotato whitefly, Bemisia tabaci

Thomas et al. 1995


Apple

Multiple insects

James et al. 1993

Tobacco mulit-domain inhibitor

Apple

Light-brown apple moth, Epiphyas postvittana

Maheswaran et al. 2007

Soybean Kunitz proteinase inhibitor

Poplar

Polyphagous moth, Lymantria dispar

Confalonieri et al. 1998

Soybean Kunitz proteinase inhibitor

Poplar

Poplar tip moth, Clostera anastomosis

Confalonieri et al. 1998

Rice cystein protease inhibitor

Poplar

Poplar Leaf-Beetle, Chrysomela tremulae

Leplé et al. 1995


Strawberry

Multiple insects

James et al. 1993


Strawberry

Vine weevil, Otiorhynchus sulcatus

Graham et al. 1997

Sweet potato trypsin inhibitor

Cauliflower

Pieris conidia

Ding et al. 1998

Sweet potato trypsin inhibitor

Cauliflower

Diamondback Moth, Plutella xylostella

Ding et al. 1998


Cauliflower

Small cabbage white butterfly, Pieris rapae

Lu et al. 2005


Pea


Charity et al. 1999

Common bean αamylase inhibitor 1

Pea

Pea weevil, Bruchus pisorum

Morton et al. 2000

Common bean α-amylase inhibitor 1

Pea

Cowpea weevil, Callosobruchus maculatus

Shade et al. 1994


Pea

Chinese bean weevil, Callosobruchus chinensis

Shade et al. 1994


Chickpea


Sarmah et al. 2004

58

Mingshun Chen, Junxiang Wu and Guohui Zhang (Continued)

Source and Name of Inhibitor Transgenic Plants

Targeted Herbivore

Reference


Chickpea


Sarmah et al. 2004


Azuki bean


Ishimoto et al. 1996


Azuki bean




Azuki bean

Graham bean weevil, Callosobruchus analis


Potato proteinase inhibitor

Rice

Rice stem borer, Chilo suppressalis

Bu et al. 2006

Winged bean trypsin inhibitor

Rice


Mochizuki et al. 1999


Rice


Xu et al. 1996


Rice

Pink stem borer, Sesamia inferens

Xu et al. 1996

Potato proteinase inhibitor 2

Rice

Pink stem borer, Sesamia inferens

Duan et al. 1996

Soybean Kunitz trypsin inhibitor

Rice

Brown planthopper, Nilaparvata lugens

Lee et al. 1999

Barley trypsin inhibitor Cme

Rice

Rice weevil, Sitophilus oryzae

Alfonso-Rubi et al. 2003


Wheat

Angoumois Grain Moth, Sitotroga cerealella

Bi et al. 2006

Barley trypsin inhibitor Cme

Wheat

Angoumois Grain Moth, Sitotroga cerealella

Altpeter et al. 1999

Nightshade serine proteinase inhibitor

Tobacco

Cotton bollworm, Helocoverpa armingera

Luo et al. 2009

Nightshade serine proteinase inhibitor

Tobacco

Tobacco cutworm, Spodoptera litura

Luo et al. 2009

Despite the detrimental effect on insect pests, inhibitor-transgenic plants have not been widely adapted in commercial field crops. One of the reasons may be due to the availability of more potent insecticidal agents (such as Bt toxin) available. The other reason is that herbivores can adapt to inhibitor defenses by maintaining diverse digestive enzymes (Ahn et al. 2004; Zhu et al. 2005) and by overexpressing the enzymes that are insensitive to inhibitors when sensitive enzymes are inhibited (Jongsma and Bolter 1997; Koiwa et al. 1997; Jongsma et al. 1995). To develop crop plants with more durable resistance, scientists are engineering inhibitors that inhibit multiple digestive enzymes (Brunelle et al. 2005).


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Proteases. Emerging evidence also suggests that proteases, particularly cysteine proteases, are involved in plant defense against insect herbivores as well (Table 3.3). A good example among proteases is a 33-kDa cysteine protease (a papain-like protease) in maize (Pechan et al. 2000). This protease accumulates rapidly at the insect feeding site and is part of the corn resistance to Lepidopteran insect pests. Overexpression of the 33-kDa protease in maize callus used for feeding experiments can retard insect growth by 80% (Pechan et al. 2002). The toxicity mechanism of this enzyme is to disrupt the peritrophic matrix of caterpillars, thus impairing the insect digestive system. Cysteine proteases may also play an important role in papaya trees against herbivorous insects (Konno et al. 2004). Papaya leaves exude latex when attacked by herbivores. Contained in latex are various cysteine proteases including papain, bormelain, and ficin. Table 3.3 Plant defensive proteases, lectins, amino acid deaminases, and oxidation enzymes. Name

Mode of Action

Plant/Insect System

Main Reference

33-kD Toxicity (Disrupting Cysteine proteinase gut membrane)

Corn/Lepidopterans

Pechan et al. 2000

Papain Toxicity? (Cysteine protease)

Papaya tree /Lepidopterans

Konno et al. 2004

A leucine aminopeptidase

Tomato/Lepidopterans

Pautot et al. 1993

Tobacco/Aphids

Dutta et al. 2005

Wheat/Hessian fly (Mayetiola destructor)

Williams et al. 2002

Proteases

Antinutrition?

Lectins and Lectin-Like Proteins Allium sativum leaf lectin

Toxicity (Surface carbohydrate binding)?

Toxicity Jacalin-like lectins (Surface carbohydrate binding)? Chitin-binding lectin


Brassica/Cabbage aphid

Cole 1994

Agglutinin


Tobacco /Peach-potato aphid (Myzus persicae)

Chang et al. 2003

Talisia esculenta Lectin


Cowpea/Bruchid weevil (Callosobruchus maculatus)

Macedo et al. 2004

Jackbean lectin


Bean/Pea aphid

Sauvion et al. 2004

Snowdrop Lectin


Multiple crops /Multiple insects

Wang et al. 2005

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Mingshun Chen, Junxiang Wu and Guohui Zhang (Continued)

Name

Mode of Action

Plant/Insect System

Main Reference

Amino acid deaminases Arginase

Antinutrition (Degrading arginine)


Chen et al. 2005

Threonine deaminase

Antinutrition (Degrading threonine)


Kang et al. 2006

Asparaginase

Antinutrition (Degrading asparagine)

Wheat/Hessian fly (Mayetiola destructor)

Liu et al. 2007

Toxicity Cholesterol oxidase (Disrupting gut membrane)

Cotton/Boll weevil

Estruch et al. 1997

Polyphenol oxidase Antinutrition

Multiple crops /Multiple insects

Thipyapong et al. 2007

Top of Form Ascorbate peroxidase

Antinutrition

Multiple crops /Helicoverpa zea

Felton et al. 1993

Lipoxygenase

?

Artificial diet

Shukle and Murdck 1983

Oxidation enzymes

Lepidopteran larvae die when fed on fig leaves that contained latex, but not when the latex is removed from these leaves by washing or when the cysteine proteases are inactivated by E64, a potent cysteine protease inhibitor. In laboratory experiments, artificial diets containing either papain, bromelain, or ficin in concentrations that occur in latex are toxic to silkworm larvae. These observations suggest that proteases are a vital part of the defense arsenal in latex exuded out of wounded sites in response to insect feeding. Other types of proteases may also involve in direct defenses against herbivorous insects. A leucine aminopeptidase is strongly up-regulated in response to insect feeding, pathogen attack, and mechanical wounding (Pautot et al. 1993). The protein is resistant to insect digestive enzymes once ingested into insect gut (Chen et al. 2007). Leucine aminopeptidase liberates arginine from the N-terminus of peptides. Arginine, an essential amino acid to insects, can be further degraded via arginase, which is also induced by insect feeding (see below). Therefore, leucine aminopeptidase may cause detrimental effects on insect herbivores by removing essential nutrients in conjunction with arginase and other amino acid catabolic enzymes. In addition to direct toxicity, proteases are widely involved in various signal transduction cascades of plant defense (van der Hoorn and Jones 2004). Considering the complexity of plant proteolytic machinery (at least 488 protease genes within the Arabidopsis genome, http://merops.sanger.ac.uk), the roles of various proteases in plant defense are still waiting to be discovered.


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Lectins. Lectins are a very diverse group of proteins that reversibly bind to specific mono- or oligosaccharides and are widely distributed in microbial, plant, and animal tissues (Komath et al. 2006). Lectins incorporated in artificial diets have been shown to result in reduced performance of several insect species (Janzen et al. 1976; Shukle and Murdock 1983; Murdock et al. 1990; Powell et al. 1993; Peumans and van Damme 1995; Rahbé et al. 1995; Sauvion et al. 2003; Subramanyam et al. 2008). One of the most widely studied, and perhaps one of the most promising chemicals for aphid control, is snowdrop lectin. The particular interest in this protein is based on the fact that it acts on sap-sucking insect pests that are not targeted by the known Bt-toxins (Rahbe et al. 1995; Sauvion et al. 1996; Powell et al. 1993, 1995, 1998; Powell 2001). Transgenic plants that express snowdrop lectin offer partial resistance to homopteran pests, as has been observed in tobacco (Hilder et al. 1995; Yuan et al. 2001), potato (Down et al. 1996; Gatehouse et al. 1996), rice (Rao et al. 1998; Foissac et al. 2000; Sun et al. 2002), and wheat (Stoger et al. 1999). In addition to effects on sap-sucking pests, insecticidal effects of snowdrop lectin were also found to lepidopteran pests (Fitches et al. 1997; Gatehouse et al. 1997; Irvine and Mirkov 1997; Legaspi et al. 1997; Fitches and Gatehouse 1998; Setamou et al. 2002, 2003) and a coleopteran herbivore (Nutt et al. 1999). The insecticidal mechanism of lectins is not clear at present. Since lectins interact with mono- and oligosaccharides, the insecticidal activity may involve a specific carbohydrate-lectin interaction with glycoconjugates on the surface of digestive tract epithelial cells (Macedo et al. 2004). Lectins are resistant to insect gut proteolysis and can form complexes with gut proteins (likely glycosylated proteins) with high affinity (Gatehouse et al. 1995; Macedo et al. 2004; Sauvion et al. 2004). Amino acid deaminases. Amino acid deaminases are another group of antinutritional proteins in plant defense against insect herbivores. These enzymes can degrade free amino acids in the insect gut, removing nutrients from the herbivore (Chen et al. 2005). Two well characterized amino acid deaminases are arginase and threonine deaminase, both of which are induced by insect attacks and wounding (Hildmann et al. 1992; Chen et al. 2004), and are resistant to proteolysis in the insect gut (Chen et al. 2007). Transgenic tomato plants that over-express arginase are more resistant to M. sexta larvae (Chen et al. 2005) whereas suppression of a threonine deaminase gene in Nicotiana attenuate impairs plant defense against M. sexta larvae (Kang et al. 2006). Polyphenol oxidase. Polyphenol oxidases (PPOs) are antinutritive enzymes that catalyze the oxidation of phenolics to quinones, which decrease the nutritive value of the wounded plant by cross-linking with the nucleophilic side chains of proteins and free amino acids. Evidence for PPO involvement in plant defense against insect herbivores includes transcriptional induction of PPO genes by insect attack (Constabel et al. 1995; Thipyapong and Steffens 1997; Haruta et al. 2001), insect growth suppression via artificial diet (Felton et al. 1992), and high stability of ingested, active PPOs in insect gut (Chen et al. 2005). In addition,

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transgenic plants that over-express PPO enhance plant resistance to bacterial pathogens (Li and Steffens 2002), whereas suppression of PPO gene expression increases plant susceptibility to the same pathogens (Thipyapong et al. 2004). Other proteins. There are many other proteins that are potentially involved in plant defenses. A few examples with preliminary supporting evidence include lipoxygenases, peroxidases, and many lypolitic enzymes (El Moussaoui et al. 2001). Lipoxygenases are involved in the biosynthesis of plant hormones such as jasmonic acids and phytoprostanes (oxylipin), which are transcription regulators of defense-related genes (Kessler et al. 2004). Lipoxygenases are also toxic directly to insects when incorporated into artificial diets (Shukle and Murdock 1983; Felton et al. 1994). Even though it is not known on how they produce toxic metabolites, lipoxygenases can generate lipid peroxidation products such as hydroperoxides and aldehydes, which are potent electrophilic reactants. These reactants can then react with the nucleophilic side chains of amino acids, eliminating nutritive amino acids (Felton et al. 1994). The roles of peroxidases in plant defense are multifaceted (Kawano 2003). Peroxidases generate reaction oxygen species, which regulate defense-related signal transduction pathways, initiate hypersensitive reaction, and strengthen cell wall through enhanced lignification and cross-linking (Ralph et al. 2004). The potential direct detrimental effect of peroxidases on insect herbivores is also from their role as “digestibility reducers” (Felton and Summers 1993; Duffey and Stout 1996). Quinones and other oxidative species produced by peroxidases can react with side chains of either free amino acids or proteins, thus reducing the nutritive value of ingested food in the herbivore’s gut.

3.3 Conclusions Plant defense against insect herbivores is complex and via different mechanisms. As a result, factors involved in direct plant defense to insect herbivores are usually additive, and vary among different plant-insect systems. Multiple factors are usually involved in defense against a single insect species, and factors of major importance to one insect species may only be secondary or not effective at all against another insect species (Blau et al. 1978). On the other hand, plant defense to insects is clearly interconnected with other biological processes. Many genes and pathways that are activated by insect attack are also involved in disease defense responses or in responses to other stress agents (Walling 2000). The differentiation of specific defense strategies in different plant-insect systems and the interconnection of insect defense pathways with other physiological processes make plant defense to insects a very complex phenomenon to be analyzed by traditional approaches. Fortunately the arrival of advanced technologies in genomics, proteomics, metabolomics, lipidomics, and bioinformatics allows us to tackle these complicated biological issues more efficiently. Extensive qualitative and quantitative analyses of temporal and spatial variations and system-wide


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assessment of genes, proteins, metabolites, and other molecular components will accelerate the identification of specific targets that are critical for plant defense in individual plant-insect systems. High throughput validation and transgenic techniques will also bring improved crop plants with enhanced resistance using the identified targets for agriculture. Recent advances in new technologies such as RNA interference-based transgenes (Baum et al. 2007; Mao et al. 2007) may enhance the effectiveness of innate and induced plant defenses and prolong resistant crops in agriculture.

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CHAPTER 4

Host Marking and Host Discrimination in Phytophagous Insects Siwei Liu, Baige Zhao and Edmond Bonjour

Abstract Host marking and host discrimination are an inevitable result of evolution in some phytophagous insects. In order to subsist and develop properly, they use a chemical signal, that is, a host marking pheromone (HMP), for communication to avoid laying eggs at the same site. This phenomenon was noted 140 years ago, but it was studied and described for the first time in the 1970’s. Since then, the purposes and functions of host marking have been determined, as well as the strategies and methods of host marking and discrimination. Currently, HMPs have been described in more than 30 species belonging to at least 20 families from five orders. This article reviews the mechanisms of host marking and host discrimination, and the range and function of HMP. It highlights the current understanding of the subject and the potential for practical application. The article addresses future directions in terms of questions asked and the complexity of the research that is needed to answer them. Keywords host marking, host discrimination, host marking pheromone, function of pheromone, phytophagous insects

Siwei Liu Department of Biological Sciences, the University of Texas at El Paso, El Paso, Texas, 79968, USA; Department of Entomology and Plant Pathology, Oklahoma State University, Stillwater, OK, 74078, USA E-mail: [email protected] Baige Zhao Department of Plant and Soil Sciences, Oklahoma State University, Stillwater, OK, 74078, USA Edmond Bonjour Department of Entomology and Plant Pathology, Oklahoma State University, Stillwater, OK, 74078, USA


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Siwei Liu, Baige Zhao and Edmond Bonjour

4.1 Introduction 4.1.1

Concepts

For insects, “Nature is not a Garden of Eden, but an abode of many competing forms of life. Even sibling insects…” (Schoonhoven 1990). For subsistence and development, insects use the oldest language, chemical signals, for communication. Many phytophagous insects are capable of distinguishing food plants based on chemical cues, and can also recognize when host plants are already occupied. There is an important ovipositional behavior of some phytophagous insects in that females avoid depositing eggs on hosts previously marked with HMPs. The HMPs produce stimuli typically perceived by chemoreception. The stimuli, which permit females to distinguish between marked and unmarked hosts, can be categorized as either cues or signals (Seeley 1998). A signal is a stimulus, such as a host marking pheromone (HMP). It has been shaped by natural selection and conveys information from sender to receiver. A cue is also a stimulus, visual or tactile, but it is not shaped by natural selection and conveys information only incidentally (Nufio and Papaj 2001). Cues can be chemical or physical. Chemical cues act as signals of egg or larval occupancy (Butkewich et al. 1987). What are the chemical signals of host marking and discrimination? They are HMPs. Also called ovposition deterring pheromones, some insects deposit chemicals on or around the eggs they lay to deter other females from laying eggs at the same site (Blaakneer et al. 1994). HMPs are generally produced by egglaying females. This event is called host marking. “Host discrimination is frequently mediated through employment of HMP, that appear to function as display or epideictic message” (Roitberg and Mangel 1988). HMPs may be produced by adults, eggs, or larvae, and discriminated by adults or larvae. They may deter other insect species from landing and ovipositing (Nordlund et al. 1981). HMPs, or ovposition deterring pheromones, are used for both parasitic and phytophagous insects. Use of these terms with parasitic insects occurred not only earlier (Nordlund et al. 1981), but also more often than with phytophagous insects. For example, in preparation for this review, 127 articles were searched, and of these, 87 articles (68.50%) reported on parasitic insects, 38 articles (29.92%) reported on phytophagous insects, and two articles (1.55%) reported on both. In these articles, phytophagous insects included herbivorous and carpophagous insects. This review will emphasize the mechanism, function and application of HMPs in phytophagous insects, although a few parasitic insects and references are used as examples. Host marking pheromone is a broad term, which includes epideictic pheromone, spacing and territoriality pheromone (Renwick and Radke 1981). The function of these pheromones, a chemical substance produced by one individual, is to maintain the proper density of individuals of that species and to also deter other females from laying eggs (Klowden 2002). HMP has many

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synonyms such as oviposition-deterring pheromone, marking pheromone, and oviposition-deterring host marking pheromone (Prokopy et al. 1977; Schoonhoven 1990; Aluija and Boller 1992; Hurter et al. 1987), fruit-marking pheromone (Katsoyannos 1975), egg-laying pheromone, and brood-tending pheromone (Nation 2002). 4.1.2

History

Kirby and Spence (1863) observed that when Pieris brassicae laid eggs on cruciferous plants, this influenced females to lay eggs again. The observation was confirmed by Rothschild and Schoonhoven (1977). Salt (1935) reported a “factor” in parasitic insects, and later this same factor was called the spoor factor (Flanders 1951). After another 20 years, the spoor factor was called dispersal pheromone, marking pheromone, and epideictic pheromone (Nordland 1981). Yoshida (1961) studied the evidence of host marking in two species of bean weevils. Prokopy (1972) and Cirio (1972) first reported fruit fly HMPs in apple maggot (Rhagoletis pomonella) and walnut husk flies (R. completa), respectively. Oshima et al. (1973) isolated an oviposition marker from the Azuki bean weevil, Callosobruchus chinensis. Hurter et al. (1987) purified and determined the chemical constitution of the cherry fruit fly Rhagoletis cerasi HMP – {N[15(βglucopyranosyl)oxy-8-hydroxypalmitol]-taurime}. The study of HMP was recently revolutionized (Aluja and Boller 1992).

4.2 The progress of study 4.2.1

Mechanisms

(1) Why phytophagous insects do host marking Host marking is an important evolutionary strategy. It not only reduces competition, but also reduces the waste of eggs and energy. Another function is to stimulate the search for new food or oviposition sites (Vinson 1981). For example, the sorghum shoot fly, Atherigona soccata Rondani, lays only one egg per sorghum plant (Ponnaiya 1951; Ogwaro 1978). The HMP is emitted from the cement used to glue the egg to the plant (Raina 1981). If one plant has more than one egg, only one larva survives to maturity (Raina 1981). HMPs deter both intraspecific (Prokopy et al. 1976) and interspecific (Giga and Smith 1985; McClure et al. 1998; Schoonhoven 1990) female insects from laying eggs at an already marked site (Prokopy et al. 1976). Of course, they prevent a female from returning to the same site as well (Schoonhoven 1990). HMPs also enhance risk–incur parasitoid (Roitberg and Lalonde 1991). (2) How to mark There are three strategies for host marking in phytophagous insects. 1. Release of pheromones–chemical signal. 2. Visual cues–color, form. Butterflies can use

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visual cues due to their well-developed eyes (Schoonhoven 1990). Rothschild and Schoonhoven (1977) reported that the presence of plastic eggs also deter oviposition of the white cabbage butterfly, Pieris brassicae (L.) 3. Physical cues – chemical substances emitted from ovipositional and feeding-puncture sites (Butkewich et al. 1987), damaged foliage (Rothschild and Schoonhoven 1977), and larval frass (deterrent is not a pheromone) deterred oviposition (Renwik and Radke 1981). HMPs are a very complex area of study that has great potential for use. Many species use their ovipositors to deposit HMPs following oviposition. For instance, the apple maggot fly, Rhagoletis pomenella (Walsh), makes a circle of marking pheromone on the fruit surface by dragging her extended ovipositor (Prokopy 1972; Prokopy et al. 1978; Averill and Prokopy 1988). The pheromone trail is about 53 mm long and 0.5 mm wide (Averill and Prokopy 1980). Adult S. neanthracina use their mouthparts to deposit HMP (Zimmerman 1979; Prokopy et al. 1984; Roitberg and Prokopy 1987). Ovipositional deterrents exist in different organs of insects depending on the species. Usually they are found in the female head, thorax, and abdomen and male thorax (Quiring et al. 1998). They are also produced by larvae in the form of oral (Corbet 1973b), anal (Hilker and Klein 1989; Merlin et al. 1996; Ruzick 1996), and exocrine glandular secretions (Hilker and Weitzel 1991; Schindek and Hilker 1996). Most HMPs are produced by the exocrine, digestive or reproductive system (Table 4.1) (Nufio and Papaj 2001). Table 4.1 Site Location of HMP production Order Coleoptera

Diptera

Sites of HMP production

Citation

Hind gut, Malpighian tubules

White et al. 1980

Prothoracic, abdominal glands

Roth 1943; Loconti and Roth 1953

Larvae

Hilker and Weitzel 1991

Head region, deposited by mouthparts Quiring et al. 1998 Midgut, released through orifices

Prokopy et al. 1982a

Larvae, mandibular glands

Corbet 1973a

Lepidoptera

Accessory gland, produce egg-coating substances

Thiery et al. 1995

Hymenoptera

Associated with the legs

Foltyn and Gerling 1985

HMPs can be released from various sources. ① In most situations, HMPs are deposited by females during or following oviposition (Nufio and Papaj 2001; Quiring et al. 1998). ② Some substances are produced by larvae, however, which can deter oviposition (Hilker and Klein 1989; Merlin et al. 1996). For example, HMPs are released by last instars in the genera Ephestia, Plodia and Heliothis (Prokopy et al. 1984). First and second instars of Diptera may also deter C. capitata oviposition attempts (Prokopy et al. 1978). ③ Eggs can release HMP signals (Butkewich et al. 1987). Raina (1981) reported that washed eggs do not


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deter oviposition of the sorghum shoot fly, Atherigona soccata. ④ Juice exudates from the oviposition puncture on olive hosts deter olive fruit flies, Dacus oleae (Gmelin), from repeated egg laying attempts (Prokopy et al. 1978) In this case the active substance is a kairomone. ⑤ In some cases also males can do host marking, which influences egg laying (Papaj et al. 1996). HMPs are water soluble. Prokopy et al. (1978) suggested that they are produced in large quantities with little energy expenditure. In general, the offspring of phytophagous insects must completely develop on or in the host in which eggs are laid (Quiring et al. 1998), including first and second instars. For instance, HMP of the apple maggot fly, Rhagoletis pomonella, can under different conditions last two weeks (Quated in Prokopy 1981a) to three weeks (Averill and Prokopy 1987). The eggs hatch in three to four days (Prokopy 1981b). Persistence of HMPs may vary over time according to the species and also the nature of the message conveyed. Sometimes the deterrent effect of the HMPs is longer than it seems to be needed. Table 4.2 shows that HMPs can remain effective for a long time under dry laboratory conditions. In the field, HMPs have a shorter efficacy because of complex and varied conditions. Table 4.2 Persistence of HMPs Species

Under the condition

Time*

Citation

1. Caribbean fruit fly Anastrepha suspense (Loew)

Fruit

6d

Prokopy et al. 1977

2. Sorghum shootfly Atherigona soccata

On plant

7d

Raina 1981

3. Apple maggotfly Rhagoletis pomonella (Walsh)

Fruit dry condition in field

3 wk

Averill and Prokopy 1986

4. Cabbage butterfly Pieris brassicae

Plant leaf surfaces under vacuum condition In lab, on a glass surface

14 d 1 wk 7 wk

Schoonhoven et al. 1981 Schoonhoven et al. 1981 Schoonhoven et al. 1981

5. Mediterranean fruit fly Ceratitis capitata

In lab hawthorn fruit

6d

Prokopy et al. 1978

*d = day(s); wk = week(s).

Table 4.2 shows that HMPs can be effective for at least six days; the longest persistence is three weeks under field conditions and seven weeks under laboratory conditions. From species to species, there is much variation (Fig. 4.1). The duration of the pheromone is long enough for egg hatching and in some species persists up to first and second instars. With enough food present, some species that usually release HMPs do not release these chemicals. This phenomenon is affected by various factors (Quiring et al. 1998) such as female age (Averill and Prokopy 1987) or egg load (van Randen and Roitberg 1996). The key factor egg-laying female insects is to save energy. On the other hand, when food is scarce, releasing HMPs may not be

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Fig. 4.1 HMPs from five insect species have different persistence under differentcon-

ditions. 1. Caribbean fruit fly; 2. Sorghum shoot fly; 3. Apple maggot fly; 4. Butterfly, Ceratitis capitata; 5. Butterfly, Pieris brassicae.

effective (McClure et al. 1998). For example, under normal circumstances, the larch cone flie, Strobilomyia species, tends to lay one egg per cone (McClure et al. 1996). When seed cones are limited, however, the number of eggs per cone is increased (Roques 1988). (3) How to discriminate If unmarked hosts are readily available, females encountering HMPs usually reject marked host. On the other hand, if unmarked hosts are not readily available, the efficacy of HMPs will rapidly decrease with time and cause females to oviposit on marked host (Quiring et al. 1998). HMPs are non-volatile in phytophagous insects, and they are detected by contact chemoceptors. HMPs can be perceived by receptors on the antennae, tarsi, mouth, ovipositor, or abdomen (Hilker and Klein 1989; Ferguson et al. 1999; Quirring et al. 1998; Crnjaret et al. 1978; Klijnstra and Roessingh 1986; Stadler et al. 1994; Kouloussis and Katsoyannos 1991). Table 4.3 shows different forms of ovopositional deterrents (Quated in Butkewick 1987; Renwick and Radke 1985). For instance, after the apple maggot fly, Rhagoletis pomonella, lays a single egg beneath the fruit skin, the female immediately trails her extended ovipositor on the fruit surface for 30 to 50 seconds while depositing HMP. She then cleans her ovipositor and departs (Prokpy 1972; Roitberg and Prokopy 1987). When a new female arrives on the fruit, receptors on her tarsus (Roitberg and Prokopy 1987) and mouthparts allow her to respond to the HMP (Bowdan 1984; Prokopy 1981a). The female acquires information on the presence or absence of HMP for several seconds to more then a minute (Roitberg and Prokopy 1987). HMPs in tephritid flies are synthesized in the midgut (Prokopy et al. 1982a) and detected with receptors on the foretarsi (Crnjar and Prokopy 1982).


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Table 4.3 The forms of ovopositional deterring Species

Form of ovoposition deterring

Citation

1. Apple maggot fly Rhagoletis pomonella (walsh)

Deters conspecifics from laying eggs –pheromone

Prokopy 1972

2. Codling moths Cydia pomonella (L.)

Other mechanisms

Roitberg and Prokopy 1982

3. European apple sawfly Hoplocampa testudinea (Klug)

Responds to wound exudates of host tissue in discrimination

Roitberg and Prokopy 1984

4. Small cabbage white Pieris rapae

Cabbage leaves release Renwick and Radke1983 volatiles, which deter oviposition

4.2.2

Current situation

(1) Milestones Pheromones have been identified in 1600 species belonging to 90 families from nine orders (Roelofs 1995). In1980’s, the HMPs in 30 species of phytophagous insects had been reported (Roitberg and Prokopy 1987; Kouloussis and Katsoyannos 1991; Rika 1994). In 1990’s, HMPs were reported for over 30 species in four orders: Lepidoptera, Diptera, Coleoptera, and Hymenoptera (Schoonhoven 1990; Quiring et al. 1998). By 2001, HMPs had been reported in Coleoptera, Diptera, Hymenoptera, Lepidoptera, and Neuroptera, among at least 20 families (Prokopy 1981a, 1981b; Roitberg and Prokopy 1987; Landolt and Avorill 1999; Nufio and Papaj 2001). During the 1970s, HMPs were discovered in many species. The frugivorous Tephritidae contains the greatest number of phytophagous insect species that releases HMPs. HMPs have been discovered in 12 species including the apple maggot fly, Rhagoletis pomonella, (Prokophy 1972, 1981a). During the 1980s, many studies were relatively rich in the description of the underlying behavioral mechanisms of communication. From the 1970s and 1980s, isolation and identification of some HMPs emerged. A cherry fruit fly HMP was suspected in 1987. HMPs were further isolated, identified and synthesized in the 1990s. The number of published articles on HMPs decreased during the1990s likely as a result of the complexity of researching the field (Fig. 4.2).

Fig. 4.2 Articales of HMPs found in each decade.

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(2) Application How HMPs can be employed to IPM is a very interesting subject. Katsoyannos and Boller (1976) and Prokopy et al. (1978) suggested using of HMPs as control agents. The most effective strategy may be to use attractants to lure females. HMPs “offer an exciting alternative to agriculture management practices, even though many avenues of research must still be explored” (Roitberg and Prokopy 1987). Katsoyannos and Boller (1976) tested field applications of the ovipositiondeterring pheromone of the European cherry fruit fly, Rhagoletis cerasi, and demonstrated the potential usefulness of this pest management technique. Although the HMPs of over 30 species of herbivorous insects have been demonstrated, only a few have been applied for population control in agriculture. Examples are the European cherry fruit fly (Rhagoletis cerasi) and the large white cabbage butterfly (Pieris brassicae) (Roitberg and Prokopy 1987). Two studies were successful in demonstrating the practical application of HMPs. Prokopy et al. (1982b) used an artificial agar apple in which the female fruit fly, Anastrpha fraterculus, deposited eggs. This study confirmed the hypothesis that the fruit fly produces and deposits a HMP which deters other individuals of the same fly species from laying eggs on this host. Their mimic HMP product has been commercially used. Second is Boller, Hurter and colleagues who also showed success in their study with the cherry fruit fly, Rhagoletis cerasi. In 1976, 1980, they isolated, characterized, synthesized, and employed HMPs in field applications (Hurter et al. 1987; Aluja and Boller 1992; Boller and Aluja 1992). Once an HMP is found, there are many steps before reaching large-scale applications (Fig. 4.3). Identification and synthesis are difficult. After confirming

Fig. 4.3

Flow chart of HMP from research to application.


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tests in the laboratory, small field tests are conducted before using HMPs in largescale applications.

4.3 Discussion Many studies have been conducted on the mechanism of HMPs, but only a few HMPs have been isolated, identified and synthesized. HMPs are reported and described for over 30 species where the system of production, release and reception of HMPs is understood. Thus, the mechanisms of host marking and host discrimination is quite clear. However, identification and synthesis of HMPs are very difficult. Only recently have a few practical applications of HMPs begun to emerge. Problems are complex and involve ovipositional behavior, the specific role of pheromones, and numerous unknown factors. These problems have severely slowed down the successful commercial use of HMPs. If several HMPs could be isolated, identified and synthesized, spraying host crops could become a successful new approach to pest management, especially as methods of trapping, capturing, and deterring females (Prokopy 1972, 1975; Boller 1981). The work that needs to be done to reach this goal requires that we are cognizant of the context and properties under which insects release, receive, process, and employ HMPs. It is a big challenge for biologists to identify and synthesize HMPs, and that way develop new methods by which to combat insect pests. Nevertheless, HMPs offer great hope and will likely become an important part of future insect control strategies. Acknowledgements We sincerely thank Dr. Thomas W. Phillips (Professor and department head, Kansas State University) for his guidance and Dr. Bjorn Martin (Professor, Oklahoma State University) for the review and helpful comments regarding this manuscript.

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CHAPTER 5

Nitrogen Modulation on Plant Direct and Indirect Defenses Yigen Chen and Xinzhi Ni

Abstract Instead of being passively attacked by insect herbivores, plants possess a myriad of defense mechanisms to protect themselves. These mechanisms function broadly either by directly reducing herbivore fitness (direct plant defense), or by indirectly attracting natural enemies of the herbivores (indirect plant defense). Many biotic and abiotic factors including nitrogen affect the expression of plant defenses. Nitrogen fertilization is by and large the most common agronomic practice in crop production. Anthropogenic use of nitrogen contributes to meeting the food supply for the rapidly increasing in human population. On the other hand, it alters plant defense capability and the global nitrogen cycle, which is increasingly threatening plant and animal biodiversity in managed and natural ecosystems. In this chapter, we first briefly review types of plant defense and ecological and environmental consequences associated with anthropogenic addition of nitrogen into ecosystems. We then review the influences of nitrogen fertilization on plant direct and indirect defenses. Finally, we discuss the possibility of developing new sustainable pest management tactics through nitrogen management minimizing the negative and maximizing the positive components of tritrophic interactions through optimized nitrogen applications in agricultural production. Keywords Plant-insect interactions, Tritrophic interactions, Volatile organic compound, Pest management, Anthropogenic fertilization

Yigen Chen Department of Entomology, Michigan State University, East Lansing, MI, 48824, USA E-mail: [email protected] Xinzhi Ni Crop Genetics and Breeding Research Unit, USDA-ARS, Tifton, GA, 31793, USA


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5.1 Introduction Although plants are challenged by many herbivorous insects at different stages of their growth, they are rarely completely destroyed by herbivores. Why this is so or as White (1993) put it, “Why is the world green?” has intrigued researchers of generations. Various hypotheses (e.g., “bottom up” and “top-down” hypotheses) have been proposed to explain this phenomenon (Hairston et al. 1960; Root 1973; Mattson 1980; White 1993; Stiling and Moon 2005). One commonality in these hypotheses is that the abundance of phytophagous insects is limited. However, the possible mechanisms that keep insect herbivores in check diverge. For example, after ruling out all other biotic and abiotic factors that possibly limit herbivore abundance, Hairston et al. (1960) concluded that “predators” limited herbivore population growth. White (1993), on the contrary, suggested that biologically available nitrogen limited the buildup of herbivore abundance. Regardless of the debate of whether “bottom-up effects” (effects through food quality of herbivores) or “top-down effects” (effects through natural enemies of herbivores), or a combination of the two, are limiting herbivore abundance, plants are increasingly documented to possess a range of phenological, morphological, and biochemical defenses against herbivores that could affect the quality of the food source in time and space (Berenbaum 1995; Schoonhoven et al. 2005). Nitrogen fertilization is by and large the most common agronomic practice in crop production. Anthropogenic use of nitrogen helps meet the food demand for the increasing human population, while it alters the global nitrogen cycle because nitrogen fertilizers often leach out readily in nature. The enriched nitrogen contents in streams, ground, underground water, and the atmosphere have serious consequences to global climate and are increasingly threatening plant and animal biodiversity (Rouse et al. 1999; Giles 2005; Camargo and Alonso 2006; Clark and Tilman 2008; Seitzinger 2008). In addition to alteration of nitrogen cycling, anthropogenic fertilization may change plant defense capacities, which can have profound effects on tritrophic interactions and pest management practices. Nitrogen can have profound effects on tritrophic interactions that are basic components of nearly all ecosystems (Fig. 5.1; for a review, see Chen 2007). Nitrogen availability may first change plant nutritional qualities and levels of plant defensive compounds. These changes affect the nutritional value of the plants as herbivore food sources, which may result in low quality prey/host for their natural enemies, in particular, for parasitoids that generally have a higher requirement for quality hosts than predators (Godfray 1994; Williams 1999). Many phytophagous insects are documented to sequester plant defensive chemicals for their own use (e.g., as a nitrogen source and defense against natural enemies; Zagrobelny et al. 2004; Gleadow and Woodrow 2002; Nishida 2002). Changed patterns of plant secondary compounds through changes in nitrogen levels may subsequently influence the patterns of sequestered plant chemicals in herbivores, which in turn may alter the fate of both herbivores and natural enemies (Campbell and Duffey 1979; van Emden 1995; Kester and

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Barbosa 1991; for a review, see Turlings and Benrey 1998). Nitrogen can also affect the plant’s indirect defense via quantitatively and/or qualitatively changing herbivore-induced plant volatile emission that is crucial for natural enemies’ foraging (discussed below). Plant architecture that might affect natural enemy foraging efficiency, and the quality of food and shelter for natural enemies might also be affected by nitrogen fertilization (discussed below).

Fig. 5.1 Scheme of potential nitrogen (N) effects on tritrophic system.

Sustainable pest management needs consideration of both economic and environmental sustainability. Sustainable pest management is becoming increasingly urgent because of rising problems brought about by traditional therapeutic approaches of pest management (Lewis et al. 1997) and increased concerns over environmental contamination (Hamrick 2002). Minimizing nitrogen fertilization not only reduces production costs, but keeps the “nitrogen pollution” at a lower level. Additionally, as will be discussed below, reduced nitrogen fertilization in many cases have positive effects on plant defenses and consequently on the basic components (i.e., tritrophic interactions) of ecosystems. Therefore, nitrogen fertilization manipulation may contribute to sustainable pest control. In this review, we intend to: 1) review the types of plant defenses and their ecological and environmental consequences in relation to anthropogenic addition of nitrogen into ecosystems; 2) review the influences of nitrogen fertilization on plant direct and indirect defenses; and 3) discuss the possibility of developing new sustainable pest management tactics through nitrogen management by minimizing the negative, and maximizing the positive effects of nitrogen application on tritrophic interactions through optimized nitrogen applications in agricultural production.


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5.2 Nitrogen and plant defenses 5.2.1

Categories of plant defenses.

Plant defenses can be phenological, morphological, and biochemical. Plant phenology affects its quality and quantity as food of herbivores (Yukawa 2000; Pfender 2004). For example, younger leaves are generally more nutritious than old leaves. Leaf pubescence (e.g. trichomes) is one of the many of the morphological properties that protect plants from herbivore damage. Plant chemical defense in many cases is the dominant defensive system against herbivory, and the widespread occurance and diversity of plant defensive chemical compounds has triggered many debates on the roles that these compounds play. Although these plant secondary metabolites were originally thought to be “waste products”, more and more ecological and physiological functions of these allelochemicals have been demonstrated (Seigler and Price 1976; Bennett and Wallsgrove 1994; Berenbaum 1995; Pickett 1999; Wink 2003; Zagrobelny et al. 2004; Chen et al. 2006; Chen et al. 2008b). A good example of the anti-herbivore function of plant secondary metabolites is the performance of beet armyworm larvae on wild-type and genetically-modified tomato plants deficient in jasmonate (Thaler et al. 2002). Jasmonate is one of the putative plant hormones that transfer wounding signals from wounded sites to other parts of the plant (Karban and Baldwin 1997; Zhang and Baldwin 1997; Schmelz et al. 2003). Significantly more beet armyworm larvae survived on jasmonate-deficient tomato plants compared to wild tomato, and the biomass of larvae reared on jasmonatefree plants was more than doubled compared to those reared on wild plants. Given choices between host plants with and without defensive compounds, many insect herbivores avoid those with defensive compounds and consume less on these plants (Glynn et al. 2003; Chen et al. 2008a). Plant defenses can also be broadly categorized into direct and indirect plant resistance based on whether plant defense is directly protecting plants from herbivores or indirectly through attracting natural enemies of the herbivores. Direct resistance includes any plant morphological (e.g. glandular trichomes) and chemical trait (e.g. terpenes) that directly and negatively affects herbivore fitness. Indirect resistance, on the other hand, includes any plant trait that increases plant fitness through interactions of herbivores and their natural enemies, for example, through the recruitment of parasitoids or predators of herbivores. Plant direct and indirect resistance can be further divided into constitutive and induced resistance. Besides maintaining diverse constitutive morphological structures and plant secondary metabolites independent of herbivory, plants can be induced to manufacture a large array of defensive compounds and structures in response to herbivory. 5.2.2

Biotic and abiotic factors affecting plant defenses

Plant defenses vary temporally and spatially. For example, phenolic acids of

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common walnut, Juglans regia L. (Juglandaceae), reached a peak in the spring and declined thereafter until summer (Solar et al. 2006). Within a plant, young plant parts generally have higher levels of plant secondary metabolites than mature plant parts. In cotton (Gossypium hirsutum L.) (Malvaceae), young expanding foliage has significantly greater terpenoid aldehyde levels than mature foliage (Chen et al. 2008b). The higher investment in defensive compounds in young than mature or senescent plant parts was demonstrated in some systems to be correlated to the higher fitness values of the young leaves than mature leaves (McKey 1979; Krischik and Denno 1983; Ohnmeiss and Baldwin 2000). In addition to temporal and spatial variations of plant secondary metabolites, many other biotic and abiotic factors (e.g. herbivore feeding, fertilizers, drought, sunlight, and CO2) affect the expression of plant defensive compounds (Coviella et al. 2002; Herms 2002; Chen et al. 2008b). Nitrogen fertilization is generally the most common agronomic practice in crop production. Therefore, in this review, we focus on nitrogen modulation on plant defenses. 5.2.3

Nitrogen nutrition versus “nitrogen pollution”

Nitrogen is one of the most important elements for all organisms, and is considered by many researchers to be a limiting factor for plants and animals (Mattson 1980; White 1993), although it is the most abundant element in the atmosphere. The apparent disparity in nitrogen contents across trophic levels in terrestrial as well as aquatic ecosystems suggests that organisms differ in their need for nitrogen. The dry biomass of nitrogen (%) of herbivores generally is much higher than that of plants (Elser et al. 2000). Fagan et al. (2002) found that insect predators possessed 15% more nitrogen overall than herbivorous insects based on reports of 152 species of insects. Variation in nitrogen content is also observed within herbivores of different phylogenetic positions. More recently derived lepidopteran and dipteran phytophagous insects have lower nitrogen levels than the more primitive coleopteran and hemipteran insects, especially when terrestrial insects are considered (Fagan et al. 2002). One explanation for the tendency of decreasing nitrogen levels in recently derived insects would be to lower their dependency on dietary nitrogen. The widespread occurrence of omnivory, defined as insects that feed on more than one trophic level (Polis and Strong 1996), might also reflect that nitrogen is a limiting factor for herbivores and those of higher trophic levels. Omnivory is found in diverse habitats and across a large number of taxa (Coll and Guershon 2002; Denno and Fagan 2003), and it is considered to be a strategy to cope with low-nitrogen food. Through feeding on non-plant sources of nutrients such as cannibalism (e.g. herbivores) and on other third-trophic level insects (e.g. predators and parasitoids), omnivorous insects may gain supplemental nutrients. Nitrate-nitrogen in most inorganic fertilizers leaches into ground and underground water, and ammonia-nitrogen can be lost from the soil by volatilization. The enriched nitrogen content in streams, underground water,


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and the atmosphere have serious consequences to global climate and are increasingly threatening plant and animal biodiversity (Rouse et al. 1999; Giles 2005; Camargo and Alonso 2006; Clark and Tilman 2008; Seitzinger 2008). In addition to alteration of nitrogen cycling, anthropogenic fertilization may change plant defense capacities, which can have profound effects on tritrophic interactions and pest management practices. 5.2.4

Nitrogen effects on plant defenses

Nitrogen can have profound effects on tritrophic interactions that are basic components of nearly all ecosystems. In this review, we focus on nitrogen effects on plant direct and indirect defenses. Further details in nitrogen effects on tritrophic interactions could be found in Mattson (1980) and Chen (2007). 5.2.4.1

Nitrogen effects on plant direct defense

Soil nitrogen fertilization is known to change the expression of plant chemical constitutive defenses (Herms 2002; Hol et al. 2003; Orians et al. 2003; Chen et al. 2008b). In cotton (G. hirsutum cv. FiberMax 989), nitrogen addition dramatically decreased constitutive carbon-based hemigossypolone, heliocide 2, and heliocide 3 contents of undamaged mature leaves (Chen et al. 2008b). The constitutive levels of jasmonic acids (the established key regulators of plant responses to insect herbivores; Browse and Howe 2008) in young and undamaged cotton leaves (systemic resistance—plant resistance induced in plant parts or organs distal to damaged plant parts or organs, in contrast to localized plant resistance that occurs in damaged plant parts) also decreased also with increased nitrogen fertilization (Chen et al. 2008b). Total concentration of the carbon-based constitutive iridoid glycoside in Plantago lanceolata (Plantaginaceae) (Darrow and Bowers 1999), phenolics in tomato plants, Lycopersicon esculentum (Solanaceae) (Stout et al. 1998), condensed tannins of quaking aspen Populus tremuloides (Salicaceae) (Hemming and Lindroth 1999), and nitrogen-based pyrrolizidine alkaloids in stinking willie Senecio jacobaea L. (Asteraceae) (Hol et al. 2003) were all reduced by nitrogen addition. However, constitutive phenolics of tulip poplar Liriodendron tulipifera and dogwood Cornus florida (Dudt and Shure 1994) and nitrogen-containing protease inhibitor levels in tomato plants L. esculentum (Stout et al. 1998) were not affected by nitrogen treatments. Furthermore, production of trypsin inhibitor in Brassica napus L. (Brassicaceae) seedlings (Cipollini and Bergelson 2001) and in tobacco Nicotiana attenuate (Solanaceae) (Lou and Baldwin 2004), and nicotine contents in tobacco (Lou and Baldwin 2004) were enhanced by nitrogen fertilization. Aside from maintaining diverse constitutive plant secondary metabolites independent of herbivory, plants can be induced to manufacture a large array of defensive compounds and structures in response to herbivory (Traw and Dawson

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2002; Chen et al. 2009). These induced plant defenses can also be affected by nitrogen gradients. For example, the levels of hemigossypolone, heliocide 2, and heliocide 3 in the undamaged young leaves on plants of which a lower mature leaf had been fed upon by 20 3-d-old S. exigua larvae were greater in the low nitrogen treatment (42 ppm nitrogen) than in high nitrogen treatments (112 and 196 ppm nitrogen) (Chen et al. 2008b). However, nitrogen fertilization increased the degree of induced insect resistance in poplar (Populus nigra) after continuous feeding of gypsy moth (Lymantria dispar) for 72 hours (Glynn et al. 2003) and the magnitude of induced trypsin inhibitor in Brassica napus following mechanical injury (Cipollini and Bergelson 2001). A number of theories have been proposed to explain the patterns of plant direct chemical defense in response to environmental factors, including the GrowthDifferentiation Balance Hypothesis (GDB; Loomis 1932; Herms and Mattson 1992), Optimal Defense Hypothesis (OD; McKey 1974, 1979), Plant Apparency Hypothesis (PA; Feeny 1976), Carbon-Nutrient Balance Hypothesis (CNB; Bryant et al. 1983), and Growth Rate Hypothesis (GR; Coley et al. 1985). Among these hypotheses, CNB, GDB, and GR incorporate resource (nitrogen) effects on production of plant defenses. Although so far none of these hypotheses can explain the effects of nitrogen on the production of plant defensive compounds, or can account for the complex of ecological factors that may influence these processes (for a detailed review of the strengths and drawbacks of the hypotheses, see Stamp 2003), nitrogen addition is generally observed to decrease plant defenses in herbaceous crop plants and woody plants (Koricheva et al. 1998; Herms 2002; Throop and Lerdau 2004). 5.2.4.2

Nitrogen effects on plant indirect defense

The recruitment of entomophagous natural enemies by plants is referred to as plant indirect defense. Plant traits that are involved in plant indirect defense are volatile organic compounds (VOCs), floral and extrafloral nectaries, and plant structures that provide shelter for natural enemies. Because the relationship can be so mutualistic, these natural antagonists of herbivores are sometimes called ‘plant bodyguards’ (Dicke and Sabelis 1988; Whitman 1994; Cortesero et al. 2000). Herbivore-induced VOCs that natural enemies rely on when foraging, as well as food and shelter of natural enemies, may be altered by plant nitrogen status. 5.2.4.2.1

Nitrogen effects on plant VOC release pattern

Plants release a blend of volatile chemicals following feeding by herbivores and/ or mechanical wounding. These VOCs are employed by many entomophogous natural enemies as foraging cues. Some of them are released around the actual feeding site, while others can be induced from plant tissue distal to and above the wounded site. Green leaf volatiles (GLVs) (e.g. (Z)-3-Hexenal, (Z)-3-Hexenol, and (Z)-3-Hexenyl acetate), some acyclic monoterpenes, sesquiterpenes,


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homoterpenes, and indole are among the typical locally-induced VOCs (Loughrin et al. 1994; Paré and Tumlinson 1997, 1998). (Z)-3-Hexenyl acetate, some acyclic monoterpenoids, sesquiterpenes and homeoterpenes can be systemically induced (Loughrin et al. 1994; Röse et al. 1996; Paré and Tumlinson 1997, 1998). Many of these herbivore-induced plant-originated VOCs provide essential cues for foraging natural enemies to locate potential host/prey. Both parasitoids and predators have been observed to respond actively to VOCs. For example, the parasitoids Cotesia marginiventris (Cresson) (Röse et al. 1998), Microplitis croceipes (Cresson) (Röse et al. 1998) and Cardiochiles nigriceps Viereck (De Moraes et al. 1998) fly more frequently to host-damaged plants compared to undamaged control plants. Similarly, the predatory mite Phytoseiulus persimilis (Acari: Phytoseiidae) and two insect predators, Scolothrips takahashii (Thysanoptera: Thripidae) and Oligota kashmirica benefica (Coleoptera: Staphylinidae), were more attracted to spider mite (Tetranychus urticae)-infested lima bean plants than control plants without herbivore feeding (Dicke et al. 1990; Shimoda et al. 2002; Choh et al. 2004) Soil nitrogen levels can alter the production and release of these plant volatiles. Although effects of nitrogen on VOC production have been negative or lacking in a few crops (Lou and Baldwin 2004; Olson et al. 2009), nitrogen addition in most cases decreased VOCs following herbivory or mechanical damage or altered (van Wassenhove et al. 1990; Schmelz et al. 2003; Chen et al. 2008b; Olson et al. 2009). For example, in tobacco (Nicotiana attenuata), oral secretion from tobacco hornworm Manduca sexta (L.) and methyl jasmonate (MeJA) induced volatile release was not affected by nitrogen, but low nitrogen availability attenuated the jasmonate and salicylate levels and reduced two nitrogen-containing antiherbivore non-volatile defense compounds, nicotine and trypsin proteinase inhibitors (Lou and Baldwin 2004). Gouinguené and Turlings (2002) found that unfertilized corn plants (Zea mays hybrid Delprim) emitted less volatiles when compared to those that had received a complete nutrient solution. Nevertheless, the role of nitrogen was not implied in this study as all the nutrients were varied. The release of seven VOCs in cotton, (Z)-3-Hexenal, (E)-2-Hexenal, (E)-βFarnesene, (E)-4,8-Dimethyl-1,3,7-nonatriene, α-Bergamotene, γ-Bisabolene, and β-Bisabolol was significantly increased in locally damaged leaves (nonsystemic) when plants were grown under a low nitrogen regime (42 ppm nitrogen) (Chen et al. 2008b). Under a medium nitrogen treatment (112 ppm nitrogen), however, the number of VOCs that was significantly induced by herbivory was reduced to (Z)-3-Hexenal and (E)-2-Hexenal. Furthermore, when plants were grown in a high nitrogen treatment (196 ppm nitrogen), herbivory reduced the release of six abundant VOCs, β-Caryophyllene, (E)-β-Farnesene, αBergamotene, α-Humulene, γ-Bisabolene, and β-Bisabolol. A similar pattern of VOCs in systemically induced plants from leaves above and distal to the mature leaf damaged by Spodoptera exigua (Hübner) was observed (Chen et al. 2008b). In corn (bybrid Delprim), the peak volatile release was detected when nitrogen concentration in the fertilizing solution was the lowest, both after mechanical

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wounding and after addition of volicitin (an elicitor isolated from oral secretion of beet army worm (Schmelz et al. 2003). Low nitrogen availability also increased the production of the main sesquiterpenes, (E)-α-Berganotene, β-Caryophyllene and (E)-β-Farnesene, to a greater extent after volicitin application, compared with mechanical damage. In addition, reduced nitrogen levels made the concentration of jasmonic acid decline at a slower rate, compared to the higher nitrogen levels. 5.2.4.2.2

Nitrogen effects on plant traits as food and shelter of natural enemies

Many insect predators and parasitoids feed on pollen, and floral and extrafloral nectar as supplemental food sources (Keeler 1977, 1978; Hespenheide 1985; Koptur 1985, 1992; Kelly 1986; Idris and Grafius 1995; Jervis and Kidd 1999; Stapel et al. 1997; Rahat et a. 2005; Lee et al. 2004, 2006). Fitness correlates of many natural enemies such as longevity, movement, and fecundity are increased by feeding on these plant foods (Hagley and Barber 1992; Olson and Nechols 1995; Morales-Ramos et al. 1996; Baggen and Gurr 1998; Eijs et al. 1998; Jervis and Kidd 1999; Irvin and Hoddle 2007). For example, male and female parasitoids Diachasmimorpha longicaudata (Ashmead) (Hymenoptera: Braconidae), of the tephritid fruit flies (Diptera: Tephritidae), lived up to 15 and 28 days, respectively, when cotton extrafloral nectaries are available (Sivinski et al. 2006). Conversely, with provision of only water male and female parasitoids lived a maximum of 7 days. Trissolcus basalis (Wollaston) (Hymenoptera: Scelionidae), an important egg parasitoid of southern green stink bug [Nezara viridula (L.) (Hemiptera: Pentatomidae)], lives longer when floral nectaries are available (Rahat et al. 2005). Provision of food sources can attract more natural enemies and increase the mortality of herbivorous insects. For instance, parasitism of the gypsy moth, Lymantria dispar L. (Lepidoptera: Lymantriidae), was higher on plants with extrafloral nectaries, although the parasitoid species richness between nectaried and nectariless plants was not different (Pemberton and Lee 1996). More bollworm Helicoverpa zea (Boddie) (Lepidoptera: Noctuidae) eggs were parasitized by Trichogramma pretiosum Riley (Hymenoptera: Tricho-grammatidae) in cotton plants with extrafloral nectaries than those without nectar (Treacy et al. 1987). Plants not only provide natural enemies with food, but also shelter. Many plant structures such as leaf domatia and leaf veins can provide shelter and overwintering sites to various natural enemies (Walter 1996; Hance and Boivin 1993; Whitman 1994; Corbett and Rosenheim 1996; Elkassabany et al. 1996). Despite the importance of plant for provision of pollen, nectar and shelter, there are no studies of the effects of nitrogen on these plant traits.

5.3 Sustainable management of nitrogen and pests synergistically in crop production In contrast to the overall pattern of reduced VOC release in relation to increased


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nitrogen fertilization under the laboratory and greenhouse conditions (Schmelz et al. 2003; Chen et al. 2008b; Olson et al. 2009), parasitism under field conditions or in large-cage studies is generally not affected by nitrogen fertilization (Loader and Damman 1991; Bentz et al., 1996; Chen and Ruberson 2008; Chen et al. 2008b). The disparity of VOC release and parasitism pattern might reflect a conflict for parasitoids between ease of host location and benefits for their progeny. Although increased VOC release in lower nitrogen plants may facilitate host searching, parasitoids need to find high-quality hosts for the development of their progeny. Parasitoids generally have a higher requirement for the quality of hosts than predators (Williams 1999) and many parasitoids are known to be selective in choosing hosts and they are equipped to detect the quality of the hosts (Godfray 1994; Quicke 1997). Alternatively, parasitoids in the field might not exclusively rely on VOCs. Other orientation cues such as visual cues may also play a role because plants with low and high nitrogen not only differ in height, but also in color and architecture. Plant morphological traits also interact with foraging efficiency of natural enemies, and mutualistic, antagonistic, and neutral relationships between plant trichomes and predators and natural enemies have been documented (Treacy et al. 1987; Cortesero et al. 2000). Therefore, parasitoid foraging cues other than VOCs and plant defensive morphological traits might have confounded the effects of VOCs under field conditions. More field investigations specifically designed for examining effects of nitrogen fertilization on VOCs and subsequently on foraging of parasitoids are needed before any generalization can be made. Predators, in contrast, may cause higher mortality of herbivores in low nitrogen conditions (Loader and Damman, 1991). Herbivores feeding on plants with low nitrogen availability are generally low in nutrition and predators need to ingest a higher number of prey to compensate. Also, increased nitrogen fertilization in many cases reduces plant secondary metabolites (Koricheva et al. 1998; Herms 2002; Throop and Lerdau 2004; Chen et al. 2008b), and consequently reduces plant resistance to herbivory. Sustainable pest management requires consideration of both economic and environmental sustainability. Minimizing nitrogen fertilization not only reduces the production cost, but keeps “nitrogen pollution” at a minimum. However, caution must be taken not to jeopardize crop yields when practicing nitrogen manipulation, because low nitrogen input is generally believed to and shown to translate into low crop biomass as well as yield in some cases (Weir et al. 1996; Chen et al. 2008a). Nevertheless, Shear et al. (1946), argued that, along with nutrition intensity nutrition balance played a critical role when other environmental conditions were kept constant. Therefore, crop yield reduction in low nitrogen fields may not be detected compared to yield in high nitrogen fields (Joham 1986). Chen and Ruberson (2008) proposed that crop yield reduction caused by low nitrogen fertilization input might be alleviated or possibly offset by lower pest damage under the following conditions: (1) a pest’s preference for high nitrogen plants; (2) higher pest mortality by natural enemies of pests on

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lower nitrogen plants; and/or (3) higher plant resistance under low nitrogen conditions because of elevated amounts of chemical defensive compounds. That herbivores are capable of distinguishing high from low nutrient plants is well known (Throop and Lerdau 2004; Prudic et al. 2005; Chen et al. 2008a). Higher VOCs and plant resistance of many plants grown in low than high nitrogen fertilization conditions has also been discussed above. Therefore, reducing nitrogen fertilization through increased plant direct and indirect defenses and a balanced nutrient without sacrificing crop yield is plausible in agricultural production. The reduction of nitrogen input will contribute to sustainable pest management. However, the optimal nitrogen level might be crop system-specific as crops, herbivorous insects, and their natural enemies may response differently to nitrogen treatment (discussed above). Acknowledgements The authors are indebted to Drs Therese Poland (U.S. Department of Agriculture, Forest Service, Northern Research Station, East Lansing, MI) and Dawn Olson (U.S. Department of Agriculture, Agricultural Research Service, Tifton, GA) for their critical review of an early draft of the manuscript.

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Section 2: Molecular Biology, Physiology, Behavior and Ecology

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CHAPTER 6

Viruses and Viral Diseases of the Honey Bee, Apis mellifera Yanping Chen

Abstract Viruses pose a serious threat to the health and well-being of honey bees, Apis mellifera, the most economically valuable pollinators of agricultural and horticultural crops worldwide. Recently, honey bee viruses have gotten a lot of international attention due to the significant disease status that viruses cause in honey bees and the recent observation of the tight correlation between Colony Collapse Disorder (CCD) in the U.S. and Israeli Acute Paralysis Virus. Over the past decades, considerable progress has been made in our understanding of virus infections in honey bees. This review chapter summarizes previous findings and recent progress in the understanding of the morphology, genome organization, taxonomy, transmission, and pathogenesis of honey bee viruses. The prospects of future research and challenges associated with the study of honey bee viruses are also discussed in detail. Keywords Honey bee, viral disease, Colony Collapse Disorder

6.1 Introduction The European honey bee, Apis mellifera L.(Hymenoptera: Apidae) plays a vital role in the global economy and food supply by assisting in the pollination of onethird of the world’s food crops and by producing honey, beeswax, pollen, propolis, royal jelly and other hive products. However, honey bees are inevitably subject to infection by a wide variety of pathogens and populations of honey bees have undergone marked declines since the 1940s (Oldroyd 2007; Stokstad 2007), threatening global agricultural production. The declines are especially alarming considering the Colony Collapse Disorder (CCD), a serious malady in which adult worker bees abruptly disappear and die, leading to the collapse of the entire Yanping Chen Bee Research Laboratory, USDA-ARS, Beltsville, MD, 20705, USA E-mail: [email protected]


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bee colony and hence the name. The CCD has resulted in a loss of 50 to 90 percent of bee colonies in 35 states in the U.S. (Van Engelsdorp et al. 2007) and has also wiped out billions of bees around the world in 2006–2007. As a result of the impact it has had, CCD has gained considerable national and international attention. A metagenomic survey of microbes in honey bee CCD conducted by the consortium of scientists showed one virus, Israeli Acute Paralysis Virus (IAPV), to be strongly associated with CCD (Cox-Foster et al. 2007), raising serious concerns about risks of virus infections in honey bees.

6.2 Strucuture and class sification So far, honey bees have been reported to be attacked by at least 19 viruses (Allen and Ball 1996; Ellis and Munn 2005; Maori et al. 2007). Aside from the filamentous virus and the Apis iridescent virus, all honey bee viruses reported so far share similar properties with viruses within the order of Picornavirales: 1) a genome of positive-sense RNA coated with a capsid protein shell, 2) RNA genome linked to a small protein called VPg (viral protein genome) at the 5’ end and polyadenylated at the 3’ end, 3) genome translation into autoproteolytically processed polyprotein(s), 4) capsid proteins organized in a module containing three related jelly-roll domains which form icosahedral, non-enveloped particles with a pseudo-T = 3 symmetry, and 5) a three-domain module containing a helicase, a cysteine protease, and an RNA-dependent RNA polymerase with the gene order , Hel-Pro-Rep (Le Gall et al. 2008) (Fig. 6.1).

Fig. 6.1 The crystal structure of dicistroviruses using the type species of the Discistroviridae family, cricket paralysis virus (CrPV), as a demonstration. Virions of CrPV consist of a capsid. Virus capsid is not enveloped, and is round with icosahedral symmetry. The isometric capsid has a diameter of about 30 nm. A) Rendering surface. Rendering of a CrPV particle was determined at 2.4Å resolution; B) T = pT3 Lattice. The crystal structure of the CrPV capsid has a typical pseudo T = 3 symmetry; C) Subunit organization. Each of the capsid proteins has the same color (VP1 = Blue, VP2 = Green, VP3 = Red) (download form PDB, primary reference Tate et al. 1999).

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To date, the complete genome sequences of seven honey bee viruses including Acute Bee Paralysis Virus (ABPV), Black Queen Cell Virus (BQCV), Chronic Bee Paralysis Virus (CBPV), Deformed Wing Virus (DWV), Israeli Acute Paralysis Virus (IAPV), Kashmir Bee Virus (KBV), and Sacbrood Virus (SBV), have been reported (Ghosh et al. 1999; Govan et al. 2000; Lanzi et al. 2006; Leat et al. 2000; Marori et al. 2007; de Miranda et al. 2004; Olivier et al. 2008). Based largely on their genomic organization, BQCV, KBV, IAPV and ABPV, belong to the family Dicistroviridae (Fauquet et al. 2005). SBV and DWV are assigned to the family Iflaviridae. Like other members of the dicistroviruses, the genomes of ABPV, BQCV, CBPV, IAPV, and KBV are monopartite bicistronic with the nonstructural proteins encoded in the 5’-proximal ORF and the structural proteins encoded in the 3'-proximal ORF. The two ORFs are separated by an untranslated region known as the intergenic region (IGR). The 5’-UTR and the IGR can both initiate translations as internal ribosomal entry site (IRES) (Fig. 6.2A). In contrast, SBV and DWV have typical genome organization of iflaviruses, monopartite monocistronic with the structural proteins encoded in the 5’proximal ORF and the non-structural proteins encoded in the 3'-proximal ORF. The capsid proteins (CPs) are preceded by a leader sequence with undefined function (Fig. 6.2B). Compared to dicistroviruses, the 5’-UTRs of SBVand DWV

Fig. 6.2 Genomes of honey bee viruses. The RNA genome is covalently attached by a genome-linked virion protein (VPg) at the 5’ and a poly-A tail at 3’ ends. A) The genomes of dicistroviridae (ABPV, BQCV, IAPV, and KBV) are monopartite bicistronic with non-structural genes at the 5’ end and structural genes at the 3’ end. The approximate positions of the helicase (Hel), protease (Pro) and replicase (RdRp) domains in the non-structural protein are shown. The structural protein is encoded by ORF 2 and processed to produce the three major structural proteins (VP2, VP3 and VP1). VP4 is presumed to be cleaved from a precursor comprising VP4-VP3 and is a minor structural component of the virion. The 5’ UTR and the untranslated intergenic region (IGR) between the two ORFs can initiate efficient translation as the internal ribosomal entry site (IRES). B) The genomes of lflaviridae (SBV and DWV) are monopartite monocistronic with structural genes at the 5’ end and non-structural genes at the 3’ end. The genome encodes a single polyprotein that is processed to produce the three major structural proteins (VP2, VP3 and VP1) and the non-structural proteins. The VP4 is in analogous to the VP4 present in some dicistroviruses. The polyprotein is preceded by a leader sequence. The approximate positions of the helicase (Hel), protease (Pro) and replicase (RdRp) domains in the non-structural protein are shown.

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are quite small and lack of an IRES-like element. The virions of honey bee viruses are composed of 60 protomers, each comprised of a single molecule of three subunits, VP2, VP3 and VP1. In addition to these three subunits, there is a smaller fourth protein, VP4 which is also present in the virions of some members such as BQCV and ABPV (Govan et al. 2000; Leat et al. 2000). VP4 is not exposed at the surface of the viral particle and is located on the internal surface of the 5-fold axis below VP1. However, the position of VP4 of CPs is different between dicistroviruses and iflaviruses. Currently, CBPV has not yet been assigned to a particular virus genus or family because of the asymmetrical morphology of the viral particles and the copackaging of multipartite positive-strand RNA genomes in a single virion (Olivier et al., 2008). Electron micrographs reveal that virions of honey bee viruses have very similar morphology, roughly spherical with a particle diameter of approximately 25–30 nm in diameter. The virions possess a buoyant density in CsCl ranging from 1.33 to 1.42 g/mL, and a sedimentation coefficient of between 100 and 190S (Ball and Bailey 1991; Bailey 1976). Therefore, it is difficult to distinguish different honey bee viruses morphologically under the electron microscope. As shown in Fig. 6.3, the virus preparation used for electron microscope analysis was determined by RT-PCR analysis to contain five different viruses, BQCV, DWV, IAPV, KBV, and SBV. No significant differences in virion size and morphology could be observed among the virus particles that comprised the four different viruses.

Fig. 6.3 The electron micrograph of honey bee viruses. Viral particles were purified by CsCl density gradient centrifugation and analyzed using electron micrograph (EM) and RT-PCR. Total RNA was extracted from virus preparation and RT-PCR was performed for the presence of seven viruses, ABPV, BQCV, CBPV, DWV, IAPV, KBV, and SBV. Five viruses, BQCV, DWV, IAPV, KBV, and SBV, were detected in the virus preparation. The electron micrograph reveals that there is no significant difference in the virion size and morphology among the five different viruses. The bar represents 100 nm.


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6.3 Infection and transmission The most crucial stage in the dynamics of virus infections is the mode of virus transmission. In order to become successfully established within the host, viruses must have ways to invade hosts and to be transmitted from one host to another to reproduce and to disrupt normal host functions, which lead to pathogenesis. Because of the importance of the transmission processes in the dynamics of virus infections, elucidation of virus transmission in honey bees represents one of the most rapidly developing research areas. Over the past few years work originating in our lab and others has significantly improved and advanced our understanding of the virus transmission in honey bees. Honey bees possess typical traits of eusocial organisms including cooperative brood care, generation overlap, and reproductive division of labor. The densely crowded populations together with the high contact rate between colony members in honey bee colonies increase the risk of disease transmission. To date, transmission of viruses in honey bees has been demonstrated to occur through horizontal or vertical transmission pathways or both (reviewed in Chen and Siede 2007). In horizontal transmission, viruses are transmitted between different individuals of the same generation. In vertical transmission, viruses are passed vertically from mother to offspring via egg. Horizontal transmission of a virus can occur by the following means: food-borne transmission, venereal (sexual) transmission, and/or vector-borne transmission. Vertical transmission can be further divided into transovum transmission in which viruses are transmitted on the surface of the egg and transovarian transmission in which viruses are transmitted within the egg. Horizontal transmission of viruses via food-borne infection can take place by eating pathogen-contaminated food and passing out viruses from the gut with feces. Natural foods in a honey bee colony consist of honey, pollen, and royal jelly. Earlier studies (Chen et al. 2006a, 2006b; Shen et al. 2005a) demonstrated that multiple viruses including ABPV, BQCV, CBPV, DWV, IAPV, KBV and SBV were detected in colony foods including honey, pollen and royal jelly. The results indicated that colony foods could act as a vehicle for food-borne transmission of viruses. Existence of food-borne transmission route in honey bees has been further supported by evidence of viruses found in gut tissue and feces (Chen et al. 2006a), suggesting that the lining of the gut was likely the primary site of the virus infection via virus contaminated food. While trophallactic chain is an important cohesive force in honey bee colonies, trophallactic activities of honey bees including processing nectar, packing pollen, feeding the brood, and attending the queen offer the potential for food-borne transmission of pathogens. It is very likely that contamination of food by viruses can occur during foraging or processing by virus-infected workers and that food-borne infection can take place by eating virus-contaminated food. Under conditions of high population density, high contact rate, and high trophallactic rate, direct food-borne transmission may be a significant route for spreading viruses in bee colonies.

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Venereal transfer of infection is a type of horizontal transmission by which pathogens are transmitted between two sexes during mating. If drones in honey bee colonies are infected with viruses, mating can pose an opportunity for horizontal transmission of viruses from infected drones to queens via semen, which in turn further contributes to the transovarial transmission of viruses from queens to their eggs. The detection of viruses in adult drones (Chen et al. 2004a), semen (Chen et al. 2006a; Yue. et al. 2006; 2007), and in the spermatheca of queens (Chen et al. 2006b) implies the existence of venereal transmission in honey bees. One of the most serious threats of honey bee virus infections involves vectorborne transmission. Vector-borne transmission requires an intermediate host, a vector, which acquires and transmits viruses from infected to an uninfected host. The Varroa destructor is an obligate parasitic mite (Anderson and Trueman 2000) of the honey bee and uses its piercing mouth-parts to penetrate the body wall of the bees to suck out the hemolymph. The Varroa mites have been catastrophic for beekeeping industry because feeding of varroa mites can result in a decline in host vigor, immunity, weight, shorter bee life span, and the eventual destruction of the colonies (DeJong et al. 1982; Korpela et al. 1992; Kovac and Crailsheim 1988; Weinberg and Madel 1985; Yang and Cox-Foxter 2005). In addition to its direct impact on host health, the feeding of mites on bees provides an entry for pathogens and also gives mites the potential to act as vectors of bee viruses. Viral disease outbreaks in the field have often been reported to be involved in the collapse of bee colonies infested with V. destructor (Ball and Allen 1988; Allen and Ball 1996; Kulincevic et al. 1990). The term Bee Parasitic Mite Syndrome has been used to describe a disease complex in which colonies are simultaneously infected with viruses and infested with Varroa mites (Shimanuki et al., 1994). The frequent field observations of the association of Varroa mite infestation with virus infections in honey bees led to several important laboratory experiments. The roles played by Varroa mites in acquiring and transmitting viruses including DWV and KBV has been experimentally demonstrated in several studies (Bowen-Walker et al. 1999; Chen et al., 2004b; Shen et al. 2005b) and further defined the role of Varroa mites in vectoring virus infections. The observation of the positive correlation between the levels of Varroa mite infestation and the levels of virus concentration in infected bees suggests that vector-borne transmission exists in honey bees and that the Varroa mite is not only a vector but also an activator of bee viruses (Ball and Allen, 1988). Recent findings that viruses replicate in the Varroa mite and that viruses are present in mite saliva (Gisder et al. 2009; Ongus et al. 2004; Shen et al. 2005b; Yue et al. 2005) suggests that Varroa mite is a biological vector of bee viruses. The association of virus infections with Varroa mite infestation in honey bee colonies is an alarming trait of infestation and heightens the necessity of timely treatment for this parasite. Vertical transmission, another mechanism of virus transmission in nature, has been shown to occur in honey bees. Detection of multiple viruses in queens of bee colonies suggests that a vertical transmission pathway might exist within the bee colonies and eggs have the opportunity to obtain viruses from their infected


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mother queens (Chen et al. 2005). Detection of DWV and BQCV in eggs and larval stages that are not normally associated with Varroa mite infestation suggests that the viruses might be transmitted vertically from queens to eggs (Chen et al. 2004a; Yue. et al. 2007). An additional indication of a vertical transmission of viruses was obtained through the detection of viruses in the surface-sterilized eggs and ovaries of the queens (Chen et al. 2006a). The detection of viruses in surface-sterilized eggs and ovaries of the queens excludes the possibility of transovum transmission and suggests the existence of a transovarial transmission pathway, in which viruses infect ovarian tissues of the queen and disseminate in developing eggs before oviposition. Quantification of virus titer in ovaries showed that virus concentration in ovaries was relatively low when compared to other tissues examined (Chen et al. 2006b), suggesting that virus infections in ovaries were retained in a non-replicate or latent stage so that viruses would not be propagated to the level that would have a deleterious effect on embryos. The observations that a simultaneous presence or absence of viruses in mother queens and their offspring, including eggs, larvae, and adults further prove the existence of a vertical transmission pathway. When queens were found to be positive for certain viruses, the same viruses were detected in their eggs, larvae and adult worker bees, though neither queens nor their offspring exhibited any overt symptoms of disease. Meanwhile, when queens were negative for specific viruses, these viruses could not be detected in their offspring. These data indicate that transmission of viruses from queens to their progeny is highly likely. Both horizontal and vertical transmission pathways have been proven to be involved in virus transmission in honey bees (Fig. 6.4) and are believed to be important survival strategies for viruses not only for their long-term persistence in the bee population but also for their establishment in nature. Viruses choose the appropriate transmission pathway based on the developmental, physiological, ecological, and epidemiological conditions. When colonies are under noncompetitive and healthy conditions, viruses remain in bee colonies via vertical transmission and exist in a persistent or latent state without causing honey bees to show any overt signs of infections. Alternatively, when honey bees live under stressful conditions such as infestation of varroa mites, co-infection of other pathogens, or decline in food supply, viruses switch to horizontal transmission and start to replicate. High numbers of produced virions then become much more infectious, leading to the death of hosts and possible collapse of the whole bee colony. Such adaptive flexibility of transmission would enable honey bee viruses to choose the appropriate pathway based on the epidemiological conditions and to profit from both horizontal and vertical transmission pathways for their own survival.

6.4 Replication and pathogenesis Pathogenesis is the process by which an infection of virus leads to disease in its target host. To produce a disease, virus must enter a host, make contact with susceptible cells, locally replicate, spread within the host, and disrupt the normal

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Fig. 6.4 Schematic representation of virus transmission routes in honey bees. Virus transmission in honey bees appears to involve both horizontal and vertical transmission pathways. Viruses infect different bee hosts of the same generation by horizontal transmission via the following means: food-borne transmission, fecal-oral transmission, venereal (sexual) transmission, and/or vector-borne transmission. Viruses are also vertically transmitted from infected queens to their offspring. Both transmission pathways are believed to be the important survival strategies for the persistence and establishment of viruses in bee populations. Solid lines represent horizontal transmission and dotted lines represent vertical transmission (from Chen and Siede 2007).

function of the host to cause damage. As a result, the pathogenesis of disease depends on multiple factors including cell susceptibility to virus replication and virus susceptibility to host defense mechanisms as well as environmental factors that affect pathogenesis. To elucidate virus pathogenesis, the investigation of biological features of the viruses and their respective hosts for efficient virus replication and propagation is essential. While transmission pathways of honey bee viruses have been studied in some detail, not very much is known about the pathogenesis of viruses in honey bees. In this section, we focus on the current available information involving pathogenic processes of virus infections in honey bees. The ability of a virus to invade and replicate in particular cells or tissues is a fundamental requirement for a successful infection. The term “tissue tropism” refers to the specificity of a virus to attach, infect, and replicate in particular cells or tissues. Elucidation of the mechanism of tissue tropism that underlie virus binding on the cellular surface of host through interaction between viral capsid proteins and specific receptors as well as spreading to different host cells require a full understanding of the structure features of a viral particle. The atomic structure of a virus particle by X-ray diffraction offers an opportunity to reveal the molecular


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determinants of the virus that are necessary for the recognition of receptors and the specificity of tissue tropism. One essential requirement for the crystallization and determination of a virus atomic structure is that viruses need to be propagated in a cell culture and isolated to a very high degree of purity. Nevertheless, the investigation into honey bee virus replication and pathgenicity has been severely hindered by lack of such an in vitro system for propagation of viruses and characterization of the interactions between host receptors and a particular virus. For many years, our knowledge of tissue tropism of honey bee viruses was mostly limited to ultrastructural observations of virus cytopathology. Earlier studies showed that different bee viruses had some differences in their tissue tropism in the hosts (Bailey et al. 1979, 1983; Bailey and Ball 1991; Ball and Bailey 1991). The primary infection site of most honey bee viruses usually occurs through the cuticle by direct contact between healthy and infected bees or in the alimentary tract when bees ingest virus-contaminated food. For example, KBV, CBPV and ABPV are most likely transmitted contagiously between crowded live bees via the cytoplasm of broken cuticle hairs, while DWV, BQCV, and SBV causes infection in bees when both young adult bees and larvae ingest the virus particles mixed in with their food. Honey bee viruses are able to spread their infections systemically from initial sites to secondary target tissues of the host via the blood circulation or nervous systems. KBV infects and replicates in most tissues of an infected bee, including the fore- and hindgut epithelial tissue, alimentary canal musculature, epidermis, tracheal epithelium, hemocytes, oenocytes, and treacheal end cells. However, no evidence of KBV multiplication has been found in the nerve tissues (Dall 1987). SBV most commonly accumulates in the hypopharyngeal glands of worker bees, but virus particles have also been found in the cytoplasm of fat, muscle, and tracheal-end cells of larvae (Lee and Furgala 1967). While CBPV particles can be found in the alimentary tract, mandibular and hypopharyngeal glands, CBPV has a particular tropism for nervous tissues. This is probably why infection of CBPV is often associated with paralysis behavior in infected bees. However, CBPV does not appear in the cytoplasm of fat or muscle tissues (Giauffret et al. 1966, 1970; Lee and Furgala 1965). ABPV particles have been seen in the cytoplasm of fat body cells, the brain, and hypopharyngeal glands of acutely paralyzed bees (Bailey and Milne 1969; Furgala and Lee 1966). Localization of BQCVand DWV infection in queens and drones by in situ hybridization and RT-PCR methods showed that infections of DWV and BQCV were spread throughout the whole body including the ovaries, fat bodies, spermatheca, and drone seminal vesicles (Chen et al. 2006b; Fievet et al. 2006). While presence of honey bees virus particles in different host tissues were illustrated previously, there is lack of information on titer of virus replication on different tissues of infected bees, which could provide important insight into the complexity of virus replication strategies leading to disease pathogenesis. Recent advances in molecular technology have greatly expanded our ability to detect and elucidate the molecular events associated with virus infections and pathogenesis.

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As with all single-strand positive sense RNA viruses, replication of honey bee viruses proceeds via the production of a negative-stranded intermediate that are used as templates for the production of positive strand progeny RNAs packaged in new virion particles. Therefore, the presence of negative-strand RNA intermediates is a key indication of active virus replication in infected hosts. The detection of negative strand RNA of viruses offers an excellent alternative to a cell culture system for the demonstration of virus replication and pathogenesis in naturally infected hosts. Using TaqMan real time quantitative RT-PCR (qRTPCR) incorporated with purification of biotinylated cDNA with magnetic beads, a study was carried out to detect and quantify DWV replication in virus infected bees. The detection of replicative intermediates, negative strand DWV RNA, in the hemolymph, gut, wings, legs, head, thorax, and abdomen indicated that active replication of DWV could spread all over the body of bees. The observations that different tissues had distinct kinetics of DWV replication and that the most abundant of negative strand DWV was detected in wings suggested that DWV had a tropism to certain host tissues and that wings were likely the predominant tissue site of DWV replication. These findings stimulate further investigation to unveil the regulatory mechanisms that honey bees use to control the replication, which would shed more light on the pathogenesis of bee virus infections.

6.5 Resistance and immunity Honey bee immunity against intruders is constituted not only by individual-level defense regulated by immune-related genes, but also by the colony-level defense mechanism. While the high level of cohesion in eusocial colonies may increase the risk of disease transmission, the highly elaborate social organization of bee colonies poses a special advantage for colony level immunity to defend against the infection of pathogens (Evans and Pettis 2005; Fries and Camazine 2001; Naug and Camazine 2002). The collective host defenses at the colony level include producing antimicrobial substances in their hive products, performing hygienic behaviors, and generating brood comb fever in response to invasion of pathogens (Spivak and Gilliam 1998; Spivak and Reuter 2001; Starks et al. 2000). It has been reported that honey bees improve their resistance to disease infections by producing antimicrobial substances in their hive products including propolis, honey, pollen and royal jelly (Burgett 1997; Greeneway et al. 1990; Kujumgiev et al. 1999; Miorin et al. 2003; Ohashi et al. 1999; Santos et al. 2005). The hygienic behaviors of worker bees such as uncapping and removing the diseased or dead brood have been shown to be effective against SBV and CBPV in the colonies (Bailey et al. 1964; Drum and Rothenbuhler 1983). Analysis of honey bee genome sequences shows that A. mellifera, compared to the genomes of the fruit fly, Drosophila melanogaster and Mosquito, Anopheles gambiae, has fewer paralogs for genes related to innate immunity (The Honeybee Genome Sequencing Consortium 2006). The reduction of immune related genes in honey bees may be a result of strengthened colony-level defense of social insects.


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Like all insects, honey bees utilize three lines of immune defenses to protect themselves from disease agents: physical and chemical barriers, cellular immune response, and humoral immune response at the individual defense level. Although the physical and chemical barriers usually keep pathogens from entering the body, pathogens occasionally penetrate the exoskeleton or alimentary canal to gain access to the internal tissues. Whenever physical and chemical barriers are breached, honey bees use innate defense mechanisms comprising cellular and humoral responses to combat the disease agents. In cellular immune responses, hemocytes confer cellular immunity and hemocytic response is mediated by phygocytosis, nodule formation, and encapsulation of invaders. Humoral responses involve activation of the phenoloxidase cascade and biosynthesis of antimicrobial peptides via a complex set of signaling pathways, Toll, Imd, and Jak-STAT (reviewed in Huszar and Imler 2008). While the humoral and cellular immune responses to bacterial and fungal infections have been characterized and documented in honey bees, relatively little is known concerning how honey bees recognize and fight viral infections. Most of our knowledge about the immune response to virus infection is derived from studies of Drosophila (Huszar and Imler 2008). Recent genome-wide screens in the fruit fly, D. melanogaster suggest that both the Toll (Zambon et al. 2005) and JakSTAT (Dostert et al. 2005) signaling immune pathways are involved with the antiviral immune response. Knowledge of the antiviral immunity demonstrated in Drosophila should provide us with important insight into the relationship between virus infections and host immune responses in honey bees. Apparent orthologs for the key members of Toll and Jak-STAT pathways are present in honey bees (Evans et al. 2006) and these pathways will be key initial targets for determining viral immune functions in bees. More recently, Dicer-2, an endonuclease that is required by RNA interference (RNAi) and cleaves double stranded RNA into the small interfering RNA (siRNA) that represses gene expression, was proven to play an essential role in host defense against non-enveloped, positive-sense RNA viruses including Flock House Virus, Drosophila C virus, and Sindbis Virus in vivo (Galiana-Arnoux et al. 2006; Zambon et al. 2006). A recent study showed that RNAi was an efficient and feasible way of controlling IAPV by feeding with pathogen-specific dsRNA (Maori et al. 2009). Since the genomes of honey bee viruses are positive-stranded RNA molecules which replicate and result in dsRNA replication intermediates that are attractive targets for siRNAs, we would expect that RNAi would be an important defense mechanism against bee viruses.

6.6 Management of virus infections An integrated pest management program for bee diseases caused by viruses should include at least the following three components: 1) good beekeeping management practice; 2) accurate diagnosis of diseases; and 3) selecting and breeding of disease-resistant strains of honey bees. Viral disease outbreaks as well

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as inapparent viral infections can dramatically affect the profitability of the beekeeping industry. Unfortunately, there are currently no specific chemotherapies to tackle viruses because they use host cells for reproduction and survival. Although vaccination is widely used to prevent viral diseases in human and domestic animals, investment in vaccine development may not be a cost effective approach to warrant sufficient economic impact for honey bee viruses due to the high population density of honey bee colonies. While direct treatment is not available to eliminate viruses, effective management of bee viral diseases can be achieved by implementation of good beekeeping practices that would help prevent the spread of viruses and enhance honey bees’ own natural immunity to combat diseases. Good beekeeping practices can be achieved through maintaining a clean environment, providing bees with good nutrition, disinfecting hive tools with virucidal agents, treating parasites and other pathogens, and replacing combs and queens when the problem is serious. A quick and accurate disease identification is critical for making correct disease management decisions. Therefore, early diagnosis of virus infections is an important component of the virus surveillance and control program. It will help to determine the epidemiology of bee viral infections and to monitor honey bee colonies for viruses to prevent the spread of diseases. For many years, the detection and identification of viral infection in honey bees were based largely on serological methods (Allen and Ball 1995; Allen et al. 1986; Anderson 1984). The development of molecular methods has revolutionized the diagnosis of viral diseases and provided powerful tools for specific, sensitive and rapid identification of viruses. The PCR-based method has become a standard method for detection, quantification, and phylogenetic analysis of honey bee viruses. With the increasing genomic information of bee viruses, we would expect that molecular assays such as Northern blotting analysis, real-time quantitative RTPCR, microarray analysis, and other emerging methods will continue to serve as predominant tools for the diagnosis of viral diseases in honey bees. Selection and breeding of disease resistant bee strains are also an effective way to defend against viral attacks in honey bees. Several traits of honey bees, such as hygienic behavior and Suppressed Mite Reproduction (SMR), are important mechanisms of disease resistance (Harbo and Harris 2005; Lapidge et al. 2002; Spivak and Gilliam 1998; Spivak and Reuter 2001).). Highly hygienic bees can efficiently suppress the virus infection and V. destructor infestation by quickly recognizing and removing the diseased brood and varroa mites from combs. Nonhygienic bee lines show a slower removal response to diseased bee brood than bee stocks selected for hygienic traits (Spivak and Gilliam 1998). Such hygienic behaviour strongly depends on gene effects and has been the basis for breeding programs. The development of an integrated program to select bee populations with desirable traits, to preserve honey bee germplasm, and to arrange the mating of queens and drones will provide an important tool to breed for disease-resistant genotypes and hold great promise for colony level disease resistance. In addition, with completion of the honey bee genomic sequence, it becomes possible to


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conduct genome-wide selection for genotypes with defensive and hygienic behaviors and to characterize the genes that confer disease resistance and to genetically manipulate the genes to enhance the disease resistance in honey bees. RNAi has emerged as an effective tool to control viral diseases. An increasing number of studies clearly demonstrate the enormous potential of RNAi in the prevention and control of viral infections. However, more research is needed to evaluate the therapeutic potential of RNAi, and to improve the efficacy of the applications. Acknowledgements I wish to express my most sincere gratitude to Dr. Yan Zhao, Molecular Plant Pathology Laboratory, USDA-ARS, Beltsville, MD for his critical review of this manuscript.

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CHAPTER 7

Biological Function of Insect Yellow Gene Family Jianyong Li and Bruce M. Christensen

Abstract The yellow-y gene in Drosophila has been recognized for many years. Its sequence shares no apparent sequence homology to proteins from noninsect species. Mutation of the yellow gene produces a yellow-colored cuticle; therefore, the yellow gene has been extensively used as a model to study the molecular regulation of protein expression because of its visible phenotype. The completion of the Drosophila genome revealed a number of yellow-y like genes and based on their sequence similarity with yellow-y they have been classified into a Drosophila yellow gene family. As more insect genomes have been sequenced, it has become apparent that a yellow gene family is present in other insect species as well. The yellow gene family is unique in insects because members of this family share no apparent sequence identity to non-insect species. This then leads to some fundamental questions as to why this group of proteins has evolved only in insects and what functions do they perform? Based on limited research with select insect yellow genes, we speculate that at least one of its primary functions involves cuticle and eggshell (chorion) formation and hardening during insect development, and that the yellow gene family is vital for insect survival. The following provides data that describe/discuss the presence of a yellow gene family in different insect species, and the function or possible functions for some individual yellow genes. We also discuss some future directions for research that lead to a more comprehensive understanding of the insect yellow gene family. Because studies dealing with the insect yellow gene family have been limited primarily to Drosophila, the Drosophila yellow gene family is commonly used in the following discussion.

Jianyong Li Department of Biochemistry, Virginia Tech, 111 Engle Hall, Blackburg, VA, 24061, USA E-mail: [email protected] Bruce M. Christensen Department of Pathobiological Sciences, University of Wisconsin-Madison, 1655 Linden Drive, Madison, WI, 53706, USA


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Jianyong Li and Bruce M. Christensen

Keywords insect yellow gene, biological function, Drosophila, insect genomes

7.1 Drosophila yellow gene In Drosophila, a loss of function in one of its genes (due to mutation) leads to the production of a yellow-colored cuticle (Fig. 7.1). Based on the observed phenotype, this particular Drosophila gene has been termed the yellow gene. Logically, the gene should have been named “brown/black gene” because it is responsible for producing a brown-black colored cuticle. This Drosophila yellow gene is related to insect melanization/pigmentation (Gompel et al. 2005; Jeong et al. 2006; Wittkopp et al. 2002a; Wittkopp et al. 2002b; Wittkopp et al. 2003). Surprisingly, however, the mechanism by which the yellow gene promotes cuticle melanization in Drosophila remains unknown. The primary sequence of the Drosophila yellow gene shares no similarity to non-insect proteins. Before the Drosophila genome became available, the yellow gene had been considered somewhat unique because there was no other similar gene or protein available from other species. Due to its easily visible phenotype upon loss of function, this Drosophila yellow gene attracted a great deal of attention since its discovery. For example, there already have been hundreds of references related to the Drosophila yellow gene in the literature prior to the completion of the Drosophila genome project.

Fig. 7.1 Physical appearance of wild type (left) and yellow-y mutant of Drosophila melanogaster. From http://www.exploratorium.edu/exhibits/mutant_flies/.

7.2 The insect yellow gene family Soon after the completion of the Drosophila genome-sequencing project in 2002, several genes, sharing apparent sequence homology with the typical yellow gene, were found in the Drosophila genome and classified as yellow genes. Since then the Drosophila yellow gene family has grown (Drapeau 2001), and now a total of 13 genes have been annotated as members of this family. These genes are grouped with the classical Drosophila yellow gene (termed yellow-y to distinguish it from other yellow genes) based on their sequence similarity and have been named yellow-b, yellow-c, yellow-d, yellow-d2, yellow-e, yellow-e3, yellow-f, yellowf2, yellow-g, yellow-g2, yellow-g3, and yellow-h. Like yellow-y, other members of this Drosophila yellow gene family share essentially no sequence similarity with proteins from bacteria to humans, except for some similarity with the

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proteins from other insects. The names of these Drosophila yellow genes bear are not related their physiological functions. The rapid advance in DNA sequencing technology has been accelerating the genome projects of various model species. Since the completion of the first Drosophila genome, a number of insect genomes, including Acyrthosiphon pisum, Aedes aegypti, Anopheles gambiae, Apis mellifera, Bombyx mori, Tribolium castaneum, etc., have become available. Database searches of their genomes, using the Drosophila yellow-y, enlist invariably a similar yellow gene family from each species. Figure 7.2 illustrates the Drosophila yellow gene family in comparison with a similar yellow gene family from An. gambiae. The yellow gene family contains 10 to 15 members in most of the insect genomes, but the Apis mellifera yellow gene family has 26 individual yellow-like genes. The evolution and potential functions of the yellow/major royal jelly protein family of Apis mellifera (Drapeau et al. 2006a) and the presence of a yellow gene family in Bombyx mori (Xia et al. 2006) have been discussed.

Fig. 7.2 Comparison of Drosophila and Anopheles gambiae yellow gene families.

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Within a given species, members of a yellow gene family share moderate to high sequence identity. For example, The Drosophila yellow genes share 14%– 78% sequence identity and the T. castaneum yellow genes share 15%–60% sequence identity. However, the corresponding orthologous yellow genes from different species share much greater sequence identity (Fig. 7.2). For example, the yellow-y and yellow-c from currently available insect genomes share 51%– 98% sequence identity. Their corresponding yellow-f and yellow-b sequences share 40%–90% identity. Through extensive comparisons of orthologous yellow genes among the available insect genomes, it has become clear that yellow-y and yellow-c are highly conversed, followed by yellow-b (>42%) and yellow-f (>40%). The levels of sequence identity for yellow-e, yellow-g and yellow-h are moderate. Therefore, it becomes questionable if the assigned yellow-e, yellow-d and yellow-g from different species are true orthologous genes across species.

7.3 Physiological functions of insect yellow gene families The physiological functions for most of the insect yellow genes are unknown, which makes it difficult or impossible to practically summarize the overall functions of the insect yellow gene family. However, the physiological importance of the yellow gene family can be somewhat predicted based on the Drosophila genome projects. One of the Drosophila genome projects has been to disrupt each Drosophila gene by the insertion of a single transposable element. In a 2004 report, 5362 individual Drosophila lines, each containing one of the 5362 disrupted genes (39% of the 13,666 currently annotated Drosophila genes), have been created (Bellen et al. 2004). Now it is anticipated that more individual lines with numerous newly disrupted genes will be produced in Drosophila. Database search of the Drosophila yellow family genes identified the presence of the disrupted yellow-y, yellow-e, and yellow-g lines. For the majority of yellow genes, however, no mutants have been created. This suggests that most of the yellow genes are essential, because their knockout mutants cannot be created or are not viable if their function is eliminated. Involvement of insect yellow genes in melanization: As indicated above, the lack of mutants for the majority of Drosophila yellow genes provides some basis to suggest the physiological importance of this gene family in fruit flies, and in turn the importance of the yellow gene family in other insects as well. However, it does little to establish the specific function for the products of individual yellow genes. Among the Drosophila yellow genes, yellow-y has been extensively studied. The yellow-y gene contains two exons and four tissue specific enhancers (i.e., wing, body, bristle and tarsal claw). Taking advantage of its remarkable visible phenotype, the Drosophila yellow gene has been extensively used as a marker gene for study of gene regulation (Drapeau et al. 2006b; Jeong et al. 2008; Labrador et al. 2008; Llopart et al. 2005; Prud'homme et al. 2006; Prud'homme et al. 2007; Simpson 2007; Soshnev et al. 2008) and for the production of mutant. The flybase (http://flybase.org/) indicates that the molecular function of yellow-y


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is unknown, but specified that the gene is involved in numerous biological processes, including cuticle pigmentation, melanin biosynthetic processes, pigmentation during development, male courtship behavior, veined wing extension, mating behavior, adult chitin-based cuticle development, instar larval or pupal development, and courtship behavior (http://flybase.org/reports/ FBgn0004034.html). Among all these suggested functions, perhaps only its involvement in cuticle melanization/pigmentation stands on reasonably solid ground, because a loss of its yellow-y function prevents the formation of a normal brown-black cuticle. This then leads to the fundamental question: what exactly does yellow-y do to promote the production of a brown/black cuticle. The close relationship between cuticle melanization and yellow-y function provides some basis to speculate that yellow-y might be directly involved in insect melanization. In the melanization pathway, tyrosine is the initial precursor. The biosynthesis of melanin from tyrosine is a very complicated biochemical process (Fig. 7.3). In these reactions, hydroxylation of tyrosine to dopa and conversion of dopachrome (DC) to 5,6-dihydroxyindole (DHI) are the two limiting steps in the production of melanin in insects. Hydroxylation of tyrosine to dopa, oxidation of dopa to dopaquinone, and oxidation of DHI to indolequinone are catalyzed by phenoloxidase, which does not seem to be related to yellow-y. Other than these phenoloxidase-mediated reactions, the remaining limiting step of insect melanization is the conversion of DC to DHI (Fig. 7.3). It has been known for many years that insects have an enzyme that catalyzes the decarboxylative structural rearrangement of DC to DHI, leading to the rapid melanin production (Li et al. 1994). However, the identity of this enzyme was unknown. Based on its function, we named it dopachrome conversion enzyme (DCE) in an earlier report (Li et al. 1994). The importance of the yellow-y in promoting insect melanization led to an initial belief that yellow-y could be a DCE that promoted insect melanization by rapidly converting DC to DHI.

Fig. 7.3 Melanization pathway in insects.

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To determine the identity of DCE, we purified his enzyme from Aedes aegypti mosquitoes (DCE activity is much greater in mosquitoes than in Drosophila), which enabled us to obtain a partial amino acid sequence and then cloned its cDNA (Johnson et al. 2001). Blast search of protein database identified Drosophila yellow-y (30% identity) as its best match (Fig. 7.4). After the completion of Drosophila genome project in 2002 and the establishment of its yellow gene family, we determined that the mosquito DCE shares sequence identity to yellow-f (41%), yellow-f2 (40%), yellow-c (34%), and yellow-b (30%), and also recognizable sequence identity (16%–27%) with other members of the Drosophila yellow gene family.

Fig. 7.4 Sequence comparison of Ae. aegypti DCE and Drosophila yellow-y.

The high sequence identity of the mosquito DCE with Drosophila yellow-y provides a reasonable basis for the belief that yellow-y was a DCE. Recombinant mosquito DCE, expressed in insect cell protein expression system, has high DCE activity (Fang et al. 2002), but recombinant Drosophila yellow-y, produced in the same manner, did not (Han et al. 2002). Because Ae. aegypti DCE also shares slightly high sequence identity with Drosophila yellow-f and yellow-f2, yellowf2, yellow-c and yellow-b, we also expressed each of these yellow genes in an insect cell protein expression system and determined that yellow-f and yellow-f2 code for DCE in Drosophila (Han et al. 2002). Drosophila yellow-f and yellowf2 shares 72% sequence identity and their biochemical characteristics are essentially the same, but yellow-f is expressed primarily during larval stages and yellow-f2 is expressed mainly during the pupal stage. These data demonstrate that at least 2 Drosophila yellow genes are directly involved in the insect melanization pathway and responsible, to a great extent, for the rapid melanization reactions in this insect (Fig. 7.5). Involvement of yellow family genes in insect eggshell formation and hardening: In Drosophila egg development, yellow-g and yellow-g2 are extensively expressed in follicle cells. Disruption of yellow-g expression through p-element transposon insertion reveals that mutant females are sterile, but male fertility is not affected. Although oogenesis proceeds in mutant females, eggs laid by mutant females are defective and collapse (Claycomb et al. 2004). In our recent analysis of protein composition in Anopheles gambiae chorions (eggshell), we identified a number of proteins expressed by members of the Anopheles yellow genes family, including EAA08479 (DCE or yellow-f equivalent),


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Fig. 7.5 Drosophila yellow-f and yellow-f2 accelerates the melanization pathway. (A) Dopa solution before addition of phenoloxidase. (B) Accumulation of dopachrome at 30 min after addition of phenoloxidase. (C) Accumulation of melanin at 5 min after addition of both phenoloxidase and yellow-f or yellow-f2 into dopa solution. Presence of DCE rapidly converts dopachrome to 5,5-dihydroxyindole, leading to rapid melanin production.

EAA03946 (yellow-h), EAA11735 (yellow-g), and EAA11686 (yellow-g2) (Fig. 7.6, unpublished data). These preliminary data suggest the involvement of select yellow genes in mosquito chorion formation and hardening. The overall physiological functions of yellow genes in insect cuticle and eggshell formation and hardening, however, remain to be substantiated.

7.4 Outstanding questions and future research directions in insect yellow genes Members of the insect yellow gene family share no apparent sequence homology to proteins of non-insect species and the inability to create mutants for the majority of the yellow genes in Drosophila suggests that this gene family as a whole is vital for survival and development. Therefore, it seems reasonable to propose that the insect yellow gene family is one of the gene families that define these species as insects. Consequently, to achieve a comprehensive understanding of insect biology, physiology, and evolution, it is necessary to understand the physiological functions of the insect yellow gene family. Among the Drosophila yellow genes, its yellow-f and yellow-f2 catalyze the conversion of DC to DHI in the insect melanization pathway; the phenotype of yellow-y mutant provides a basis to suggest its involvement in insect melanization; and the presence of yellow-g, yellow-g2 in both Drosophila and mosquito eggshell indicates their function in eggshell formation. A very recent report indicates that yellow-e in Bymbox mori is involved in pigmentation of head and tail (Ito et al. 2009). These limited data provide a reasonable basis for suggesting that one of the primary functions of the insect yellow gene family concerns cuticle and eggshell formation and hardening. The cuticle (also named exoskeleton) provides insects with protection against physical injury and water loss, rigidity for muscle attachment and mechanical support, and flexibility for

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Fig. 7.6 Two-dimensional profile of An. gambiae chorion proteins. Solubilized chorion protein samples were first separated by two-dimensional gel electrophoresis. Individual protein spots were excised from the gel and in-gel digested with trypsin. Tryptic peptides extracted from individual gel spots were analysis by Liquid chromatography and tandem mass spectrometry (LC/MS/MS). Protein spots labeled with 1, 34, 46 and 58 corresponding to Anopheles gambiae EAA08479 (DCE or yellowf), EAA03946 (yellow-h), EAA11735 (yellow-g) and EAA11686 (yellow-g2), respectively.

joints. The highly protective cuticle is one of the reasons why insects are the most successful animals on earth. During larval development, continued growth requires that insects periodically shed their old cuticle and produce a new one. The newly formed cuticle is soft and elastic, which allows it to stretch and expand to accommodate the increased body size, but at this time it also is vulnerable to adverse environmental conditions and must be hardened or solidified shortly after insects shed their old cuticle. The importance of the yellow gene family in cuticle formation is strongly supported by the fact that Drosophila yellow-f and yellowf2, and mosquito DCE (yellow-f equivalent), accelerate greatly the melanin biosynthesis. And this gene family’s role in chorion formation is documented by the collapse of Drosophila eggshell upon loss of yellow-g function and presence of DCE (yellow-f), yellow-g, yellow-g2 and yellow-h in mosquito eggs. Among the insect yellow genes, Drosophila yellow-y is extensively studied. Yellow-y has been extensively used as a model in studies dealing with the molecular regulation of protein expression (Drapeau et al. 2006b; Jeong et al. 2008; Labrador et al. 2008; Llopart et al. 2005; Prud'homme et al. 2006; Prud'homme et al. 2007; Simpson 2007; Soshnev et al. 2008). This is obviously


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due to the visible phenotype of yellow-y mutants. Compared to other yellow genes, however, yellow-y does not seem to be critical for fly survival because a loss of its function does not result in death or reduced longevity. However, yellow-y remains the most popular yellow gene in experiments involving the yellow gene family; therefore, it seems apparent that we should increase our understanding of this “well-like, extensively utilized gene”. At the very least we should determine the specific function of the product of yellow-y. In Drosophila, the functional elucidation of individual genes has relied heavily on producing their corresponding gene mutants. Although this approach can be powerful for identifying the general function of a given gene, it has limitation. For example, these types of studies clearly show the involvement of yellow-y in insect pigmentation, but provide no information relative to the mechanism(s) that might be controlled by this gene. To truly understand the specific role of yellow-y, it will be necessary to thoroughly characterize its protein. Among the insect yellow genes, the role Drosophila yellow-f and yellow-f2 in mediating the conversion of DC to DHI is well defined (Fig. 7.3). Therefore, these gene products can be called DCE. Recombinant DCE proteins (including yellow-f, yellow-f2 and mosquito DCE) can also catalyze the conversion of dopaminechrome to DHI (an isomerization-tautomerization process), but at a lower catalytic efficiency. Based on their activity to dopaminechrome, it is clear that insect DCE can be classified as an intermolecular oxidoreductase (i.e. the oxidation of one part of a molecule with a corresponding reduction of another part of the same molecule) (Fig. 7.7). When DC is used as the substrate, DCE apparently catalyzes the same electron transfer process, leading to the production of an intermediate (the true enzymatic reaction product) that is highly unstable and that undergoes rapid non-enzymatic decarboxylation to produce DHI as a relatively stable product. The determination of DCE primary sequences and the type of reaction they catalyze is a major achievement in efforts to gain a comprehensive understanding of insect DCE, but the detailed chemical mechanism and the structural basis of its catalytic action is far from clear. For example, the factor or residue(s) responsible for electron transfer, the conformation of its active site, the residues interacting with DC, the residue(s) involved in catalysis, etc., cannot be derived from primary sequences. Details concerning chemical mechanisms of insect DCE also cannot be derived through comparison with existing information, because DCE sequences share no similarity with any functionally characterized proteins from non-insect species. Therefore, we still have much to learn, even for the functionally defined Drosophila yellow-f and yellow-f2 (or DCE from other species). To understand the structural basis of DCE-mediated reactions, it will be necessary to determine the 3-D structure of DCE and the structures of its complexes with substrates or inhibitors. Yellow-f and yellow-f2 share more than 30% sequence identity with yellow-b, yellow-c, and yellow-y. These proteins likely have similar folds in their 3-D structures. Consequently, a 3-D structure of yellow-f or yellow-2 should provide valuable insight for elucidating the specific functions of other yellow gene family members.

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Fig. 7.7

DCE-catalyzed isomerization-tautomerization of dopaminechrome to DHI

A bottleneck in attempts to elucidate the specific function of the individual insect yellow genes has been the difficulty in obtaining reasonable quantities of functional proteins for extensive biochemical and structural characterization. In an earlier study, we purified a DCE from mosquitoes, which led to the subsequent identification of Drosophila yellow-f and yellow-f as DCE. However, it is a painstaking process to isolate individual yellow proteins from any insects. It also is problematic to express their recombinant proteins in cell lines. In spite of these difficulties it is absolutely necessary to analyze these insect yellow gene products at protein level if one wants to clearly understand their physiological functions. Acknowledgements This work is supported by an NIH grand AI 19769.

References Bellen H J, Levis R W, Liao G, et al. The BDGP gene disruption project: single transposon insertions associated with 40% of Drosophila genes. Genetics, 2004, 167: 761–781. Claycomb J M, Benasutti M, Bosco G, et al. Gene amplification as a developmental strategy: isolation of two developmental amplicons in Drosophila. Dev. Cell, 2004, 6: 145–155. Drapeau M D. The family of Yellow-related Drosophila melanogaster proteins. Biochem. Biophys. Res. Comm., 2001, 281: 611–613. Drapeau M D, Albert S, Kucharski R, et al. Evolution of the Yellow/Major Royal Jelly Protein family and the emergence of social behavior in honey bees. Genome Res., 2006a, 16: 1385– 1394. Drapeau M D, Cyran S A, Viering M M, et al. A cis-regulatory sequence within the yellow locus of Drosophila melanogaster required for normal male mating success. Genetics, 2006b, 172: 1009– 1030. Gompel N, Prud'homme B, Wittkopp P J, et al. Chance caught on the wing: cis-regulatory evolution and the origin of pigment patterns in Drosophila. Nature, 2005, 433: 481–487. Fang J, Han Q, Johnson J K, et al. Functional expression and characterization of Aedes aegypti dopachrome conversion enzyme. Biochem. Biophys. Res. Commun., 2002, 290: 287–293. Han Q, Fang J, Ding H, et al. Identification of Drosophila melanogaster yellow-f and yellow-f2 proteins as dopachrome-conversion enzymes. Biochem. J., 2002, 368: 333–340. Ito K, Katsuma S, Yamamoto K, et al. Yellow-E determines the color patterns of the larval head and tail spots of the silkworm, Bombyx mori. J. Biol. Chem., 2010, 285: 5624–5629. Jeong S, Rebeiz M, Andolfatto P, et al. The evolution of gene regulation underlies a morphological difference between two Drosophila sister species. Cell, 2008, 132: 783–293. Jeong S, Rokas A, Carroll S B. Regulation of body pigmentation by the Abdominal-B Hox protein


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and its gain and loss in Drosophila evolution. Cell, 2006, 125: 1387–1399. Johnson J K, Li J, Christensen B M. Cloning and characterization of a dopachrome conversion enzyme from the yellow fever mosquito, Aedes aegypti. Insect Biochem. Mol. Biol., 2001, 31: 1125–1135. Labrador M, Sha K, Li A, Corces V G. Insulator and Ovo proteins determine the frequency and specificity of insertion of the gypsy retrotransposon in Drosophila melanogaster. Genetics, 2008, 180: 1367–1378. Li J, Zhao X, Christensen B M. Dopachrome conversion activity in Aedes aegypti: significance during melanotic encapsulation of parasites and cuticular tanning. Insect Biochem. Mol. Biol., 1994, 24: 1043–1049. Llopart A, Lachaise D, Coyne J A. Multilocus analysis of introgression between two sympatric sister species of Drosophila: Drosophila yakuba and D. santomea. Genetics, 2005, 171: 197–210. Prud'homme B, Gompel N, Carroll S B. Emerging principles of regulatory evolution. Proc. Natl. Acad. Sci. USA, 2007, 104 Suppl. 1: 8605–8612. Prud'homme B, Gompel N, Rokas A, et al. Repeated morphological evolution through cis-regulatory changes in a pleiotropic gene. Nature, 2006, 440: 1050–1053. Simpson P. The stars and stripes of animal bodies: evolution of regulatory elements mediating pigment and bristle patterns in Drosophila. Trends Genet., 2007, 23: 350–358. Soshnev A A, Li X, Wehling M D, Geyer P K. Context differences reveal insulator and activator functions of a Su(Hw) binding region. PLoS Genet., 2008, 4: e1000159. Wittkopp P J, Vaccaro K, Carroll S B. Evolution of yellow gene regulation and pigmentation in Drosophila. Curr. Biol., 2002a, 12: 1547–1556. Wittkopp P J, True J R, Carroll S B. Reciprocal functions of the Drosophila yellow and ebony proteins in the development and evolution of pigment patterns. Development, 2002b, 129: 1849– 1858. Wittkopp P J, Williams B L, Selegue J E, Carroll S B. Drosophila pigmentation evolution: divergent genotypes underlying convergent phenotypes. Proc. Natl. Acad. Sci. USA, 2003, 100: 1808– 1813. Xia A H, Zhou Q X, Yu L L, et al. Identification and analysis of YELLOW protein family genes in the silkworm, Bombyx mori. BMC Genomics, 2006, 7: 195.

CHAPTER 8

Bursicon, a Neuropeptide Hormone That Controls Cuticle Tanning and Beyond Qisheng Song and Shiheng An

Abstract Insects are encased in a semi-rigid exoskeleton, which provides protection, locomotion, and internal attachments sites for muscles and internal organs. However, it also limits insect’s growth. Insect must shed its old exoskeleton periodically (molting) in order to grow. After each molt, the newly formed exoskeleton is usually soft, lightly pigmented and flexible, permitting insect to extend its body size for growth.Sclerotization of newly formed cuticle must occur in a relatively short period of time after each molt and completion of body expansion in order for the new exoskeleton to be functional for protection. Insects could not survive without properly hardened cuticle. Molting and cuticle sclerotization in insects are regulated by a precise coordination of at least 6 hormones including prothoracicotropic hormone (PTTH), 20-hydroxyecdysone (20E), eclosion hormone (EH), ecdysis triggering hormone (ETH), crustacean cardioactive peptide (CCAP) and bursicon. In our previous chapter (Song and Sun 2005 edited by Liu and Kang), we already introduced signal transduction pathways of PTTH, 20E and JH. Here in this chapter, we would like to summarize the molecular mechanisms of EH, ETH, CCAP and bursicon and their coordination in regulating the final steps of the molting process i.e. ecdysis and cuticle sclerotization. In particular, we will focus on the action of bursicon. The elucidation of the key biochemical events leading to cuticle sclerotization would add an important body of knowledge towards understanding insect development and for designing more efficient pest control strategies to disrupt the cuticle sclerotization process. Keywords prothoracicotropic hormone, hydroxyecdysone, juvenile hormone, eclosion hormone, ecdysis triggering hormone, bursicon

Qisheng Song, Shiheng An Division of Plant Sciences, University of Missouri, Columbia, MO, 65211, USA E-mail: [email protected]


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8.1 Introduction Insects are encased in a semi-rigid exoskeleton, which provides protection, locomotion, and internal attachments sites for muscles and internal organs. However, it also limits insect’s growth. Insect must shed its old exoskeleton periodically (molt) in order to grow. Molting is a process common to all arthropods, during which a larger new cuticle is synthesized and the old one is partially digested and cast off (ecdysis) allowing the animal to grow. The newly formed exoskeleton after each molt is usually soft, lightly pigmented and flexible, permitting insect to extend its body size for growth. Sclerotization (hardening and tanning) of newly formed cuticle must occur in a relatively short period of time after each molt and completion of body expansion in order for the new exoskeleton to be functional for protection. Insects could not survive without properly hardened cuticle. Studies on ecdysial behavior in insects showed that at least six different hormones are released in an orderly manner during the final molting cycle to regulate the synthesis and sclerotization of new cuticle (Truman 1992; Zanassi et al. 2001; Zitnan and Adams 2005; Frankel and Hsiao 1965), including prothoraci cotropic hormone (PTTH) ecdysteroid 20-hydroxyecdysone (20E), eclosion hormone (EH), ecdysis triggering hormone (ETH), crustacean cardioactive peptide (CCAP), and bursicon (Ewer et al. 2002). The final hormone released in this cascade is a neuropeptide, bursicon, which triggers sclerotization of the new cuticle. Although hormonal regulation of cuticular sclerotization in insects was first reported in 1962 (Cottrell 1962a; Fraenkel and Hsiao 1962), molecular identity of bursicon was not characterized until 2005. It is now clear that functional bursicon is a heterodimer consisting of two cystine knot subunits, referred to as bursicon (burs) or bursicon α (burs α) and partner of bursicon (pburs) or bursiocn β (burs β) (Lou et al. 2005; Mendive et al. 2005). Several excellent reviews summarize early physiological studies of bursicon action (Seligman 1980; Reynolds 1983) and recent advances in molecular identification and functional characterization of bursicon (Ewer and Renolds 2002; Honegger et al. 2008). This chapter will give a brief introduction to identification and functional characterization of bursicon and provide insight on novel bursicon functions beyond cuticle sclerotization and wing expansion.

8.2 Identification of bursicon 8.2.1

Discovery of bursicon

Hormonal control of cuticle sclerotization or tanning in insects was first reported in the blowfly, Calliphora erythocephala, by Cottrell (1962a) and Fraenkel and Hsiao (1962). When the blowfly was neck-ligated immediately after emergence, neck-ligation prevented the thoracic and abdominal cuticle from being tanned. In order to demonstrate the probable hormonal nature of the tanning factor,

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hemolymph collected from a fly shortly after emergence was injected into the posterior portion of a neck-ligated fly with unsclerotized cuticle. The cuticle of the later animal quickly darkened as a result of the hemolymph transfusion. Fraenkel and Hsiao (1965) concluded that a blood-borne factor must be responsible for triggering the cuticular tanning process of newly emerged flies and named the factor 'bursicon' (from the Greek bursikos, pertaining to tanning). The newly-emerged, neck-ligated blowflies with unsclerotized thoracic and abdominal regions were adopted as standard bioassay animals for detecting and localizing bursicon. Over the past four decades, bursicon activity and most recently bursicon sequences have been identified in many arthropods including insects, crustaceans and arachnids, suggesting conservation of the two bursicon monomers burs and pburs among arthropods (see reviews by Seligman 1980; Reynolds 1983; Ewer and Renolds 2002; Honegger et al. 2008). Surprisingly, burs and pburs sequences have also been identified in echinoidea, a non-arthropod class in the phylum echinodermata (Van Loy et al. 2007). All burs and pburs sequences identified in arthropod and non-arthropod species contain 11 cysteine residues in conserved positions, indicating a pivotal role in the structural integrity of these ligands (see review by Honegger et al. 2008). 8.2.2

Molecular nature of bursicon

The molecular nature of bursicon was first reported by Cottrell (1962b). Evidence from his studies indicated that the active factor from the blowfly Calliphora was nondialysable and relatively insoluble in organic solvents. It was also inactivated by alcohol and by bacterial protease Subtilisin, suggesting its protein or polypeptide nature. Fraenkel and Hsiao (1965) also found that bursicon in Calliphora could be precipitated like a protein with loss of activity in reagents such as trichloroacetic acid, alcohol and acetone. Full bursicon activity was retained after ammonium sulfate precipitation, but could be destroyed after treatment with trypsin or pronase in Manduca (Taghert and Truman 1982a). After the protein nature of bursicon was established, bursicon was thought to be a single protein with an estimated molecular size around 30–60 kDa, depending on gel filtration methods used and insect species analyzed (Fraenkel et al. 1966; Seligman and Doy 1973; Reynolds 1976, 1977; Mills and Lake 1966; Mills and Nielsen 1967; Taghert and Truman 1982a). Using more precise 2dimensional (2D) SDS-PAGE as the key approach, Kaltenhauser et al. (1995) characterized the Tenebrio molitor bursicon with a molecular weight of 30 kDa. Similarly, Kostron et al. (1995) also attributed bursicon activity to proteins around 30 kDa in species such as C. ervthocephala, Glycymeris bimaculata, Periplaneta americana, and Locusta migratoria. Using partially purified bursicon from the American cockroach, P. americana, Honegger et al. (2002, 2004) were able to obtain several partial amino acid sequences of P. americana bursiocn. By comparing the partial sequences with the available Drosophila genomic sequence, Dewey et al. (2004) identified the


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Drosophila melanogaster gene CG13419 as a candidate bursicon gene. A further study revealed that mutations in CG13419 showed defects in cuticle sclerotization and wing expansion in Drosophila adults (Dewey et al. 2004), indicating that CG13419 should be a bursicon gene. But the molecular weight of CG13419 product is about 15 kDa, which is only one-half molecular weight of the detected bursicon from gel filtration and 2D gel electrophoresis. This indicated that bursicon might be a dimer (Honegger et al. 2002). The break-through studies came simultaneously from two independent groups that CG15284, the only other cystine knot protein present in the Drosophila genome, was identified as the heterodimer partner of functional bursicon using the available CG13419 sequence (Luo et al. 2005; Mendive et al. 2005). These two groups reported that only the conditioned media of cells co-transfected with CG13419 and CG15284 initiated tanning of the neck-ligated flies. Thus, after over four decades’ effort, the functional bursicon has finally been identified as a heterodimer consisting of two cystine knot subunits named burs or burs α (CG13419) and pburs or burs β (CG15284) with molecular weight of 15 kDa each (Dewey et al. 2004; Luo et al. 2005; Mendive et al. 2005). 8.2.3

Localization of bursicon

To identify bursicon synthesis site, extracts from variety parts of central nerve system (CNS) and hemolymph samples at different time periods of test insects were injected into neck-ligated blowflies. Bursicon activity was demonstrated in almost all parts of CNS including brain, corpora cardiaca (CC) and corpora allata (CA) complex, and thoracic and abdominal ganglion, depending upon insect species analyzed (see reviews by Seligman 1980; Reynolds 1983; Ewer and Renolds 2002; Honegger et al. 2008). Taghert and Truman (1982) were able to bioassay single neuron for bursiocn activity and localized bursicon activity to four large neurons in the lateral region of each abdominal ganglion in Manduca sexta. Localization of bursicon in the central nerve system was not definitive until specific antibodies and DNA probes to burs and pburs were developed. Both burs and pburs have been shown to co-localize in a set of neurons in subesophageal, thoracic and abdominal ganglia in Drosophila and several other insect species (for details see the review by Honegger et al. 2008). Fig. 8.1 shows burs and pburs mRNA expression patterns in the subesophageal, thoracic and abdominal ganglia of different development stages of Drosophila and Musca domestica (Wang et al. 2008).

8.3 Mode of action of bursicon 8.3.1

Bursicon receptor

In recent years, a subfamily of G protein-coupled receptors, named LGRs, was

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Fig. 8.1 Fluorescence in situ hybridization of cuticle sclerotization neuropeptide bursicon α (red) and β (green) transcripts in the central nervous system (CNS) of the third instar larvae, prepupae and newly emerged adult of Drosophila melanogaster (right panel) and Musca domestica (left panel). The CNS samples were visualized under a confocal microscope and photographed. Yellow color in the merged pictures indicates the neurons with the co-localized bursicon α and β signals. Transparent pictures show the images of the CNS.

identified, which has a large ecto-domain containing leucine-rich repeats. Genes in this family are conserved in invertebrates and vertebrates (Nishi et al. 2000). LGRs can be divided into three groups: group A, vertebrate glycoprotein hormone receptors; group B, D. melanogaster LGR2 (DLGR2) (Eriksen et al. 2000) and vertebrate orphan receptors LGR4, 5, 6 (Hsu 2003); and group C, mammalian relaxin_INSL3 receptors (Hsu et al. 2002; Kumagai et al. 2002). Genetic analyses in D. melanogaster reveal that bursicon mediates the cuticle sclerotization and wing expansion process via a specific G protein–coupled receptor DLGR2, encoded by the rickets gene. Mutation of rickets or burs causes defects in cuticle sclerotization and wing expansion (Baker and Truman 2002; Dewey et al. 2004). It has been demonstrated recently that the recombinant bursicon (r-bursicon) heterodimer binds to and activates DLGR2, which in turn leads to dose-dependent intracellular increase in adenyl cyclase activity and cAMP production in the mammalian 293T cells and COS-7 cells that overexpress DLGR2 (Luo et al. 2005; Mendive et al. 2005). Thus DLGR2 has been well recognized as the bursicon receptor. 8.3.2

Bursiocn signaling pathway

Down-stream of bursiocn receptor DLGR2, cyclic AMP (cAMP) has been shown


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to be possibly involved in the bursicon-mediated cuticle sclerotization process. cAMP is an important secondary messenger for many signal transduction pathways. Earlier studies on blowflies have implicated an increase in cAMP upon the action of bursicon (extract or hemolymph) (Reynolds 1980; Seligman and Doy 1972, 1973; Von Knorre et al. 1972). Recent studies using an in vitro binding assay also indicated that r-bursicon stimulated cAMP production in a dosedependent manner (Luo et al. 2005). Injection of the rickets mutatedflies with a cAMP analog resulted in melanization of the abdominal tergites. These results strongly indicated that DLGR2 in fact is the receptor of bursicon and that DLGR2 mediates bursicon signaling pathway through cAMP. Upon activation by bursicon, DLGR2 is hypothesized to activate the cAMP/ PKA signaling pathway, eventually leading to cuticle sclerotization (Kimura et al. 2004; Luo et al. 2005; Mendive et al. 2005). Most recently, Davis et al. (2007) showed that tyrosine hydroxylase, the enzyme mediating the conversion of tyrosine to DOPA in the metabolic pathway leading to cuticle tanning, is activated by PKA via bursicon stimulation of DLGR2, providing another evidence that bursicon activates cAMP/PKA pathway via G-protein coupled receptor DLGR2 in Drosophila. Despite these reports, our knowledge of signaling components involved in the bursicon-stimulated cuticle sclerotization process remains rudimental.

8.4 Bursicon function 8.4.1

Cuticle sclerotization (tanning and hardening)

Bursicon was first described over four decades ago as a bioactive cuticle sclerotization factor present in the central nervous system (CNS) and hemolymph of newly emerged adults in a neck-ligated blowfly assay (Fraenkel & Hsiao 1962, 1965; Cottrell 1962a). The extracts from central nerve system or hemolymph of several dipteran species including, but not limited to, Sarcophaga bullata (Baker and Truman 2002; Cottrell 1962b; Fogal and Fraenkel 1969), Phormia regina (Fraenkel and Hsiao 1965), and Lucilia spp (Cottrell 1962b; Seligman and Doy 1972) have also been shown to have cross-species activities in neck-ligated fly assay. The bioactive cuticle sclerotization factor present in the central nervous system (CNS) and hemolymph of newly emerged adults has now been identified through molecular cloning and confirmed in the neck-ligated fly assay using rbursicon heterodimer (Luo et al. 2005; Mendive et al. 2005). The cross-species activity of bursicon has also been demonstrated using r-bursicon heterodimer (An et al. 2009). For example, Drosophila r-bursicon heterodimer expressed in mammalian cell line exhibited a strong tanning activity in the house fly, Musca domestica and vice versa (An et al. 2009). Surprisingly, lobster nervous system homogenates were reported to have bursicon activity in the ligated fly bioassay (Kostron et al. 1995) and this observation was supported by the presence of pburs-like transcript in the cDNA date base of the loster Homarus americanus (Van Loy et al. 2007).

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The cuticle sclerotization process in fact is an enzymes-mediated metabolic process. The enzymes participated in the process include, but not limit to, diphenoloxidases, laccases, peroxidase, tyrosine hydroxylase and dopa decarboxylase. In the red flour beetle, Tribolium castaneum, laccase 2 is the only phenoloxidase that its activity is necessary for cuticle sclerotization. RNAi of laccase 2 gene causes defects of cuticle sclerotization (Arakane et al. 2005). However, it should be pointed out that not all genes are regulated at the transcriptional level. For example, tyrosine hydroxylase mediates the conversion of tyrosine to dihydroxyphenylalanine (dopa) and is a key enzyme during the tanning process. However, tyrosine hydroxylase mRNA level is at its highest 24 h before eclosion and remains unchanged until 12 hours after eclosion (Davis et al. 2007). It seems that the activity of tyrosine hydroxylase during the sclerotization process is not regulated at the transcriptional level. Tyrosine hydroxylase is transiently activated during cuticle sclerotization by a post-translational mechanism, i.e. phosphorylation by PKA (Davis et al. 2007). Another example is dopa decarboxylase, which catalyzes the conversion of dopa to dopamine, a compound of central importance for sclerotization. The dopa decarboxylase mRNA level peaks at 24 hours before eclosion and decreases thereafter. It is almost unchanged or decreases minutely from 0 h to 3 h after eclosion. Dopa decarboxylase is transcribed and translated before eclosion to ensure that enzyme activity is present when substrate becomes available (Davis et al. 2007). 8.4.2

Wing expansion

In addition to inducing the insect cuticle sclerotization process, bursicon is also involved in insect wing expansion. The insect wing expansion process is another physiological event occurring after ecdysis and accompanying cuticle sclerotization. These processes occur at the same time, but the mechanisms under which they are controlled and regulated are still not clear. Recent research showed that mutation in the bursicon genes causes the failure of initiation of the behavioral program for wing expansion and results in defects in wing expansion in D. melanogaster (Dewey et al. 2004). Similarly, RNAi of burs gene also results in defects in wing expansion in the silkworm B. mori (Huang et al. 2007). These studies indicate that bursicon is the hormone required to initiate both cuticle sclerotization and wing expansion processes in insects. Another line of support to this observation is from genetic study of bursion receptor DLGR2. It has been shown that mutation of the rickets gene, which encodes the bursicon receptor DLGR2, inhibits wing expansion in Drosophila (Kimura et al. 2004). Although it is not clear how wing expansion in insects is controlled, it has been demonstrated that wing development and expansion are strongly associated with programmed cell death and removal of cell debris from the wing tissue (Johnson and Milner 1987; Kiger et al. 2007; Kimura et al. 2004; Natzle et al. 2008; Seligman et al. 1972, 1975). Shortly after eclosion, hemolymph pressure forces the expansion of the folded wing blade. At the same time, the dorsal and ventral


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epithelial cell layers delaminate from the outside cuticle, undergo a process called epithelial-mesenchymal transition, and then exit the wing accompanied by the initiation of a programmed cell death under the control of a yet to be identified signaling pathway (Kiger et al. 2007; Natzle et al. 2008). The wing epidermis is removed by cell death at the time of wing spreading in the large fly Lucilia cuprina (Seligman et al. 1975) and in D. melanogaster (Johnson and Milner 1987). Using a TUNEL assay and transmission electron microscopy, Kimura et al. (2004) demonstrated that cell death was accompanied by DNA fragmentation and that this cell death exhibited extensive vacuoles, implying possible autophagy. Ectopic expression of an anti-apoptotic gene, p35, inhibited the cell death, indicating the involvement of caspases. It was reported that both cell death and the removal process is controlled by changes in the levels of the steroid hormone 20-hydroxyecdysone (20E) through regulation of a series of early response and late response genes, including the expression of the cell death activator genes, rpr and hid, which then repress the Drosophila inhibitor of apoptosis proteins (IAPs)Diap2 (Jiang et al. 1997). Changes in 20E titer, in combination with the fine-scale expression of the ecdysone receptor, control the timing and spacing of cell death during metamorphosis. Though ecdysone-dependent regulation is the possible pathway controlling cell death, it does not exclude the possibility of the involvement of other hormones in wing expansion and programmed cell death. It is very likely that bursicon co-regulates the programmed cell death with other factors during the post-eclosion wing maturation process. It has been shown that stimulation with components downstream of bursicon, such as a membrane permeate analog of cAMP, or ectopic expression of constitutively active forms of G proteins or PKA, induces precocious cell death; and conversely, cell death was inhibited in wing clones lacking G protein or PKA function. Therefore, bursicon activation of the cAMP/PKA signaling pathway is likely required for transduction of the hormonal signal that induces wing epidermal cell death after eclosion (Kimura et al. 2004).

8.5 Evidences for bursicon’s roles beyond cuticle tanning and wing expansion Bursicon mediates cuticle sclerotization and wing expansion via a specific G protein-coupled receptor DLGR2, encoded by the rickets gene (Baker and Truman 2002; Dewey et al. 2004; Luo et al. 2005; Mendive et al. 2005). DLGR2, once activated, is hypothesized to activate the cAMP/PKA signaling pathway, leading to phosporylation of tyrosine hydroxylase, the enzyme mediating the conversion of tyrosine to DOPA in the metabolic pathway leading to cuticle tanning (Davis et al. 2007). To identify signaling pathway downstream of the bursicon receptor DLGR2 and adenylate cyclase, as well as the genes regulated by bursicon, An et al. (2008) performed a DNA microarray analysis in the neckligated flies (Fig. 8.2) using r-bursicon as a probe (Fig. 8.3) and identified a set of

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87 genes (Table 8.1) whose expression were up or down regulated by r-bursicon in D. melanogaster. Among total 87 transcripts, 54 were identified at 1 h post rbursicon injection and 33 at 3 h post r-bursicon injection. Table 8.1 Microarray identification of the genes that are up- and down-regulated by r-bursicon in the neck-ligated fly assay at 1 and 3 h post r-bursicon injection (³2.0 folds, p < 0.05) (from An et al. 2008). 1h

3h

Function

Transcript

Gene name or function domain

Fold

Transcript

Gene name or function domain

Fold

Signaling

CG2849-RB

Ras-related protein

3.9

CG17226-RA

Odorant receptor 59c

– 2.7

CG6407-RA

Wnt oncogene analog 5

2.3

CG11348-RB

Nicotinic acetylchol. receptor

2.1

CG32659-RA

Tenascin accessory (EGF- Domain)

2.1

CG2346-RA

FMRFamiderelated

2.0

CG10125-RA

Zero population growth (Innexin domain)

2.0

CG6386-RA

Ballchen (Kinase domain)

2.0

CG10146-RA

Attacin

7.9

CG31193-RA

Humoral factor Turandot X

3.8

CG33202-RB (Immunoglobulin domain)

2.3

CG31691-RA

Humoral factor Turandot F

2.3

CG18372-RA

2.0

CG31508-RA

Humoral factor Turandot C

2.2

CG1878-RA

Cecropin B

2.3

Immune response

Channel & transporter

Attacin

CG9500-RA

(Fibrinogenrelated domains)

2.4

HDC14466

Magnesium/ cobalt transporter

3.0

CG12348-RA

Shaker ( BTB/POZ domain )

2.1

CG10369

Inward rectifier K+ channel

2.3

CG4110-RA

Pickpocket 11 (Sodium channel)

2.0

CG33098-RC

Ca2+ sensors & Ca2+ signal MODs

2.1


141

(Continued) 1h

Transcription

3h

CG1522-RA

Cacophony (Ion transport protein)

2.0

HDC02744

Iron-sulfur cluster-binding protein

2.0

CG14872-RB

(Lipocalin /cytosolic fattyacid binding protein)

2.1

CG10844

Ryanodine-sensitive Ca2+-release channel

– 2.6

CG8221-RA

Amyrel (Alpha amylase)

– 2.5

CG30080-RA

Iduronate-2- like sulfatase

3.0

CG30080-RA

Iduronate-2- like sulfatase

2.1

CG32491-RA Modifier of mdg4

2.4

CG32491-RA Modifier of mdg4

2.4

CG11501-RA

Transcriptional regulator

9.6

CG1624-RC

Dappled (Zinc binding domain)

2.0

CG6157-RA

Discontinuous actin hexagon

2.0

(Zinc binding domain)

DNA/RNA binding

Metabolic enzyme

CG8260-RA

(BTB/POZ domain)

2.0

CG3497-RA

Suppressor of Hairless (DNA binding domain)

2.4

CG12924-RA

Eukary. Sm & Sm-like (LSm) protein

3.2

CG3019-RA

Suppressor of white-apricot

2.1

CG32438-RA

SMC5 proteins (ATPase family)

– 2.2

CG8920-RA

Tudor domain

2.1

CG7776-RA

Enhancer of Polycomb

2.0

CG10897-RA Toutatis (MethylCpG binding domains)

2.0

CG10327-RC

TBPH (RNA recognition motif)

2.0

CG3238-RA

(DUF889 domain)

2.6

CG12660-RA

Retinal degeneration A (Phorbol-esters /diacylglycerol binding domain)

2.5

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(Continued) 1h

3h

CG13927-RA

γ-glutamyl carboxylase

2.3

CG10140-RA

Chitin binding Peritrophin-A domain

2.2

CG6822-RB

Ergic53(Legume lectin domain)

2.1

CG4181-RA

Glutathione S transferase

2.1

CG3649-RA

Sugar transporter domain

2.0

Cytoskeletal compnt

CG13338-RA

Cuticular protein 50Ca (Chitin-binding domain)

2.1

CG32050-RA

Titin

2.0

Cell adhesion

CG3938-RE

Cyclin E

2.2

CG16826-RA

Lipoprotein

2.2

CG7294-RA

Trophinin

2.0

CG4793-RB

Trypsin-like serine protease

4.9

CG11066

Serine-type endopeptidase

2.9

CG30287-RA

Trypsin-like serine protease

2.1

Proteolysis & peptidolysis

Unknown function

CG30482-RA

N

2.8

CG7985-RA

N

2.6

CT32157

N

2.4

HDC18092

N

2.6

CG16863-RA

N

2.3

HDC15090

N

2.4

HDC03087

N

2.2

HDC13939

N

2.3

HDC16822

N

2.2

HDC15545

N

2.2

CG32027-RA

N

2.2

HDC05639

N

2.0

HDC09490

N

2.2

HDC14431

N

– 2.0

LD33458

N

2.2

HDC09972

N

– 2.1

CG2217-RA

N

2.1

CT33997

N

– 2.2

CG17816-RA

N

2.1

HDC18592

N

– 2.5

CG40172-RA

N

2.1

CG4440-RA

N

– 2.5

CG31813-RA

N

2.1

S.CX001941

N

– 4.0


143

(Continued) 1h GH12319

3h N

2.0

CG32132-RA

N

2.0

CT29521

N

2.0

CG15699-RA

N

2.0

CG40386-RA

N

2.1

HDC10707

N

– 2.2

The genes in bold were detected at both 1 and 3 h post r-bursicon injection. The name in parenthesis indicates a functional domain. N: gene with no known function associated with it.

Fig. 8.2 Functional assay of the r-bursicon heterodimer in neck-ligated flies. Newly emerged flies were ligated between the head and thorax at emergence and injected with 0.5 ml of cell culture transfected with blank pcDNA 3.1 vector as a sham control (a) or with the purified r-bursicon α (b) or r-bursicon β (c) or r-bursicon heterodimer expressed in insect High Five™ cells (e) and in mammalian HEK293 cells (f). A CNS homogenate (0.5 CNS equivalent/fly) from newly emerged flies was used as a positive control (d). The arrow indicates the area with the unsclerotized cuticle (light color) in control (a-c) and the sclerotized cuticle (darkened) in the flies injected with Drosophila r-bursicon heterodimer. A strong cuticle tanning was observed when the neck-ligated house flies were injected with Drosophila r-bursicon or Drosophila CNS homogenate (bottom panel).

Analysis of these genes by inference from the fly database (http://flybase.bio. indiana.edu) revealed that only 57 genes show significant similarities to known proteins or functional domains in the Drosophila data base while 30 did not. The 57 known genes encode proteins with diverse functions, including cell signaling, gene transcription, DNA/RNA binding, ion trafficking, proteolysis-peptidolysis, metabolism, cytoskeleton formation, immune response and cell-adhesion. Most

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Fig. 8.3 Western blot analysis of recombinant bursicon heterodimer proteins expressed from mammalian 293 cells after SDS-PAGE. Lane 1–3 refers to burs α subunit only, burs β subunit only, and burs α + β expressed together, respectively.

of these known genes could be linked to cuticle sclerotization and wing expansion, directly or indirectly. For example, 8 genes are found to control cell signaling and these genes include a Ras-related protein (CG2849-RB), a Gprotein activator-odorant receptor 59C (CG17226-RA), etc. Ras proteins are considered to be very important molecular switches for a wide variety of signaling pathways that control such processes as cytoskeletal integrity, proliferation, cell adhesion, apoptosis, and cell migration (Howard et al. 2003) Ras-related protein is often activated via phosphorylation by PKA. Perhaps in Drosophila the Ras-related protein CG2849-RB identified in the microarray analysis (Table 8.1) might be activated directly by PKA i.e. act as a downstream component of PKA in the bursicon signaling pathway. Another example is Su(H) gene. We have shown that Drosophila r-bursicon regulates the expression of Su(H) gene in both Drosophila (An et al. 2008) and Musca (Wang et al. 2009). Su(H) is one of the integral components in the Notch signaling pathway, a cell signaling system present in most multi-cellular organisms (Fortini and Artavanis-Tsakonas 1994). During insect development, Su(H) and Notch signaling pathway control alternative cell fate in adult epidermis and pupal notum cells of Drosophila (Schweisguth 1995; Schweisguth and Posakony 1994). Su(H) contributes to imaginal disc-derived wing morphogenesis and wing vein formation (Curtiss et al. 2002; Crozatier et al. 2003; Johannes and Preiss 2002; Klein et al. 2000). Su(H) influences expression of several cuticlepatterning genes, such as Dfrizzled 2, hairy, shaggy and patched, in Drosophila (Wesley 1999). Su(H) is also involved in cell proliferation and cell death in development of insect wings and other tissues (Giraldez and Cohen 2003). The development and expansion of insect wings after adult emergence is to a great extent related to the programmed cell death of wing hypodermis and to the


145

removal of the cell debris (Johnson and Milner 1987; Kiger et al. 2007; Kimura et al. 2004; Seligman and Doy 1972; Seligman et al. 1975). The factor that triggers wing epidermal cell death is probably bursicon (Kimura et al. 2004). These results, together with the involvement of Su(H) and Notch signaling pathway in controlling wing cell proliferation and death, suggest the likely crosstalk between the Su(H)/Notch and the bursicon-induced cuticle sclerotization and wing development processes. However, some identified genes could not be related to the cuticle sclerotization and wing expansion processes, suggesting that bursicon might have unidentified novel functions. From the microarray identified genes (Table 8.1), seven immune-related genes were up-regulated by r-bursicon. These genes include 3 from turandot gene family (turandot X, turandot F and turandot C), 2 from attacin gene family and 1 each from cecropin and immunoglobulin families (An et al. 2008). The turandot gene family is considered to be immune-related in insects and is induced under a wide range of adverse conditions such as infection, heat, oxidizing agents, wounding and aging (Ekengren et al. 2001) while attacin, cecropin and immunoglobulin are well characterized anti-microbial peptides in insect immune system. Although no direct association between bursicon and antibacterial peptides is obvious, the newly ecdysed insect is soft (before cuticle tanning and hardening) and more susceptible to injury and infection. Perhaps a more vigorous anti-bacterial defense is necessary at this time when there may be perforations in the soft cuticle due to predators or simple contact injuries. Up-regulation of the immune response genes by r-bursicon suggests, certainly not approve, bursicon’s role in mediating immune defense mechanism in insects. Bursicon is a heterodimer consisting of two structurally different subunits. It is quite possible that in addition to its classic roles in cuticle sclerotization and wing expansion, each subunit may have its own physiological function. As a matter of fact, our preliminary data indicate that each bursicon subunit indeed induces a different set of genes in the final molting cycle of D. melanogaster (unpublished information). Although bursicon’s roles in larval and pupal development have not been explored, the fact that each of bursicon subunits (burs and pburs) is expressed throughout larval and pupal stages, also suggest their potential roles in larval and pupal development.

8.6 Prospect The studies of bursicon have been over four decades since its discovery. Physiological and molecular characterizations of bursicon have been wellstudied, but many questions still remain unsolved. For examples, what are the functions of one third of bursicon-regulated unknown genes identified in DNA microarray? What signal transduction components are involved in bursiconstimulated cuticle sclerotization and wing expansion? Does bursicon regulate immune response genes in insects? Does bursicon have a role in larval and pupal

146


development? Does each bursion subunit have its own physiological function? With the recombinant bursicon protein, well-established bursicon bioassay systems, and available genetic and molecular approaches, the answers to these questions will be unfolded in the near future.

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(H) in the house fly Musca domestica. Insect Science, 2009, 16: 207–217. Wesley C S. Notch and Wingless regulate expression of cuticle patterning genes. Mol. Cell Biochem., 1999, 19: 5743–5758. Zanassi P, Paolillo M, Feliciello A, et al. cAMP-dependent protein kinase induces cAMPresponse element-binding protein phosphorylation via an intracellular calcium release/ERK-dependent pathway in striatal neurons. J. Biol. Chem., 2001, 276: 11487–11495. Zitnan D, Adams M E. Neuroendocrine Regulation of Insect Ecdysis. // Gilbert, L.I., Iatrou K, S.S., Gill, S.S. Comprehensive Insect Biochemistry, Physiology, Pharmacology, and Molecular Biology, Volume 3. London: Elsevier Press, 2005: 1–60.

CHAPTER 9

Chemical Ecology of Bark Beetles in Regard to Search and Selection of Host Trees John A. Byers and Qinghe Zhang

Abstract Bark beetles (Coleoptera: Scolytidae), especially pests in the genera Dendroctonus, Ips, Scolytus, Trypodendron, Tomicus, and Pityogenes of the Northern hemisphere are reviewed regarding aspects of their chemical ecology during host finding and selection. Most of the species covered here feed on conifers, primarily pines (Pinus) in the Northern hemisphere and Norway spruce (Picea abies) of Europe and Asia. Bark beetles use a variety of olfactory strategies to discriminate suitable host trees from among less suitable, overcolonized, or decaying hosts as well as nonhosts. Bark beetles also use olfactory strategies to find mates and select attack sites. These strategies have implications for coevolution of trees and bark beetles. Knowledge of the chemical ecology of insect-insect and insect-plant relationships is necessary to develop improved methods for monitoring and controlling bark beetles that are predators of trees. Keywords host selection, pheromones, semiochemicals, olfaction, coleoptera, scolytidae, host finding, mate location, competition, monoterpenes

9.1 Introduction Conifer-inhabiting bark beetles (Scolytidae) are among the most economically important forest insects, mainly due to species that infest and kill apparently healthy standing trees. The problems with bark beetle outbreaks are serious in many parts of the world as a result of air pollution, global climate change, and John A. Byers Arid-Land Agricultural Research Center, USDA-ARS, 21881 North Cardon Lane, Maricopa, Arizona, 85238, USA Qinghe Zhang Sterling International, Inc., 3808 N. Sullivan Rd, Bldg 16p, Spokane, WA 99216-1630, USA E-mail: [email protected]


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many other factors influencing forest health. The Scolytidae (bark and ambrosia beetles) is considered a family of beetles, although some think these beetles are a subfamily of weevils (Curculionidae). In any case there are at least 6,000 species that, although appearing similar, may differ widely in their ecology and adaptations to host trees (S.L. Wood 1982 2007; Byers 2004). Bark beetle species that feed on phloem tissue (a few mm-thick layer just below the exterior bark) are usually found only on one or a few host tree species, whereas ambrosia beetle species that are xylomycetophagous (wood-fungus feeding) and use symbiotic fungi for "cultivation" in their galleries generally colonize a larger range of hosts (S.L. Wood 1982). Most biological knowledge on bark and ambrosia beetles comes from studies on relatively few pest species in the genera Dendroctonus, Ips, Scolytus, Xyleborus, Trypodendron, Tomicus = Blastophagus (Fig. 9.1), Pityogenes, Hypothenemus, Pityophthorus, Hylastes, and Gnathotrichus.

Fig. 9.1 Electron micrograph of male (top) and female Tomicus piniperda from Sweden.

Most of the pest species are obligate or facultative tree-killing bark beetles that comprise only about 10% of scolytid species in the United States and Canada (Raffa et al. 1993). However, these species that kill trees are the most likely to significantly influence the evolution of the host trees and the forest ecosystems. The diversity of bark beetle biology, in which each species is adapted to only one or a few host tree species, probably has resulted from natural selection pressures from the great variety of toxic tree biochemicals. Each species of tree may have coevolved various chemicals to defend against the selection pressures of the treekilling bark beetles (Byers 1989a, 1995, 2004). Semiochemicals (behavior modifying chemicals) from both the tree and the beetle have many functions during the life cycle of a bark beetle (for reviews see D.L. Wood 1982; Borden 1997; Byers 1989a, 1989b, 1995; 2004; Raffa et al. 1993; Schlyter and Birgersson 1999; Zhang and Schlyter 2004). Host and non-host plant chemicals can be attractive, repellent, toxic, or nutritious to bark beetles. These chemicals

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may have affects on: (1) finding and accepting the host tree (host selection and suitability), (2) feeding stimulation and deterrence, (3) plant resistance, (4) pheromone/allomone biosynthesis and communication, and (5) attraction of predators, parasites and competitors of bark beetles. In this chapter, we will focus primarily on bark beetle finding and selecting the host tree. Knowledge of bark beetle chemical ecology and insect-tree relationships is important to developing methods of managing bark beetle populations and their colonization damage to trees. The strategies that bark beetles use to avoid competition within and between species, to avoid unsuitable host and nonhost trees, to find their host trees and mates, and to maximize their reproductive success can be investigated with the purpose of eventually manipulating these processes to the disadvantage of the beetles. Most of the following presentation involves species in the genera Dendroctonus, Tomicus, Ips, and Pityogenes. In general, adults of bark beetles in the above genera overwinter in either forest litter (e.g., Ips, Pityogenes) or the brood tree (e.g., Dendroctonus, Ips, Pityogenes). In species that have several generations during the summer, emergence is from the brood tree. Tomicus piniperda has a more complex life cycle in which adults over-winter in living, nonbrood trees. After emergence, the adults of all species attempt to locate a host tree (termed the dispersal flight), either by orienting to pheromone or plant volatiles. Host suitability may be determined in flight or after landing on the tree. In the monogamous genera Tomicus and Dendroctonus (subfamily: Hylesininae), the females select the host tree and an entrance hole (attack) to begin construction of oviposition galleries in the phloem. In contrast, males of the polygynous genera Ips and Pityogenes (subfamily: Scolytinae) begin the entrance hole and later accept several females. In most cases, individuals of only one sex begin the attack, releasing a speciesspecific blend of chemicals comprising an aggregation pheromone (Byers 1989a, 1995, 2004). However, in D. brevicomis, the female and the joining male each produce a unique synergistic pheromone component that when combined elicit maximal attraction response. In T. piniperda, there is no evidence of an aggregation pheromone (Byers et al. 1985); instead, monoterpenes from the hosttree induce aggregation (Fig. 9.2). Species of bark beetles feed only on one or a few host tree species, which are perceived and located by means of the beetle’s various sensory receptors that are located on their antennae and mouthparts. Except for morphological studies on D. ponderosae and I. typographus, little is known about the sensilla on the maxillary and labial palpi mouthparts of bark beetles. In these species there is clearly a large number of chemosensilla, which appear to be important for host selection and food discrimination. In other insects, the tarsi and ovipositor have chemosensilla that are involved in host acceptance (Städler 1984), but these structures have been little studied in bark beetles. Most work with bark beetles has focused on the antennae (Fig. 9.3), which are known to have sensilla responsive to volatile pheromone and host components, as well as other air-borne chemostimulants (Payne 1979, Mustaparta 1984, Faucheux 1989).


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Fig. 9.2 Storm-damaged Scotch pine, Pinus sylvestris, releases monoterpenes attractive to both sexes of Tomicus piniperda in the early spring.

Fig. 9.3 Electron micrograph of antenna and eye of Tomicus piniperda.

Vision plays a vital role during orientation flights of bark beetles, and in conjunction with antennae enable bark beetles to locate semiochemical sources. The eyes of many bark beetles (e.g., Ips, Scolytus, and Pityogenes) consistently have only about 100–240 ommatidia (Fig. 9.3), which is less than many insects (Lanier 1983; Byers et al. 1989a). Based on electrophysiological recordings, two color receptor types have been identified in the eyes with a maximum absorbance at 450 nm (blue) and 520 nm (green) (Groberman and Borden 1982). Observations of I. paraconfusus, I. typographus, D. brevicomis, P. chalcographus, and T. piniperda in flight chambers under dim red light or in complete darkness using an electronic vibration detector indicate they will not fly after dark (Lanne et al. 1987; Byers and Löfqvist 1989; Byers unpublished). Bark beetles are attracted in greater numbers to traps baited with host odor or pheromone that are placed next to "tree trunk silhouettes" than to traps without such visual stimuli, indicating that beetles orient to the tree trunk during landing (Tilden et al. 1983). Beetles of some species prefer to land on horizontal silhouettes rather than

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on vertical ones of the same size. Another indication that bark beetles have relatively poor visual acuity is that T. piniperda males must walk within 1 cm of a female beginning her entrance hole before they appear to detect her and initiate guarding behavior (Byers 1991). Both T. piniperda and D. brevicomis individuals can be induced to drop off a tree by movements of the human body about 2 m away (about the same angle of resolution and relative size, Byers unpublished). Bark beetles search for breeding habitat by flying toward odiferous sources of host chemicals or aggregation pheromone (Fig. 9.4). We consider long-range attraction in bark beetles to be flight orientation using optomotor anemotaxis (Byers 1996b) over a meter or more to a semiochemical source. In optomotor anemotaxis, a bark beetle attempts to fly directly upwind when in contact with a packet of pheromone-laden air of the plume, but casts (flies from side to side with respect to the source) when contact is lost. The beetle senses the wind direction while flying by observing the ground below: in no wind, or head-on wind, the ground moves directly underneath during flight. However, if the visual ground field also moves from right to left somewhat, for example, then wind is coming from the left, and the beetle turns to the left to minimize the transverse ground shift and keep the ground moving directly underneath, in this manner the insect heads upwind and toward the pheromone source (Byers 1996b). After the beetle orients and lands on a host tree and begins to release an aggregation pheromone, the likelihood of successful colonization depends on (1) the population level of beetles available for recruitment to the attack and (2) the resistance and health of the tree and its ability to produce defensive resin (Fig. 9.5). Resistance of conifers, especially pines, to bark beetle attacks has long been attributed to the amount of resin exuded and pitch tubes formed (Hodges et al. 1985). Dead beetles can often be seen in crystallized resin of pitch tubes. However, species of Dendroctonus and other aggressive bark beetles have a great ability to survive the "toxic" monoterpenes and suffocating mucilage and may struggle for hours in copious resin flows (D. frontalis, Hodges et al. 1979, D. brevicomis, Byers 1995). Drought and poor water balance lower the resistance of conifers (Hodges et al. 1979) probably by lowering the turgidity of resin duct cells, which lowers the oleoresin exudation pressure (OEP). Oleoresin and the monoterpenes indicate host resistance and therein are repellent to bark beetles in concentrated amounts (Byers et al. 2000; El-Sayed and Byers 2000; reviewed in Byers 1995, 2004). Chemodiversity of the tree may play a role in acceptance or resistance of the host tree: ponderosa pines with fewer different types of monoterpenes escaped colonization by Ips lecontei more often than trees with more kinds of monoterpenes (Thoss and Byers 2006). Successful colonization and reproduction by a bark beetle in a living tree requires release of enough aggregation pheromone to ensure the attraction of sufficient conspecifics to overwhelm the host tree defenses (Fig. 9.5); however, after killing the tree and securing mates, pheromone should stop in order to avoid further competition for bark areas (Byers et al. 1984; Berryman et al. 1985).


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Semiochemicals play a role in "cooperation" among beetles when killing the tree and in their avoidance of competition (discussed later). “Pioneer” beetles that attack the tree first may suffer most from the tree's defensive resin, but these beetles may have no choice but to attack due to low fat reserves. The later that a beetle arrives in the colonization sequence of the host, the poorer is the quality of the bark substrate due to (1) space utilization by established conspecifics (intraspecific competition) and (2) degradation by microorganisms (discussed subsequently). The question arises then: How does a newly emerged pioneer bark beetle find a suitable breeding host tree in which to produce aggregation pheromone and draw in other individuals in a mass attack. The process begins with dispersal from the brood tree or overwintering site.

9.2 Dispersal from the brood tree or overwintering site and search for host trees Insects disperse when their habitat becomes unsuitable. This can be from a lack of food resources, mating possibilities, territories and suitable domiciles, or from the need to escape the local buildup of parasites and predators. Apparently for the same reasons, bark beetles emerge from the dead brood tree, or litter near the brood tree, and begin a dispersal flight seeking suitable host trees from among many non-host and unsuitable host trees (Fig. 9.6). During dispersal, bark beetles and associated predators and parasites feeding or living in brood trees must locate a new host tree from among the relatively few widely scattered suitable hosts in the forest (Fig. 9.7). The host tree is restricted usually to one or a few species and in most cases the bark beetles seek weakened, less resistant trees, or trees that are in the initial stages of death and decay. Also, beetles try to avoid feeding and reproduction in areas heavily colonized by conspecifics and competing species (Byers 1984, 1995, Fig. 9.8). Thus, it is expected that species have evolved behavioral responses to volatile host-plant chemicals that indicate the presence of a suitable host in which reproduction can occur. The dispersal flight of a bark beetle may vary from only a few meters (as observed during epidemics) to possibly several kilometers. Several factors interact to cause the dispersal flight distance to vary between individuals. The most obvious is that a beetle encounters a susceptible tree early in the dispersal flight. However, whether this tree is attacked may depend on the level of fat reserves that can be mobilized for flight (Byers 1999). A beetle should have higher reproductive fitness if it flies far from the brood tree since it can both avoid inbreeding with siblings and, probably more importantly, escape predators and parasites that are locally denser near the brood tree. Thus, the dispersal distance has been optimized over evolutionary time to balance the probably logarithmically increasing benefits of flying farther against the probably exponentially increasing likelihood of exhaustion and failing to find a host. The fat level

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Fig. 9.4 Pheromone components of bark beetles. Key: aggregation component (a), inhibitor of aggregation (i). Row 1: 2-methyl-3-buten-2-ol (a, I. typographus ); 3methyl-3-buten-1-ol (a, I. cembrae), 4-methyl-3-heptanol (a, S. multistriatus). Row 2: sulcatol (a, G. sulcatus, G. retusus); seudenol (D. pseudotsugae, D. rufipennis, D. simplex); MCH (i, D. pseudotsugae); lanierone (a, I. pini). Row 3: cis-verbenol (a, I. paraconfusus, I. typographus, I. calligraphus); trans-verbenol (a, D. ponderosae, T. minor; i, D. brevicomis); verbenone (i, Dendroctonus); chalcogran (a, Pityogenes). Row


Fig. 9.5

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Dendroctonus brevicomis in resin of ponderosa pine.

Fig. 9.6 Ips typographus preparing to begin the dispersal flight after emerging from the duff in late May in Sweden.

required for lengthy dispersal will depend on the conditions in the brood tree during larval development; for example, disease, insect, and climatic factors will affect the nutritional quality of the host (Fig. 9.9). Severe competition among the larvae will reduce the size of adults as well as their fat content (Anderbrant et al.

4: ipsenol (a, Pityokteines curvidens, and many Ips: e.g. I. paraconfusus, I. grandicollis), ipsdienol (a, many Ips, e.g. I. paraconfusus, I. duplicatus, I. pini, I. calligraphus, I. avulsus); amitinol (a, I. amitinus); E-myrcenol (a, I. duplicatus). Row 5: (+)-exo-brevicomin (a, D. brevicomis, Dryocoetes); (-)-exo-brevicomin (a, Dryocoetes); methyl decadienoate (P. chalcographus). Row 6: frontalin (a, many Dendroctonus); endo-brevicomin (i, D. frontalis); multistriatin (a, S. multistriatus); lineatin (a, T. lineatum). References to above pheromones are in reviews by Byers (1989a, 2004).

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Fig. 9.7 Many species of bark beetles disperse through a stand of Norway spruce in the spring (Torsby, Sweden).

Fig. 9.8 Spacing of attacks of Ips typographus (larger circles) and Pityogenes chalcographus (smaller circles) to avoid competition in the bark of Norway spruce (bark surface scraped smooth on left to reveal attacks that were circled with ink pen).

Fig. 9.9 Theoretical curve for the acceptance of host trees by bark beetles depending on prerequisite flight exercise (asymptotic y-axis) and level of fatigue (amount of flight) and suitability of the host for reproduction (which depends on nutritional quality and density of colonization by competing bark beetles).


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1985). Parasites would reduce the size and fat content of some adults while predators would lessen competition for those remaining locally, thereby increasing the variability of dispersal range in the population. The population density of bark beetles should be stabilized by a frequency-dependant competition for the susceptible trees. This would produce increasingly stronger, longer-flying individuals with decreasing attack and larval density while producing weaker, shorter-flying individuals with increasing competition. Knowledge of how far and where bark beetle populations disperse comes mainly from (1) mark-release-recapture studies using pheromone traps and from (2) the geographical occurrence of new infestations relative to previous ones. Both lines of investigation are inconclusive since (1) only a few pheromone traps were used, usually some tens to hundreds of meters from the release site, so that a large proportion of released beetles escaped, or (2) the origins of attacking beetles were uncertain. Several studies have placed various sized rings of pheromone traps around a source of marked beetles. For example, the spruce bark beetle of Europe, I. typographus, was recaptured at various outer distances from 120 to 1000 m (Zumr 1992; Zolubas and Byers 1995; Duelli et al. 1997). In California, I. paraconfusus was recaptured in outer traps at 2 km (Gara 1963). The ambrosia beetle, Trypodendron lineatum, was recaptured at 500 m (Salom and McLean 1989). As expected, the widely spaced outer traps captured a small proportion of the released beetles, and the large gaps between traps probably allowed many to slip through as they drifted with the wind (e.g. gaps of 785, 1257, and 393 m in Zumr 1992; Gara 1963; and Salom and McLean 1989, respectively). An adverse effect of marking, although discounted, might also influence the dispersal. The view that bark beetles can fly some tens of km is based less on markrecapture studies and more on collections of beetles far from forests. Nilssen (1978) found two I. typographus in the stomach of a salmon 35 km from any spruce forest. Miller and Keen (1960) report results of studies by the US Forest Service in California where the western pine beetle, D. brevicomis, infested ìslands' of ponderosa pine, initially free of beetles, that were separated from the main forest by open sagebrush areas. They concluded that significant numbers of bark beetles must have flown a minimum of 3.2 km in one study, and 9.6 or 20 km in another study, to reach the infested trees. Increasing competition among larvae due to increasing densities of parents laying broods was shown to reduce size and fat content of bark beetles (Anderbrant et al. 1985) and thus should decrease dispersal ranges. However, this seems in conflict with the statement of Forsse (1991) that flying time of I. typographus on flight mills was "similar among populations and appeared unaffected by outbreak conditions". Earlier, Forsse and Solbreck (1985) could not find any affect of sex or body size on the duration of flight on mills. Botterweg (1982) also concluded that there was little, if any, affect of beetle size or fat content on dispersal distance as monitored in field traps. However, he did find that fat content of beetles declined over the flight period. This was probably due to consumption of fat during host-seeking rather than later emergence of lower-fat

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beetles since beetle's sizes (elytral weights) did not decrease over the spring season. Birgersson et al. (1988) reported that newly emerged I. typographus averaged about 10% fat, while after 24 hours of flight exercise in a plastic box, they declined to only 5% since fat is used for energy. In trees, males with nuptial chambers had about 8% fat (possibly replenishing some after feeding), but after several more days of feeding after females had joined them, the males had 10% fat again. Bark beetles appear capable of flying quite far in the forest since newly emerged D. pseudotsugae flew an average of 2 h on flight mills before resting (3 h total), while some individuals flew up to 8 h uninterrupted (Atkins 1961). Jactel and Gaillard (1991) flew I. sexdentatus on rotary flight mills and found that 50% of the beetles could fly more than 20 km, and 10% more than 45 km, based on about 50 interrupted flights. About 25% of I. typographus taken from litter in an outbreak area flew for over 1 h, and 10% for more than 2.5 h on flight mills, with a maximum flight of 6 h and 20 min recorded (Forsse and Solbreck 1985). At freeflying speeds of 1.9 to 2 m/s (Gries et al. 1989; Byers 1996a), a maximum range would be 41 to 45.6 km without wind transport. However, wing beat frequency declines with flight duration, which may affect flight range (Byers 2004).

9.3 Deciding whether to accept a host tree or continue to disperse Tradeoffs regarding host plant acceptance by insects have been reviewed by Miller and Strickler (1984). They present a model (their Fig. 9.6.1) by Dethier (1982) where the decision by the insect whether to accept the plant is dependent on external (olfaction, vision, mechanoreception, and gustation) stimulatory and inhibitory inputs balanced against internal excitatory and inhibitory inputs. A graphical, and simplified, model of host acceptance is shown in Fig. 9.9 that is directly applicable to pioneer bark beetles. In this model, as the bark beetle flies around searching for suitable host trees (usually trees already under attack by conspecifics) they use up energy reserves of lipids and probably become increasingly willing to accept substandard hosts (Byers 1999). The beetle may, by chance, encounter several hosts during the dispersal flight that are more or less suitable for reproduction. The beetle will accept a host if the combination of the host suitability and fatigue level of the beetle is above the curve (Fig. 9.9); otherwise the beetle will continue searching for more suitable hosts. The curve is asymptotic to the Y-axis for those beetles that require flight before responding to semiochemicals, whereas the curve would intersect the Yaxis for species that are immediately responsive after emergence. The suitability of the host is determined by the nutritional quality, as well as by the density of established attacks by the same or other species of bark beetle that indicate the potential for damaging competition. At the beginning of a dispersal flight, bark beetles are considered rather unresponsive to pheromone or host volatiles. The theory is that fat reserves are


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higher in freshly emerged beetles so that they have the ability for extended flight and can gain adaptive benefits from dispersal before responding to hosts (Borden et al. 1986; Anderbrant et al. 1985; Gries et al. 1990). Freshly emerged T. lineatum and D. pseudotsugae required 30 or 90 min of flight, respectively, before responding to pheromone from female frass (Bennett and Borden 1971). Atkins (1966) found that female D. pseudotsugae with more than 20 % fat (dry weight) were usually not responsive to the host, while those under 20 % fat were responsive and still could fly. Beetles with less than 10% fat had trouble flying since fat was required as an energy source (Atkins 1969). The fat metabolized by D. pseudotsugae consists mainly of C16 and C18 fatty acids (Thompson and Bennett 1971). Other studies have found that scolytid beetles in the genera Trypodendron, Dendroctonus, Scolytus, and Ips increased their responsiveness or upwind orientation to host and pheromone after continued flight exercise (Borden et al. 1986). However, some bark beetles appear responsive to pheromone upon emergence. Lindelöw and Weslien (1986) found that overwintered I. typographus, collected and marked as they emerged from tents over forest litter, were caught in synthetic pheromone traps within minutes of release. Schlyter and Löfqvist (1986) suggested that preliminary experiments indicated I. typographus became more responsive to pheromone with flight exercise, but further details were not reported. The majority of I. paraconfusus in California responded to aggregation pheromone soon after emergence (Hagen and Atkins 1975). Botterweg (1982) also found that I. typographus can immediately respond to pheromone when beginning dispersal, and this is in accordance with his finding that beetles lost 40%–50% of their fat over the winter. Possibly, second generation beetles in southern Europe would have higher fat content and consequently disperse further.

9.4 Theories of how bark beetles find suitable host trees without aggregation pheromone There are two general theories on how bark beetles find suitable host trees that have not been previously colonized by conspecifics releasing pheromone (Byers 1989a, 1999). The first is that beetles locate such trees by orienting over several meters to volatile chemicals usually released by damaged or diseased trees (called "primary attraction"). It seems that Tomicus piniperda finds hosts by primary attraction as the species is attracted to monoterpenes in the resin (Fig. 9.10). The second theory is that beetles fly about and encounter suitable host trees at random, whereupon they land and test them by short-range olfaction or by taste. The two theories are not mutually exclusive, and one or the other may primarily operate in a particular species. In California, host finding by the important pests D. brevicomis and I. paraconfusus is thought to be a random process. Ponderosa pines that were killed by freezing with dry ice and then screened to prohibit bark beetle attack, did not have higher landing rates for the prevalent D. brevicomis

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Fig. 9.10 Attraction of Tomicus piniperda to monoterpenes released from oleoresin exuding from cut end of a Scotch pine log.

and I. paraconfusus bark beetles (among other species) than did living trees. Landing rates on diseased and healthy trees also were similar. It was estimated that about one D. brevicomis beetle visited each tree in the forest each day (Moeck et al. 1981, Wood 1982). Logs of freshly cut ponderosa pine placed in sticky screen traps did not catch beetles of these species, while at the same time high numbers were attracted to synthetic pheromone or infested logs (Moeck et al. 1981). In addition to I. paraconfusus and D. brevicomis, many species probably visit trees at random, whereupon the tree’s resistance is tested during an attack. For example, Scolytus quadrispinosus was caught equally on traps placed in host shagbark hickory, Carya ovata, and nonhost white oak, Quercus alba (Goeden and Norris 1965). Berryman and Ashraf (1970) found attacks by Scolytus ventralis in the basal section of 74% of grand fir examined, while only 3.5% of these trees were colonized. Most unsuccessful attacks were abandoned before beginning the gallery. The attacks on grand fir appeared random during the early part of the flight period before aggregations resulted. Hynum and Berryman (1980) caught D. ponderosae in traps on 96% of the lodgepole pines (P. contorta) sampled, but only 66% of these pines were killed. Also, they found no differences in landing rates between killed and surviving lodgepole pines or between host and nonhost trees. A direct relationship between the numbers of D. ponderosae caught on unattacked trees and the numbers of trees upon which beetles landed was found in a study of lodgepole pines (Raffa and Berryman 1979). I. grandicollis landed equally on sticky traps on trees judged resistant or susceptible based on crown area (Witanachchi and Morgan 1981). However, Schroeder (1987) found an average of 35 T. piniperda landing on lower vigor Scotch pine, P. sylvestris (as judged by less crown area), than on higher vigor trees (mean of 22 landing per tree). These differences could be due to secondary release of monoterpenes by beetles boring in the low vigor trees that were less able to resist attack.


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There is some evidence that I. typographus is weakly attracted to host volatiles (Lindelöw et al. 1992) or monoterpenes such as α-pinene, but other studies have not observed any attraction to host volatiles or synergism of pheromone and host volatiles (Schlyter et al. 1987a). A computer model by Gries et al. (1989), in which "beetles" must take a series of flights between trees in a grid (each flight to one of eight neighboring trees) and test each tree for suitability, showed that few beetles would find the widely scattered hosts designated as susceptible. Thus, they concluded that a mechanism of long-range primary attraction would be required for maintenance of the population. However, a more recent computer model, in which beetles "fly" more naturally among randomly dispersed susceptible trees indicated that a significant proportion of the population could find the susceptible trees by chance interception of the trunk diameter (Byers 1993a, 1996a). If beetles then test the defenses of the potential host (although this rarely has been observed) then weaker, more susceptible, trees will not exude adequate resin and allow the beetle to produce aggregation pheromone. According to the later model, this will in effect greatly increase the effective attraction "radius" of the tree (or EAR) so that many more in the population can quickly find and colonize these trees (Byers 1993a, 1996a). In addition to the “random landing” and “primary attraction” to host volatile theories, some bark beetles may find weakened and susceptible host trees by orienting to volatiles produced by competing species during colonization. The volatiles can be host compounds that virtually any bark beetle would release upon attack (e.g. monoterpenes) or pheromone components of these other species. For example, D. brevicomis responds to pheromone components of I. paraconfusus in the laboratory (Byers and Wood 1981a); I. typographus responds to exo-brevicomin (from D. micans and Dryocoetes spp., Borden et al. 1987) when combined with its pheromone components (Tommerås et al. 1984); and several sympatric species of Ips in the southeastern United States are cross-attracted to infested pine logs in the field (Birch et al. 1980b). Finally, bark beetles may be aided in locating suitable host trees by avoiding volatiles from (1) hosts fully colonized by conspecifics or competing species and (2) nonhost trees and plants. Avoidance of these types of substrates will be covered in more detail in later sections on discriminating suitability of hosts, avoidance of competition and decaying colonized hosts, and avoidance of nonhosts.

9.5 Strategies of “Pioneer” bark beetles A beetle that lands on a tree and attempts to find a place on the bark to bore is termed a "pioneer" if there are few others present. Pioneers are presumed to encounter significant host resistance and resin when attacking compared to later arrivals ("joiners") when the tree is weakened or has succumbed (Wood 1982; Byers 1995). Only males, in the case of Ips and Pityogenes, or females, in the case of Dendroctonus, initiate the entrance tunnel and can be pioneers, but the joining

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sex in the early stages of colonization must incur some increased risks of resinosis as well. One hypothesis is that since a few pioneers must attack the tree and survive to produce pheromone before numerous others of the population can exploit the resource, pioneers must be the largest and most vigorous beetles of the population. In Fig. 9.11, an alternative theory is presented for the dispersal and host-seeking flight under various conditions and circumstances. An individual should prefer to orient to pheromone and a tree under colonization, but if fat reserves become relatively low during dispersal, then a pioneer strategy becomes advantageous compared to exhaustion in flight (as in Fig. 9.9). As fat reserves become dangerously low, the beetle might attempt to bore into any host tree in the expectation of encountering one of low resistance (Fig. 9.2). Thus, smaller beetles, such as those that suffered severe larval competition and have low fat (Anderbrant et al. 1985; Anderbrant and Schlyter 1989), or those that have used up their fat reserves during a host-seeking flight, regardless of size, are hypothesized to be the pioneers (Byers 1999). If the pioneer beetle lands on a tree of low resistance that cannot produce sufficient resin to repel the beetle, then it has time to feed and excrete pheromone

Fig. 9.11 Conceptual model of dispersal and host-seeking ecology of "aggressive" bark beetles that use aggregation pheromones. Factors such as the beetle's fat reserves, chances of encountering pheromone, and level of competition and host suitability determine whether a beetle joins resident beetles in colonizing a tree or is the first "pioneer" to attack.


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components with the fecal pellets. This then functions as a beacon to individuals of the population in the surrounding area that a weakened host can be exploited as a food and mate resource (Byers 1996a). Aggregation pheromone is an evolutionarily adaptive signal since only trees too weak to vigorously repel beetles with resin will allow beetles to produce pheromone and joining beetles will likely suffer less mortality than as a pioneer. Some species, usually termed less aggressive ones, such as the European pine shoot beetle, Tomicus piniperda, are attracted to monoterpene volatiles produced after injury to the host tree by biotic or abiotic factors that indicate susceptibility (Byers et al. 1985; Byers 1995; 1996a).

9.6 Host semiochemicals attractive to bark beetles Species of bark beetle that regularly attack and kill living trees (termed "aggressive") have been shown nearly always to possess an aggregation pheromone, usually of two or more components, but are weakly, if at all, attracted by host volatiles alone (reviews: Byers 1989a, 1995). However, socalled "secondary" bark beetle species (those that arrive later after the tree has already been killed by the aggressive bark beetles or that feed as saprophytes in decaying trees) may not use an aggregation pheromone, but generally are strongly attracted to either host monoterpenes, ethanol or a combination (Klimetzek et al. 1986; Schroeder and Lindelöw 1989). Host volatiles are attractive to a number of forest scolytids including species in the genera Scolytus, Dendroctonus, Hylurgops, Trypodendron and Tomicus (Byers 2004). Ethanol, probably released by microorganisms in decaying woody tissue and by alcoholic fermentation processes in stressed plants (Kelsey and Joseph 1999), is attractive to a wide variety of species of bark beetles (Byers 2004). Primary alcohols other than ethanol have not been reported as being attractive to scolytids. However, only a few studies have tested methanol (Montgomery and Wargo 1983; Byers 1992a), longer chain alcohols up to hexanol did not attract Scolytids in Sweden when they were known to be flying (Byers 1992a). Electroantennogram (EAG) responses of T. piniperda to a series of straight-chain alcohols indicated that the antennae respond increasingly to longer chain length up to a maximum between pentanol, and heptanol, and then decrease in responsiveness (Lanne et al. 1987). The response spectrum could be due in part to differences in volatility. Thus, although ethanol plays a role in host selection (discussed subsequently), the EAG response for ethanol is lower than for longer-chain alcohols, which are not attractive but rather repellent. 1-Hexanol (from deciduous trees) inhibits response of T. piniperda to attractive monoterpenes (Schlyter et al. 2000). Ethanol and CO2 are the usual end products of sugar fermentation by microorganisms whereas methanol is not, which probably explains the evolution of the use of ethanol by forest insects. Moeck (1970) found methanol to be a minor constituent and ethanol a major constituent of extracts from Douglas-fir sapwood attractive to T. lineatum.

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Various tree monoterpenes (e.g. α-pinene, myrcene, terpinolene, α-pinene, Fig. 9.12) and turpentine are also attractive to a large number of bark beetle species (Byers 2004). Synergism between ethanol and various monoterpenes (or turpentine) is also of widespread occurrence (Chénier and Philogène 1989; Schroeder and Lindelöw 1989; Phillips 1990). These compounds are not only important for primary attraction to plants, but also may play a role in enhancing the bark beetles' response to aggregation pheromone (Byers et al. 1988; Miller and Borden 1990). Host-tree compounds, ethanol and monoterpenes, elicited increased entering rates of bark beetles T. lineatum and P. chalcographus, respectively, into pipe traps baited with aggregation pheromone (Vité and Bakke 1979; Bakke 1983; Byers et al. 1988). ß-Phellandrene (Fig. 9.12) is slightly attractive alone to I. pini and enhances response to pheromone (Miller and Borden 1990), and so the monoterpene might induce entering of holes. In most of the previously discussed studies, the discovery of host compounds attractive to bark beetles has been by the comparative approach (similar species are known to be attracted) or by surmising that identified chemicals in the host would be attractive. Thus, most studies are incomplete because of the possibility that there are still undiscovered chemicals important for attraction to the host. Byers et al. (1985) used the subtractive-combination bioassay and fractionation method (Byers 1992b) to rigorously identify the host volatiles responsible for aggregation of T. piniperda. A combination of (-)-(S)-α-pinene, (+)-(R)-α-pinene, (+)-3-carene, and terpinolene, or each alone, was effective in attracting both

Fig. 9.12 Major monoterpenes of conifers. Note that the enantiomers of α-pinene are identical except that they are non-superimposable (mirror images). Camphene, ß-pinene, 3-carene, ß-phellandrene, and limonene also have two enantiomers, although only (-)-ßpinene and (+)-3-carene are found in trees. Myrcene and terpinolene are achiral.


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sexes (Fig. 9.10 and 9.12). During the isolation study, designed for detection of synergistic pheromone components, no evidence was found for beetle-produced compounds being attractive, in contrast to most bark beetles that aggregate en masse on hosts (Byers 1989a). Byers et al. (1985) quantified the release rates of αpinene, terpinolene, and 3-carene from a freshly cut log of Scotch pine (28 cm x 13 cm diam.) and found them each to be about 15 mg/day. Release of comparable amounts in the field competed favorably with a host log in attracting T. piniperda. They theorized that the beetle’s attraction to monoterpenes functioned in the selection of host species because other common tree species have less monoterpenes. In addition, this attraction to monoterpenes served in the beetle’s recognition of its host's susceptibility since storm-damaged or felled trees have resinous wounds releasing monoterpenes and are less able to resist attack due to the injuries. In the isolation of host volatiles attractive to T. piniperda, a gas-chromatographic adsorbent (Porapak Q), widely used for trapping insect pheromones, was used to collect headspace air from the infested pine logs. Unfortunately, Porapak Q will not retain ethanol molecules due to their small size. Thus ethanol could be a constituent of the attractive host odor. Vité et al. (1986) presented evidence that ethanol enhanced the attraction of T. piniperda to α-pinene and terpinolene (identified above) by about eight-fold, but these results are difficult to confirm since the chemical release rates were not given. They proposed that ethanol would be released from diseased trees and thus indicate their suitability to T. piniperda. Ethanol is attractive when released on healthy trees since T. piniperda were caught in ethanol-baited traps on trees; and these beetles also attacked trees baited with ethanol (Schroeder and Eidmann 1987; Byers 1992a). However, the attraction to ethanol in traps away from trees is weak or negligible, while monoterpenes in these traps are attractive (Schroeder 1988, Schroeder and Lindelöw 1989; Byers 1992a). Ethylene is another chemical that may be released by diseased or dying trees that could be attractive to bark beetles. Campos et al. (1994) found that the olive bark beetle, Phloeotribus scarabaeoides, was attracted to logs of olive that released higher amounts of ethylene. Also, a chemical reaction of 2chloroethylphosphonic acid caused ethylene to be released, which attracted the beetles in the laboratory (Campos and Pena 1995). Trees treated with the chemical released more ethylene and caught more beetles on traps than traps on control trees (Gonzalez and Campos 1995). Treated wood also released more ethylene that resulted in higher densities of attacks by olive bark beetles (Gonzales and Campos 1996). Inoculation of bark beetle-vectored fungi, Ophiostoma minus and O. nigrocarpa, into slash and loblolly pines induced ethylene release (Popp et al. 1995). Ethylene was also released from needles of Monterey pine inoculated with the pitch canker fungal pathogen, Fusarium circinatum. However, the twig-infesting Pityophthorus spp. was not attracted to ethylene, cut branches, or to branches plus ethylene (Bonello et al. 2001), although fungal infected branches were not tested. The authors concluded that

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host discrimination occurs after landing. More research is needed to determine if ethylene plays a role in primary attraction of other bark beetles to weakened hosts.

9.7 Avoidance of competition and unsuitable areas of hosts Bark beetles would gain reproductive benefits if they could select bark areas relatively free of competitors of both their own species and other species utilizing the same phloem resources. It appears that many bark beetle species have evolved olfactory mechanisms to avoid competition for bark resources and one important signal compound is verbenone. For example, verbenone is found in relatively large amounts (mg) in male hindguts of several bark beetles of North America, D. frontalis, D. brevicomis, D. ponderosae, and D. pseudotsugae (Borden 1997) but in low amounts (ng) in T. piniperda (Lanne et al. 1987), or essentially absent in I. paraconfusus, I. typographus, and P. chalcographus (Byers 1983a; Birgersson et al. 1984, 1990). Verbenone (Fig. 9.4) inhibits the attraction of these beetles to their respective aggregation pheromones (Borden 1997; Byers 2004). Exposure of male and female D. brevicomis to (+)- and (-)- enantiomers of αpinene for several hours caused them to produce large amounts of (+)- and (-)trans-verbenol in their hindguts (Fig. 9.4, Byers 1983b). However, the biosynthesis of verbenone in these beetles was not affected by exposure to αpinene enantiomers, even though verbenone is structurally similar to α-pinene (Fig. 9.12) and is found in males landing on trees (Renwick and Vité 1970; Byers et al. 1984). The (-)-enantiomer of trans-verbenol (Fig. 9.4) inhibits female D. brevicomis from entering holes and may serve as a signal to arriving females that they should avoid areas colonized by conspecifics (Byers 1983b). Both verbenone and trans-verbenol are produced by D. brevicomis beetles in the greatest amounts early in colonization so it was suggested that they play a role in reducing intraspecific competition (Byers et al. 1984), as well as interspecific competition with I. paraconfusus (Byers and Wood 1980). However, verbenone (and possibly trans-verbenol) are also produced increasingly in ageing logs infested by bark beetles (Birgersson and Bergström 1989; Byers et al. 1989b). A common bacterium, Bacillus cereus, also isolated from I. paraconfusus, can make cis- and trans-verbenol from α-pinene (Brand et al. 1975). Several yeasts from I. typographus can interconvert the verbenols, and when grown in a phloem medium they produced the oxygenated monoterpenes α-terpineol, borneol, myrtenol, terpenene-4-ol and trans-pinocarveol, compounds also shown to be released increasingly from bark beetle holes with age of attack (Birgersson and Bergström 1989). A mycangial fungus grown in culture media converted alcohol products of αpinene to verbenone, the end product (Brand et al. 1976). These microorganisms are introduced by bark beetles during colonization and after buildup may release verbenone, thus signaling to flying beetles that remaining in these bark substrates would entail competition with established bark beetle colonies. Recently,


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verbenone was found in twigs of eastern cottonwood, Populus deltoids (personal comm. from Carlos Flechtmann, in Huber et al. 1999). Myrcene and α-pinene from host trees can be used as precursors of pheromones and allomones in some species of conifer-feeding bark beetles. However, recent research has shown that the majority of ipsdienol and ipsenol in Ips bark beetles results from de novo biosynthesis from simple acetate and mevalonate precursors (Seybold and Tittiger 2003). The later steps in the isoprenoid pathway lead to geranyl diphosphate and myrcene before being stereoslectively, and species-specifically, converted to various enantiomers of myrcene-like alcohols (e.g., ipsdienol, ipsenol, E-myrcenol, amitinol) that are pheromone components (Seybold et al. 2000, Seybold and Tittiger 2003). D. brevicomis, I. paraconfusus and I. pini occur sympatrically in California and Oregon and compete for ponderosa pine bark. Myrcene vapors can be converted only by male D. brevicomis to (+)-ipsdienol (Fig. 9.4), also a pheromone component of its competitor I. paraconfusus (Byers 1982). (+)-Ipsdienol inhibits response of both D. brevicomis and I. pini to their synthetic aggregation pheromones (Birch et al. 1980a; Lanier et al. 1980; Byers 1982). In Europe, ipsdienol, a pheromone component of I. duplicatus could act as an allomone to inhibit response of I. typographus (Byers et al. 1990b; Schlyter et al. 1992). Mated males of I. typographus produce small amounts of ipsdienol and ipsenol during colonization that might function in avoiding competition (Birgersson et al. 1984; Birgersson and Leufvén 1988; Birgersson et al. 1988). Although, ipsdienol was previously thought to be an aggregation pheromone component of I. typographus (Bakke et al. 1977), other studies have shown the compound to be either inactive or inhibit attraction of these beetles to cis-verbenol and methyl butenol, the most potent components (Schlyter et al. 1987b, c). Also, ipsdienol was not found in single males in a nuptial chamber, while it was detected in small amounts in later phases of attack when males had been joined by one or more females (Birgersson et al. 1988). P. chalcographus of Europe (Fig. 9.13) produces a two- component aggregation pheromone consisting of chalcogran and methyl decadienoate (Francke et al. 1977; Byers et al. 1988, 1990a). Both chalcogran and methyl decadienoate (Fig. 9.4) cause I. typographus to avoid landing on traps releasing their aggregation pheromone components (cis-verbenol and methyl butenol) (Byers 1993b). However, P. chalcographus attraction to its pheromone was not inhibited by pheromone components of I. typographus even though the latter, and larger, beetle wins in competitive situations when both species attack simultaneously (Byers, unpublished). However, if P. chalcographus precedes I. typographus on the host by just a few days then the former species will win in competitive situations (Byers, unpublished). Verbenone is increasingly released as colonized areas of I. typographus age (Birgersson and Bergström 1989) and the compound is inhibitory to P. chalcographus (Byers 1993b), as well as to I. typographus (Schlyter et al. 1989).

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Fig. 9.13 spruce.

Pityogenes chalcographus searching for a boring site on the bark of Norway

9.8 Avoidance of decaying or fully colonized hosts As mentioned above, some microorganisms isolated from bark beetles or their gallery walls, may convert α-pinene to cis- and trans-verbenol, or trans-verbenol to verbenone. It was proposed that this process may account for termination of attack. Verbenone is increasingly released from ageing logs of spruce and pine colonized by bark beetles (I. typographus, Birgersson and Bergström 1989; T. piniperda, Byers et al. 1989b), possibly due to the activity of microorganisms. Byers (1989a, b) speculated that if verbenone is a consistent signal of microbial activity in decaying hosts, then bark beetle species may have evolved an avoidance to this compound (a kairomone) in order to avoid unsuitable hosts. The bark beetle then could have evolved to produce verbenone as a pheromone that reduced intraspecific competition, since the avoidance response was already existent. Other bark beetle species might then evolve to avoid species that produced verbenone (as an allomone), and so avoid interspecific competition. Sympatric species on the same host might coevolve responses to, and/or production of, verbenone since the chemical could serve as a signal for several types of beneficial information (kairomone, pheromone, and allomone). Verbenone was reported to inhibit pheromone responses of over 10 species of bark beetle (Borden 1997). Verbenone also deterred attack on conifer logs or in conifer stands treated with a grid of release devices affixed to trees or with aerially-applied verbenone-impregnated plastic pellets/flakes (Schlyter et al. 1988; Gillette et al 2006, 2009; Borden et al. 2007). However, verbenone does not always inhibit bark beetles. For example, H. palliatus feeds in unhealthy or dying Scotch pines that release verbenone (Byers et al. 1989b), and the beetle's attraction to ethanol was not inhibited by verbenone (Byers 1992a). Verbenone efficacy has been inconsistent at stand levels of several conifer species probably due to breakdown in ultraviolet radiation to inactive chrysanthenone (Francke et al 1995; Borden 1997). Angiosperm trees in a state of decay may not release verbenone since they probably do not have α-pinene, thus T. domesticum could evolve to avoid degrading nonhost pines by avoiding verbenone (Byers 1992a).


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Another bark beetle, P. bidentatus, attacks diseased limbs of Scotch pine and is not affected by release rates of verbenone, which inhibit more aggressive bark beetles. This is probably because verbenone is expected to be present from the diseased host limbs (Byers et al. 2000). In the case of conifer bark beetles, Verbenone is increasingly implicated as a general sign of host unsuitability in conifer-killing bark beetles (due to microbial decay or competition with bark beetles). Therefore, it is paradoxical that conifers have not evolved the capacity to convert α-pinene, which they have in abundance, to verbenone in order to repel aggressive bark beetles. Ethanol, also, sometimes reduces response to attractive baits. Klimetzek et al. (1986) tested different release rates of ethanol (24 to 125 mg/day) with an unreported release rate of α-pinene and terpinolene and found that the higher releases of ethanol inhibited attraction of T. piniperda. However, a control with either ethanol alone or terpenes alone was not reported. Schroeder (1988) increased the release of ethanol in five dosages over an even wider range from 0 to 50 g/day in combination with a 240-mg/day α-pinene release. In this case, the attraction of T. piniperda declined linearly with the logarithm of ethanol release, which is in conflict with the theory of Vité et al. (1986) that ethanol was synergistic with monoterpenes. Schroeder and Lindelöw (1989) provided the first evidence that could integrate the disparate results. They found that high release rates of α-pinene were most attractive to beetles and that ethanol releases alone from 0 to 3 g/day were barely attractive. At a low release rate of α-pinene (2.4 or 22 mg/day), and thus low attraction, lower release rates of ethanol from 0 to 3 g/day had a synergistic effect when combined with α-pinene in attracting beetles (Schroeder and Lindelöw 1989). Their results are supported by Byers (1992a); i.e., a weak enhancement of attraction by ethanol at low release rates when blended with three host monoterpenes, but no observable effect of ethanol on the greater attraction to higher release rates of monoterpenes. Ethanol released at even higher rates, 120 mg/day (Klimetzek et al. 1986) or 50 g/day (Schroeder 1988), inhibited the response of T. piniperda to monoterpenes. Therefore, the beetle could find diseased, but physically uninjured, trees by a weak response to a synergism between low monoterpene release rates and moderate ethanol rates- the hypothesis of Vité et al. (1986). Beetles would occasionally penetrate these trees, and if low in resistance would permit continued feeding. Resinosis and monoterpene release from the entrance holes would elicit increased numbers of beetles joining in a mass attack. Injured trees with wound oleoresin, and trees under attack with "pitch tubes", would have a higher monoterpene release and attract the greatest numbers of beetles, according to Byers et al. (1985). Trees with high ethanol release rates would indicate a tree in advanced decay and unsuitable for reproduction, and thus to be avoided, as theorized by Klimetzek et al. (1986). High monoterpene releases from trees (freshly wounded and not dead) would not naturally coincide with high ethanol release rates (presumably during decay after death). In addition, other

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compounds such as verbenone from decaying hosts would inhibit response to monoterpenes from unsuitable hosts (discussed in the next part). These studies emphasize the need for releasing semiochemicals at known rates during tests in the field. In addition, measurements of the natural release rates of ethanol and monoterpenes from various host and nonhost substrates are necessary for further understanding of bark beetle chemical ecology.

9.9 Avoidance of nonhosts Plant suitability in insects was reviewed by Scriber (1984). A plant's suitability to bark beetles varies with its nutritional quality and composition of deterrents and toxins. Nonhost trees are probably less nutritional to a particular beetle than its hosts. The beetle in most cases would not be expected to be able to detoxify some of the toxins in nonhosts (which are avoided or not usually encountered) that may have evolved for use against herbivorous insects. A beetle would save much time and energy if it could discriminate between the host and the nonhost and determine the suitability of the host by olfactory means from a distance without the need to land. Sometimes host and nonhost trees are adjacent and the beetle could land by mistake on the nonhost; however, shortrange olfactory cues might indicate the inappropriateness of the nonhost bark substrate (Byers et al. 1998, 2000). If the beetle still could not decide, boring a short distance into the nonhost might reveal the lack of feeding stimulants or the presence of deterrents causing the beetle to leave (Elkinton and Wood 1980; Byers et al. 2000). According to studies discussed previously, bark beetles find their host tree by attraction to host volatiles (or after random landing and probing), as well as by avoiding chemicals from colonized hosts or decaying hosts. However, it is becoming increasingly apparent that many beetles avoid nonhost trees due to specific odors. It is inherently more difficult to isolate repellents and inhibitors used in avoidance behavior than to isolate attractants since tests of avoidance require one to first isolate the attractive host odors and then present these with and without the possibly inhibitory nonhost odors. Several studies indicate that at least some species of bark beetle avoid nonhost volatiles during their search for host trees (reviewed in Borden 1997; Schlyter and Birgersson 1999, Byers et al. 2004; Zhang et al. 2007b). Common green leaf volatiles (GLVs) are emitted at high levels from leaves of angiosperm trees that may indicate non-host habitats (Huber 2001; Zhang 2001). Specific bark volatiles signalling non-host angiosperm species, like the trans-conophthorin, 3-octanol, 1-octen-3-ol, and some benzenoids released from the bark of Betula spp., Populus spp. and Acer spp. (Fig. 9.14; Borden et al. 1998; Schlyter and Birgersson 1999; Huber 2001; Zhang 2001). The attraction of both T. piniperda and H. palliatus to ethanol (1–6 g/day) was reduced by odors from cut logs of nonhost trees, birch, Betula pendula, and aspen, Populus tremula (Schroeder 1992). In future experiments, host logs (or


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Fig. 9.14 Major nonhost volatile compounds from bark or leaves of angiosperm trees proven to be behaviourally or antennally active in conifer-inhabiting scolytids (Zhang and Schlyter, 2004).

monoterpenes and ethanol) should be tested instead of ethanol alone to simulate the host tree. Dickens et al. (1992) reduced the attraction response of D. frontalis, I. grandicollis and I. avulsus to aggregation pheromone by releasing the greenleaf volatiles, 1-hexanol and hexanal. T. domesticum colonizes wood of deciduous trees (e.g. Fagus sylvatica, Quercus spp. Betula spp.) and is known to be attracted to ethanol (Magema et al. 1982; Paiva and Kiesel 1985). Monoterpenes of Scotch pine and verbenone (from decaying conifers) reduced response of this species to ethanol (Byers 1992a) and would provide a mechanism for avoiding nonhosts and unsuitable colonization areas. This also is valid for the hardwood-breeding species, Anisandrus dispar (Schroeder and Lindelöw 1989). The spruce bark beetles Ips typographus and Pityogenes chalcographus were shown to avoid volatiles of nonhost birch trees (both from bark and leaves, Fig. 9.15), which suggests the possibility that beetles may not enter areas of primarily birch (Byers et al. 1998). However, it is more certain that the beetle would leave a birch tree after landing due to a relatively high concentration of repellent nonhost volatiles at the surface of the bark. In addition to the above evidence, numerous studies hae shown that aggregation responses to semiochemicals by conifer-infesting bark beetles in several genera are reduced by volatiles from nonhost angiosperm trees (e.g. Betula, Populus, Acer) (Dickens et al. 1992; Schroeder 1992; Schlyter et al. 1995; Wilson et al. 1996; Guerrero et al. 1997; Borden et al. 1997, 1998; Deglow and Borden 1998; Byers et al. 1998, 2000; Poland et al. 1998; Zhang et al. 1999a, b,

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Fig. 9.15 Pityogenes chalcographus were induced to land on birch trees by baits of aggregation pheromone but the individuals did not stay more than about 2 minutes due to odors from the bark (probably 1-hexanol, trans-conophthorin, and other unidentified compounds, Byers et al. 1998).

2000, 2001; Huber et al. 1999, 2000, 2001; Huber and Borden 2001a, b; Poland and Haack 2000; Schlyter et al. 2000; Zhang 2003; Zhang and Schlyter 2003). These studies have found that some of the most important nonhost angiosperm compounds are (Z)-3-hexen-1-ol, (E)-2-hexen-1-ol, and 1-hexanol mostly from leaves, as well as trans-conophthorin from bark (Fig. 9.14). Volatiles from leaves or bark of nonhosts birch (Betula pendula) and Norway spruce (Picea abies) also dramatically reduced the attraction of the bark beetle, Pityogenes bidentatus, to their aggregation pheromone components (cis-verbenol and grandisol) in the field (see image of the authors collecting odors of nonhost trees, Fig. 9.16). Surprisingly, odors from either the needles or bark of the host Scotch pine, Pinus sylvestris, similarly inhibited attraction. Monoterpenes of pine and spruce (α-pinene, β-pinene, terpinolene, and 3-carene), as well as ethanol, chalcogran and some nonhost green leaf alcohols [(Z)-3-hexen-1-ol, (E)-2-hexen1-ol, and 1-hexanol], also reduced catches. Collections of volatiles from the field-

Fig. 9.16 Authors (left to right: John Byers and Qing-He Zhang) collecting odors on Porapak Q adsorbent from thermal oven bags containing bark of Scots pine, Norway spruce, and various hardwood species in southern Sweden in April 2001.


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tested plant tissues indicated they released monoterpenes in amounts similar to the synthetics that inhibited responses. The varied plant and insect sources of these inhibitory compounds indicate that P. bidentatus bark beetles have evolved several strategies to increase their fitness by avoiding nonhost and unsuitable host trees in a complex olfactory landscape. Recent electrophysiological and behavioural studies clearly indicate that conifer-inhabiting bark beetles are not only able to recognize but also to avoid non-host angiosperm habitats or trees by olfactory means (Zhang and Schlyter 2004). Antennal responses to non-host leaf and bark volatiles have been found by using the coupled gas chromatography – electroantennographic detection (GCEAD) techniques in over 20 species of bark beetles (Pureswaran et al. 2004; Zhang and Schlyter 2004; Shepherd et al. 2007; Zhang et al. 2007a; Zhang et al. 2008). The latest study by Andersson et al. (2009) using Single-Cell Recording (SCR) technique found that 25% of the strongly responding neurons on Ips typographus antennae were tuned to compounds typical of angiosperm non-host plants. These antennally active NHVs (Non-Host Volatiles such as GLVs and angiosperm bark volatiles; Fig. 9.14) in individuals or various combinations also showed strong disruption to the pheromone/kairomone-positive responses of many conifer-inhibiting bark beetles (Zhang and Schlyter 2004; Fettig et al. 2005). The most disruptive individual NHV components and blends vary among different scolytid species. The response diversity to NHVs by conifer-inhabiting bark beetle species might reflect differences in the odor characteristics of their particular host and non-host habitats and ecosystems (Poland et al. 1998; Zhang and Schlyter 2004). Combined NHV signals in blends showed both redundancy and synergism in their inhibitory effects (Zhang and Schlyter; 2003). A recent SCR study on I. typographus also showed that part of these behavioral redundancy among NHVs probably is integrated exclusively at the peripheral level, whereas the synergism appears to be integrated in the CNS (Andersson et al. 2009). The coexistence of redundancy and synergism in negative NHV signals may indicate different functional levels (non-host habitats, species, and unsuitable hosts) in the host selection process (Zhang and Schlyter; 2003). In addition to volatile repellents or inhibitors, a bark beetle could avoid a potential host or nonhost by lack of feeding stimulants or presence of deterrents in the bark or phloem (Byers and Wood 1981b; Elkinton et al. 1981; Raffa and Berryman 1982; Byers et al. 1998, 2000; Byers 2004).

9.10

Control of bark beetles with semiochemicals

Attractive pheromones have been used in the field to disrupt mate finding in insects (references in Byers 2007). In most cases, relatively large quantities of pheromone (consisting of several pheromone components) are more or less evenly distributed throughout the field to adapt (overload) sensory receptors or habituate behavioral response (`confusion') or to exhaust the individuals in orientation attempts (i.e. “wild-goose chases”). The best successes so far have

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involved straight-chain olefinic acetates, alcohols, and aldehydes of moths (Byers 2002, 2007). The effective attraction radius (EAR) has been used in mating disruption and mass trapping models for moths (Byers 2007, 2008, 2009) and bark beetles (Byers 1996a). Bark beetles that colonize forest trees may present problems for disruption techniques for several reasons, one is that their pheromone components, usually oxygenated monoterpenes, are more volatile than moth straight-chain hydrocarbons (Byers 1989a). More important perhaps is that compared to moths even larger quantities are expected to be required for disruption of bark beetles since the latter individuals generally release higher rates (ng to mg/h) of pheromone components than moths (pg to ng/h) (Schlyter et al. 1987a; Birgersson and Bergström 1989; Byers et al. 1990a, b; Ramaswamy and Cardé 1984; Du et al. 1987). Furthermore, even higher quantities of synthetic pheromone are required to compete with pest bark beetles that typically release semiochemicals in large aggregations on their host tree as compared to individual female moths. Possibly because of these reasons, as well as the fact that both sexes are attracted by pheromone, several attempts to control bark beetles have used the mass-trapping method. This method employs traps, either sticky-screen or cylinder with holes/barrier type, baited with synthetic pheromone components. Traps releasing pheromone components have been used in control programs to lure other pest insects such as moths to their death (Bakke 1989; Haniotakis et al. 1991; Sternlicht et al. 1990). The first major attempt to control bark beetle populations using pheromonebaited traps was done in 1970 in California (Wood 1980). Large (1 x 2 m) sticky screens baited with exo-brevicomin and frontalin, pheromone components of the western pine beetle, Dendroctonus brevicomis, plus the host monoterpene, myrcene, were placed in ponderosa pine forests at Bass Lake, California. In four plots of 1.3 km2 each, 66 pheromone traps were deployed in a grid of about 161 m spacing. Over a million beetles were caught and the test appeared to be successful since the number of trees killed by the beetle declined to 10% the pre-treatment level for several years (Wood 1980). Norway and Sweden have extensive conifer forests, and in the 1970's a major outbreak of the European spruce engraver, I. typographus, devastated many areas (Austarå et al. 1984). Since the pheromone of this beetle had recently been identified as a mixture of 2-methyl-3-buten-2-ol and (1S,4S,5S)-cis-verbenol (Bakke et al. 1977), an extensive mass-trapping control program was initiated in 1979 and may have led to the decline of outbreaks after 1980 (Bakke 1989; Vité 1989). Several other European studies have reported successful control of bark beetles with the intensive use of pheromone-baited traps (Vrkoc 1989; Richter 1991; Jakus 1998). Recently, Schlyter et al. (2001) reported a successful case of pheromone mass trapping of the bark beetle Ips duplicatus in a forest island, analyzed by 20-year time-series data in Inner Mongolia, China. Previous theoretical attempts at determining the effectiveness of pheromone mass trapping have used population dynamic models (Fisher et al. 1985; Barclay 1988; Barclay and Li 1991). These models are mathematically complex and make several assumptions about beetle survival and mating rates, as well as attraction


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rates to pheromone traps, which limits their application. There have been no models where ìnsects' are moved in `real' time and space in relation to traps of specific dimensions and positions, although two-dimensional models of ‘correlated random walks’ are probably close to reality (Byers 1993a, 1996a, b, 1999, 2000, 2001, 2007). In many past control programs that used pheromone trapping, there has been the problem of finding control areas to determine whether the treatment has been effective. However, several monitor traps placed inside the control area (or even the control traps themselves) will indicate the population density and the progress of the control program. If no more insects are being caught, then obviously the control is a success, unless the flight period is over. This can be determined by monitoring traps placed in untreated areas, some distance away, but still within the same general biotope and climatic regime. Usually only one beetle or pair of bark beetles begin attack of a tree and at this time pheromone release is relatively low compared to a few days later when thousands of beetles participate in the mass attack. Thus, it seems advantageous to initiate mass trapping before beetles swarm in the spring and have time to build aggregations that can compete with traps for attraction of dispersing beetles. The population levels need only be reduced below the thresholds required to kill trees. Synthetic NHVs have been tested in several tree protection experiments in North America and Europe. Wilson et al. (1996) first demonstrated that a blend of the GLVs: (Z)-3-hexenol and (E)-2-hexenol, released at ca. 8 mg/day, reduced attacks of Dendroctonus ponderosae (Hopkins) on attractant-baited trees (Wilson et al. 1996). Benzyl alcohol, a volatile found in non-host callus, at high doses (700 mg/day per log) reduced attacks on host logs by Tomicus destruens (Guerrero et al. 1997). Field studies in North America also showed that combination of NHVs and verbenone significantly reduced attack densities and tree mortalities caused by D. ponderosae (Huber and Borden 2001; Borden et al. 2003), D. brevicomis LeConte (Fettig et al. 2008; Fettig et al. 2009), D. valens LeConte (Fettig et al. 2008) and Ips perturbatus (Eichhoff) (Graves et al. 2008) at individual tree level. Jactel et al. (2001) reported that trans-conophthorin combined with a blend of NHValcohols achieved 89% reduction of I. sexdentatus (Boerner) attacks on un-baited pine logs, and 62% on pheromone-baited logs in France (Jactel et al. 2001). A strong area effect, protecting forest edge zones with treated trees, was observed in a spruce tree protection experiment on I. typographus in the Czech Republic, 2000–2001 (Jakus et al. 2003). In this experiment with a mixture of NHVs plus verbenone, there were significantly less overall attack rates in the zones with treated trees than in the untreated control zones (Fig. 9.17) (Jakus et al. 2003). Further investigations on economically optimal blends for operational use are still in progress in both North America and Europe. These findings on non-host angiosperm volatiles lead to a new hypothesis (Zhang 2001) (‘semiochemical-diversity hypothesis’) that because mixed forests have greater semiochemical diversitythan pure host stands, they would disturb

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Fig. 9.17 Full scale spruce forest edge protection experiments against Ips typographus with a mixture of non-host volatiles (NHVs) plus verbenone in Šumava, Czech Republic, 2000–2001. Synthetic blends of NHVs such as 1-hexanol, C8-alcohols, and trans-conophthorin, were applied at 2 m and 6 m above the ground on every second tree within treatment sections, each consisting of 10 trees. Control sections were of the same size, without any treatment, established only for observations. A) 6 Adjacent sections of each type, in total 60 trees in 2000. B) 13 Sections of each type, paired but not adjacent, in total 130 trees in 2001. In both the cases the observed difference in attack rates was highly significant (χ2 tests with Yates’ correction for continuity both P < 1% **) (Jakus et al. 2003).

olfaction-guided host selection and would reduce the likelihood of outbreaks of conifer-infesting bark beetles. A meta-analyses of 118 studies by (Jactel and Brockerhoff 2007) recently showed the generality of a lower risk of herbivore outbreaks in mixed forest. The semiodiversity has been shown by a manipulation experiment to be one causative factor of lower damage by a moth folivore and a bark beetle (Jactel & Schlyter in prep.). This ‘semiochemical-diversity hypothesis’ would provide new support to the general ‘stability-diversity hypothesis’ of population dynamics (Zhang and Schlyter 2004).

9.11

Ecological aspects and concluding remarks

Bark beetles are a keystone species, meaning that they are evolutionarily and historically a dominant component of the natural forest ecosystem. Without bark beetles, many thousands of microbial, nematode, mite, and insect species would


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become extinct since they rely on bark beetles to create a habitat for them (Byers 2004). The aggressive interactions between bark beetles and trees also must affect population genetics and evolution. Trees and parts of trees (shoots, cones, limbs) are killed when eaten by bark beetles or by fungi introduced during feeding so there is certainly a severe selection pressure on trees to evolve resistance mechanisms, which they have, but bark beetles have also evolved counterresistance mechanisms (Byers 1995). This so-called evolutionary arms race continues, especially with man's activities introducing new species of tree and bark beetle. In spite of a general resistance having evolved in conifers, older, weak and unhealthy trees are removed by bark beetles, and microbial diseases, which greatly influences the age and species structure of the forest. The main reason bark beetles have been studied so extensively is that they are perceived as pests that damage forests. Certainly at outbreak levels, bark beetles kill vast areas of conifer forests, which drastically affect the ecology and species composition for a considerable time. In forest plantations used only for the production of fiber, bark beetles are indeed pests. In forests with residential tracts, bark beetles are a threat to the desired stability of old-growth stands. In wilderness areas and recreational forests, bark beetles may be tolerated as part of the natural ecosystem. In fact, bark beetles are a keystone species that naturally fluctuates in abundance as the forest ages and succession processes occur. Many forests are designated as multi-use, meaning that they are for recreation and for timber production. Obviously some uses preclude other uses, or are at least in conflict. There often must be a compromise between producing the most fiber per unit area (short term interest) and the maintenance of natural forest biodiversity (long term interest). Over the long term (longer than a human’s lifetime), most forest ecosystems require disturbances that remove old trees and open the land to a succession of plant and animal species guilds. Either bark beetles or fire, or both, are well known to carry out this natural long-term cycling function. Man, however, usually does not appreciate these perturbative cycles that progress over many decades. The discussion about the role of bark beetles, fire, and overuse of forests by mankind will continue for many years to come, but research in all these areas will help to understand how to better utilize and enjoy nature.

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CHAPTER 10

Infochemical-tritrophic Interactions of Soybean Aphids-host Plants-natural Enemies and Their Practical Applications in Pest Management Junwei J. Zhu

Abstract The soybean aphid, Aphis glycines Matsumura, is a newly invasive insect species that seriously threatens U.S. soybean production. This aphid pest has kept haunting many soybean growers by developing large colonies on soybeans in North America since 2000. Since its first appearance in Wisconsin, it has spread to over half of US states and southern provinces in Canada. The heavy infestation of this pest whittles soybean growers’ profits and causes hundreds of million dollar losses. The present chapter will mainly describe efforts in studying aphid chemical ecology and sensory physiology for understanding how male aphids find their mates and host plants. It will also cover research efforts to understand host plant associated volatiles being used as cues for overwintering host plant location. In addition, findings on how soybean plant defensive system works against aphid infestation, as well as how those induced plant volatiles are used by aphid’s natural enemies for prey location will be presented. Finally, the use the basic understandings for developing useful tools for soybean aphid practical control will be discussed. Keywords infochemicals, tritrophic interactions，host plant, natural enemy, mating disruption, integrated pest management

10.1

General introduction

Soybean aphids, Aphis glycines Matsumura, are newly-invasive aphid pests that have caused significant economic losses to U.S. soybean production. This aphid pest tends to develop large colonies on soybeans, Glycine max, in North America (Ragsdale et al. 2004). Since their first being found in Wisconsin soybean fields, Junwei J. Zhu Agroecosystem Management Research Unit, USDA-ARS, Room 5, Entomology Hall, University of Nebraska, East Campus, Lincoln, NE 68583-0938, USA E-mail: [email protected]


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they have spread to over 20 US states and southern provinces in Canada (Soybean aphid watch 2003). The infestation of this pest whittles soybean growers’ profits and causes hundreds of million dollar losses (CNN News, Nov. 25, 2003; Chicago Tribune Business News, Oct.11, 2003, and data from Midwest Soybean Aphid Workshop, Feb. 5, 2004). The soybean aphid has a complex life cycle with more than 15 generations per season living on the secondary host, G. max. During the autumn, the soybean aphid producing winged females (gynoparae) that fly from soybean fields searching for their primary host plant, the common buckthorn, Rhamnus cathartica or R. alnifolia in the U.S. (Voegtlin et al. 2004). Once on buckthorns, gynoparae produce pheromone-emitting wingless female offspring (oviparae). Winged males are attracted to oviparae via a specific sex pheromone blend produced from glands on the hind legs of female aphids (Zhu et al. 2006). After mating, oviparae lay eggs that overwinter on the buckthorn.

10.2

Sex pheromone of aphids

Oviparae of soybean aphids produce both (1R,4aS, 7S,7aR)-nepetalactol and (4aS,7S,7aR)-nepetalactone, which are the two most common aphid pheromone compounds identified from a number of other aphid species (Goldansaz et al. 2004, Boo et al. 2000; Pickett et al. 1992; Dawson et al. 1990). The pheromone blend identified at a specific ratio of 35∶65 of the two compounds, (1R,4aS,7S,7aR)-nepetalactol and (4aS,7S,7aR)-nepetalactone, from A. glycines is highly attractive to conspecific males and gynoparae (Fig. 10.1, Zhu et al. 2006). This also indicates that gynoparae are capable of using pheromone compounds as cues for overwintering host plant location. Further evidence has shown that leaves of buckthorns release less volatiles after the appearance of gynoparae and pheromone-producing females (Zhu, unpublished).

10.3

Pheromone applications in aphid control

The use of synthetic sex pheromone to disrupt mating behavior has become a widely accepted and increasingly used integrated pest management (IPM) tool for suppressing populations of several key lepidopteran pests of agricultural crops and tree fruits around the world (Baker et al. 1997; Sanders 1997). Recent studies have shown that males of several aphid species can be selectively attracted to traps releasing synthetic aphid pheromones and gynoparous female aphids to host plant associated volatiles and sex pheromones at a relatively long distance (Campbell et al. 1990; Hardie et al. 1992, 1996; Boo et al. 2000; Lösel et al. 1996a, b). Some of the males have been further observed to orient towards pheromone traps against surprisingly strong winds. The sensory adaptation of soybean aphid antennal responses to the two pheromone compounds have been first demonstrated by Zhu et al. (2006) (Fig. 10.2). These recent findings are

Infochemical-tritrophic Interactions and Their Practical Applications in Pest Management 193

Fig. 10.1 Total number of male and gynoparous soybean aphids caught in traps with different combinations of identified sex pheromone compounds in 2001. Means with different letters on top of the bars indicate significant differences. (N = 10, for gynoparae, F = 75.42; df = 3,20; P < 0.001, for males, F = 99.04; df = 3,36; P < 0.001).

Fig. 10.2 Comparisons of absolute EAG responses of male soybean aphids preexposed with higher dosages of aphid pheromones to those without pre-exposure. Means with different letters on top of the bars indicate significant differences (Student T-test, N = 12, for nepetalactone, t = 5.08, P < 0.001; for nepetalactol, t = 3.54, P < 0.005, for blank, t = 1.18, P> 0.05).

encouraging for the potential deployment of mating disruption using sex pheromones in soybean aphid control.

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Another IPM component, mass trapping of insect pests using sex pheromones or host plant volatiles (as synergists), has also shown renewed promise as a population management tool (Kobayashi et al. 1981; Smit et al. 2001) for both moth and beetle pests. Mass trapping of male soybean aphids, as they leave soybean fields to locate females on the winter host, using inexpensive trap designs placed at the edges of soybean fields may be a feasible approach. Likewise, mass trapping of gynoparae leaving soybean fields may reduce population densities of overwintering aphids. The field trapping results shown in Fig. 10.1 indicate gynoparous soybean aphids respond to their female pheromones as well. Disruption of the ability of these gynoparae to fly to winter host plants by diverting them with the same sex pheromone dispensers that disrupt mate finding may further reduce the production of sexually reproductive female soybean aphids, thereby adding to the reduction in the overwintering population caused by mating disruption alone. Sex pheromones have become very important tools for monitoring agricultural pest populations, and provide critical information for growers to take actions on pest outbreaks. Currently, soybean aphid population density monitoring in the field relies heavily on traditional field scouting, which is laborious and timeconsuming. Morphological studies indicate that antennae of spring and summer winged soybean aphids (alatae) soybean aphids contain placoid sensilla (secondary rhinaria) with olfactory receptor neurons that respond to the identified sex pheromone components (Du et al. 1995; Zhu and Park, 2005; Zhu et al. 2006). Field trapping tests have also shown that these alatae are caught in traps baited with 10 mg of the soybean aphid pheromone blend. By comparison of total number of these alatae caught from pheromone traps and mean number of soybean aphids per plant (based on scouting 30 plants randomly in each field), the peak catch is approximately 10–14 days earlier than the highest abundance of soybean aphids in the field (Fig. 10.3). These results suggest that it may be possible to use the trap catch to establish an early warning system for monitoring soybean aphid outbreaks.

10.4

Semiochemical-based biological control

The use of predatory insects and parasitoids, as biological control agents to suppress population of pest species on either economically important crops or in home gardens, is well recognized by the general public and by biological control practitioners (Obrycki and Kring 1998). There have been significant successes in using coccinellids and chrysopids to suppress whitefly, aphid, mealybug, scale, and mite populations (Gerling 1990; Frazer 1988; New 1991). The use of natural enemies as biological control agents for soybean aphids has great potential to suppress soybean aphid populations (Rutledge et al. 2004). However, the biggest concern for the success of soybean aphid biocontrol is how to recruit predaceous insects into natural or damaged soybean fields and synchronize their presence with the targeted aphid pest, thereby increasing their predatory efficacy. Several


Fig. 10.3 Comparisons of total number of alatae soybean aphids caught in pheromone traps and mean number of soybean aphids per plant in two soybean fields (Iowa, USA, 2005).

predatory insects and parasitoids have been demonstrated to use semiochemicals associated with host or the host plant habitat to locate their prey (Barbosa and LeTourneau 1988; Turling et al. 2002; Vet and Dicke 1992). Zhu and Park (2005) studied the soybean plant response to aphid infestation, and showed that the plant induced defensive compound, methyl salicylate, and 2-phenylethanol (also associated with soybean plant) are attractive to several beneficial insects of soybean aphids (Fig. 10.4). Van der Werf et al. (2000) and James (2003) have reported on the use of artificial lures to attract and retain predators in alfalfa fields and hop yards. Field trials to determine the impact of predators on soybean aphid dynamics have

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Fig. 10.4 Mean number of predatory insects (both sexes) caught in traps baited with 100 mg of methyl salicylate and 2-phenylethanol, and the control from an Iowa soybean field in 2003. Columns with no letters in common in four different categories are significantly different (n = 10, ANOVA followed by FPLSD test, P < 0.05).

revealed that early season predation on soybean aphids can cause significant reductions of their later immigration to soybeans (Rutledge et al. 2004; Fox et al. 2004). The manipulation of these predatory insects to be synchronized with the soybean aphid appearance can be achieved by applying beneficial insect attractant lures during the earlier season in aphid-infested soybean field. Field applications using controlled release packets (developed by MSTRS Technologies, Inc.)1) containing these two attractant compounds (Predalure with 2phenylethanol and Me-SA lure with methyl salicylate) have shown significant increases in number of beneficial insects and the suppression of soybean aphid populations in the treated fields (Fig. 10.5). Meanwhile significant more pods are produced from soybean. Plants in plots treated with the attractant lures or combined with sugar water, compared to those from plots treated with the control, or sugar water alone (Fig. 10.6, upper). An increase of 50% on pod weight has also been shown in treated plots (Fig. 10.6, lower). A further field trial with the application of beneficial insect lures in aphid-infested soybean fields showed that soybean yields from the treated field was 1,424 kg versus 1,284 kg from the control field (two fields at a size of ~ 30,000 m2, each, and the treated field was deployed with 30 lures of each, and the lures were replaced monthly during the 5month experimental period).

1) This article reports the results of research only. Mention of a proprietary product does not constitute an endorsement or a recommendation for its use by USDA


Fig. 10.5 Effects of applications of beneficial insect attractants on soybean aphid and their beneficial insect populations in the soybean field. Lure: Predalure and Me-SA lure; Lure + Sugar water: lures with the spray of sucrose application (250 grams of sucrose in one liter of water); Sugar water: sucrose application only; Control: blank. (ANOVA followed by a type III test, F = 23.3; df = 3, 184; P < 0.01, SAS Institute 1999).

10.5

Summary

In summary, the present chapter presents a unique case of exploring novel integrated aphid management strategies using infochemical tritrophic interactions among the aphid, host plants, and natural enemies. Such practices can also benefit other agricultural pest management, particularly for organic crop growers where application of chemical pesticides is not an option. However, some of these practices may be laborious and costly than the traditional approach, and need additional developments to reduce costs before the fully deployment of this new strategy.

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Fig. 10.6 Comparisons of number of pods and pod weights from soybean plants collected from the treated fields and control fields (ANOVA followed by a type III test, F> 4.3; df = 3, 16; P < 0.05, SAS Institute 1999). Acknowledgements This project was funded by National Science Foundation-SBIR program, Iowa Department of Natural Resources and The Leopold Center for Sustainable Agricultures to J. Zhu. I will also thank J. Jones, J. Heath, T. Liu and B. Ellingson for their technical supports during the course of research. Suggestions and comments from Drs. J. Obrycki and B. Wienhold on an earlier version of the manuscript are especially appreciated.

References Baker T C, Mafra-Neto A, Dittl T, Rice M E. A novel controlled-release device for disrupting sex pheromone communication in moths, // P. Witzgall and H. Arn. Technology transfer in mating disruption. Montpellier: IOBC wprs Bulletin, 1997, 20: 141–149. Barbosa P, LeTourneau D K. Novel Aspects of Insect-Plant Interactions. Wiley, New York, 1999.

Infochemical-tritrophic Interactions and Their Practical Applications in Pest Management 199 Boo K S, Choi M Y, Chung I B, et al. Sex pheromone of the peach aphid, Tuberocephalus momonis, and optimal blends for trapping males and females in the field. J. Chem. Ecol., 2000, 26: 601– 609. Campbell C A M, Dawson G W, Griffiths D C, et al. Sex attractant pheromone of damson-hop aphid Phorodon humuli (Homoptera, aphididae). J. Chem.Ecol., 1990, 16: 3455–3465. Chicago Tribune – Knight Ridder/Tribune Business News. 2003. Soybean aphids drain production of Midwestern cash crops. Oct. 11. CNN News (Associated Press). 2003. Aphids whittling soybean farmers’ profits. Nov. 25. Dawson G W, Griffiths D C, Merritt L A, et al. Aphid semiochemicals-a review, and recent advances on the sex pheromone. J. Chem. Ecol., 1990, 16: 3019–30. Du Y J, Yan F S, Han X L, Zhang G X. Olfaction in host selection of the soybean aphid Aphis Glycine. Acata. Entomol. Sinica., 1995, 37: 385–391. Fox T B, Landis D A, Cardoso F F, Difonzo C D. Predators suppress Aphis glycines Matsumura population growth in soybean. Biol. Control, 2004, 33: 608–618. Frazer B D. Coccinellidae. // AK Minks, P Harrewijn. Aphids—Their Biology, Natural Enemies and Control, Vol. B, New York, Amsterdam: Elsevier, 1988: 231–247. Gerling D. Natural enemies of whiteflies: predators and parasitoids. // D. Gerling. Whiteflies: Their Bionomics, Pest Status and Management. Andover: Intercept Ltd., 1990: 147–85. Goldansaz S H, Dewhirst S, Birkett M A, et al. Identification of two sex pheromone components of the potato aphid, Macrosiphum euphorbiae (Thomas). J. Chem. Ecol., 2004, 30: 819–834. Hardie J, Nottingham S F, Dawson G W, et al. Attraction of field-flying aphid males to synthetic sex pheromone. Chemoecology, 1992, 3: 113–117. Hardie J, Storer R J, Cook F J, et al. Sex pheromone and visual trap interactions in mate location strategies and aggregation by host-alternating aphids in the field. Physiol. Entomol., 1996, 21: 97–106. James D G. Synthetic herbivore-induced plant volatiles as field attractants for beneficial insects. Environ. Entomol., 2003, 32: 977–982. Kobayashi M, Wada T, Inoue H. A comparison of communication disruption technique and masstrapping technique for controlling moths using sex pheromone of Spodoptera litura (F.) (Lepidoptera: Noctuidae). Proc. Japan/USA Symp. On IPM., 1981, p32–40. Lösel P M, Lindemann M, Scherkenbeck J, et al. The potential of semiochemicals for control of Phorodon humuli (Homoptera: Aphididae). Pestic. Sci., 1996a, 48: 293–303. Lösel P M, Lindemann M, Scherkenbeck J B, et al. Effect of primary-host kairomones on the attractiveness of the hop-aphid sex pheromone to Phorodon humuli. Entomol. Exp. Applic., 1996b, 80: 79–82. Midwest States Soybean Aphid Management Workshop, Feb. 5, 2004. http://www.ipm.uiuc.edu/ fieldcrops /insects/soybean_aphids/Workshop/ index.html New TR. 1991. Insects as Predators. Kensington, Aust.: NSW Univ. Press. 178 pp. Obrycki J J, Kring T J. Predaceous Coccinellidae in biological control. Annu. Rev. Entomol., 1998, 43: 295–321. Pickett J A, Wadhams L J, Woodcock C M. The chemical ecology of aphids. Annu. Rev. Entomol., 1992, 37: 67–90. Ragsdale D W, Voegtlin D J, O'Neil R J. Soybean aphid biology in North America. Ann. Entomol. Soc. Am., 2004, 97: 204–208.

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Rutledge C E, O’Neil R J, Fox T B, Landis D A. Soybean aphid predators and their use in IPM. Ann. Entomol. Soc. Am., 2004, 97: 240–248. Turlings T C J, Gouinguené S, Degen T, et al. The chemical ecology of plant-caterpillar-parasitoid interactions. // Tscharntke and Hawkins. Multitrophic Level Interactions. Cambridge: Cambridge Univ. Press, 2002: 148–173. SAS Institute Inc., SAS OnlineDoc®, Version 8, Cary, NC: SAS Institute Inc., 1999. Smit N E J M, Downham M C A, Laboke P O, et al. Mass-trapping male Cylas spp. With sex pheromones: a potential IPM component in sweet potato production in Uganda. Croprot., 2001, 20: 643–651. Snaders C J. Mechanisms of mating disruption in moths. // Cardé R T, Minks A K. Insect Pheromone Research New Direction. New York: Chapman & Hall, 1997: 333–346. Soybean Aphid Watch 2003. http://www.ipmcenters.org/northcentral/ saphid/ Van der Werf W, Evans E W, Powell J. Measuring and modeling the dispersal of Coccinella septempunctata (Coleoptera: Coccinellidae) in alfalfa fields. Eur. J. Entomol., 2000, 97: 487– 493. Vet L E M, Dicke M. Ecology of infochemical use by natural enemies in tritrophic context. Annu. Rev. Entomol., 1992, 37: 141–172. Voegtlin D J, O’Neil R J, Graves W R. Test of suitability of overwintering host of Aphis glycines: Identification of a new host association with Rhamnus alnifolia. Ann. Entomol. Soc. Am., 2004, 97: 233–234. Zhu J, Park K C. Methyl salicylate, a soybean aphid-induced plant volatile attractive to the predator Coccinella septempunctata. J. Chem Ecol., 2005, 31: 1733–1746. Zhu J, Zhang A, Park K C, et al. Sex pheromone of the soybean aphid, Aphis glycines Matsumura, and its potential use in semiochemical-based control. Environ. Entomol., 2006, 35: 249–257.

CHAPTER 11

The Responses of Insects to Global Warming Kun Guo, Osbert Jianxin Sun and Le Kang

Abstract Climate warming has been largely concerned with its impacts on natural ecosystems. Both flora and fauna are shown to respond to climate warming in a variety of processes across multiple organizational scales — from simple behaviors of individuals to complex structural and functional dynamics of communities. As poikilothermal animals, insects are particularly sensitive to climate warming. Here we examined current evidence on responses of insects in development, survival, reproduction, phenology, distribution, abundance, population dynamics, and community structure to global climate change. A case study was reviewed in more depth and showed that warming advanced the timing for egg hatching and eclosion in three grasshopper species, and that grasshopper diapause and increased precipitation could offset the effect of warming on egg development. Based on the experimental evidence, we draw conclusion that insects can, to some extent, adapt to global warming and escape from potential risks. Keywords climate warming, insect, phenology, distribution, ecosystem

Natural ecosystems have been experiencing influences of a undebated climate change (King 2004). A growing body of evidence shows that climate warming has affected each component of natural ecosystems (Parmesan 2006; Parmesan and Yohe 2003; Root et al. 2003) and brought strong impacts on terrestrial

Kun Guo, Le Kang State Key Laboratory of Integrated Management of Pest Insects and Rodents, Institute of Zoology, Chinese Academy of Sciences, Beijing, 100101, China E-mail: [email protected] Osbert Jianxin Sun Key Laboratory for Silviculture and Conservation of Ministry of Education, College of Forest Science, Beijing Forestry University, Beijing, 100083, China


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ecosystems across a wide range of processes—from the behavior of individual organisms to biodiversity and the function of entire ecosystems (Parmesan 2006; Parmesan and Yohe 2003; Root et al. 2003; Walther et al. 2002). Climate warming has been found to directly affect many metabolic and developmental processes in plants and animals (Hufnagel and Gaal 2005; Hughes 2000; Kiritani 2006; Musolin 2007), with known impacts including advancement in the occurrence of first leaf (Menzel 2000; Menzel et al. 2001) and flowering (Cayan et al. 2001; Fitter et al. 1995; Luo et al. 2007; Menzel and Dose 2005) and timing of harvests (Menzel 2005) in plants, earlier calling and breeding of amphibian (Beebee 1995; Gibbs and Breisch 2001), and laying and migration of birds (Both et al. 2004; Gaston et al. 2005). The current trend of global climate change is expected to cause shifts of species distribution to higher latitude (Smith 1994; Thomas and Lennon 1999; van Herk et al. 2002; White et al. 1999) or altitude (Grabherr et al. 1994; Luckman and Kavanagh 2000; Pauli et al. 1996; Pounds et al. 1999) as well as extended growing season (Menzel, 2000; Menzel and Fabian 1999; White et al. 1999). More examples of the impacts of warming include changes in population dynamics (Cullen et al. 2001; Sillett et al. 2000), community composition (Harte and Shaw 1995; Sagarin et al. 1999) and carboncycle of ecosystems (Cox et al. 2000; Luo 2007), and reduction in species richness (Barbraud and Weimerskirch 2001; Bradshaw and Holzapfel 2001; Gross 2005). Insects, with more than 1.5 million species identified in the group, are known as poikilothermal animals and therefore can be more sensitive to temperature changes than most of vertebrates (Roy and Sparks 2000). The earlier researches on temperature responses of insects were mostly carried out in incubators with quite a few species (Andrews and Donoghue 2004; Chladny and Whitman 1998; Lamb and Gerber 1985; Petersen et al. 2000). Realization of the limitation in indoor experimental conditions later leads to utilization of field manipulative studies to examine the responses of insects to climate warming. Studies with Odonata, Lepidoptera, Hemiptera, and Diptera etc. (Hickling et al. 2005; Hodkinson and Bird 2006; Kiritani 2006; Parmesan et al. 1999), have generally demonstrated that climate warming induces alteration in insect physiology, behavior, abundance, phenology, distribution, and voltinism, and drives some local populations to extinction and causes changes in community structure (Forister and Shapiro 2003; Roth and Masters 2000). There were studies showing that warming increased the overwinter survival (Kiritani 2006), accelerated growth, and enhanced fecundity (Miles et al. 1997) of insects. Many insect species, especially those in the temperate zone, have been found to shift their distribution to higher latitude in concert with climate warming (Battisti et al. 2005; Haeger 1999; Hassall et al. 2007; Hickling et al. 2005; Jordano et al. 1991; Paulson 2001). For example, 63% of the butterfly species investigated in Europe expanded their distribution northward (Parmesan et al. 1999); the northern boundary of Ampittia dioscorides in America extended 420km further north over the past four decades (Crozier 2003, 2004). As a result of global

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temperature increase, some species have shifted their distribution to higher altitude (Battisti et al. 2005). Apart from changes in distributional range, many insects have responded to climate warming by displaying advanced phenophases. In England, it was found that warmer spring advanced the appearance and flight of butterflies and dragonfly (Forister and Shapiro 2003; Roy and Sparks 2000); higher winter temperature caused by heating pipes was shown to advance the phenophase in species of Auchenorrhyncha (Mastes et al. 1998). Global warming has also been attributed to habitat loss or alteration of habitat conditions that leads to population decline or extinction. It has been found that in just 30 years, 1.3°C rise in temperature reduced habitat area by one-third for 16 butterfly species in central Spain (Wilson et al. 2005). Overall, studies to date shows that global warming has different effect on different species, which may result in changes in trophic structures of ecosystems (Andrew and Hughes 2005; Roth and Masters 2000), potentially leading to mismatches of insects with their host plants. In the following sections, we present brief reviews of experimental evidence on some of the specific responses of insects to climate warming and of processes that are susceptible to warming. Growth, development and survival Different species has different temperature requirements for growth, development, and survival. The best fitness of a given insect species is often achievable only under its optimum growth temperature. The effects of warming on growth, development, and survival have been examined through manipulation experiments. In Wales of UK, 3°C elevation of temperature in greenhouse environment increased egg development and reduced their hatching time in Operophtera brumata (Buse and Good 1996). In Ainsdale, 2.5°C elevation in temperature accelerated the larvae development of Craspedolepta nebulosa and C. subpunctata (Hodkinson and Bird 2006). Over the last 40 years in Japan, the winter mortality of adults of Nezara viridula and Halyomorpha halys was reduced by 13.5–16.5% with the mean surface temperature rose by 1.0°C (Kiritani 2006). Reproduction and behavior In North Pennines, UK, warming created by cloches enhanced the fecundity of Strophingia ericae (Miles et al. 1997). In Northern Italy, due to increased temperature of summer, more adults of Arocatus melanocephalus were found to move to construction of cities for cooling (Zandigiacomo 2003). Distribution Insects would die of stress if they are exposed to extreme low temperature for a short time period or generally low temperature for a prolonged time. So, the winter extreme cold temperature is the key factor that determine the northern limit of insects (Crozier 2003, 2004). With the climate warming, in a sample of 35 nonmigratory European butterflies, 63% have ranges that have shifted to the north by 35 – 240 km during last century, and only 3% have shifted to the south (Parmesan et al. 1999). In the most-extreme cases, the southern edge contracted concurrently

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with northern edge expansion. For example, the sooty copper (Heodes tityrus) was common in the Montseny region of central Catalonia in the 1920s, but modern sightings only occurred in the Pyrenees, 50 km to the North. Symmetrically, H. tityrus entered Estonia for the first time in 1998, by 1999 had established several successful breeding populations, and by 2006 had reached the Baltic Sea (Parmesan et al. 1999). Within 35 years, the northern range limit of Ampittia dioscorides expanded 420 km northward, from California to Washington (Crozier 2003, 2004). In the hottest year (1998), the northern range limit expanded northward by 75km. The desert orange tip (Colotis evagore), which historically was confined to northern Africa, has established resident populations in Spain while maintaining the same ecological niche (Jordano et al. 1991). Over the past 32 years, the pine processionary moth (Thaumetopoea pityocampa) has expanded 87 km at its northern range boundary in France and 110–230 m at its upper altitudinal boundary in Italy (Battisti et al. 2005). Between 1960 – 1995, among all 37 species of resident Odonates in the United Kingdom, 23 of the 24 temperate species had expanded their northern range limit, with mean northward shift of 88 km (Hickling et al. 2005). In tropic area, five dragonfly species expanded their distribution area from Cuba and the Bahamas to Florida (Paulson 2001). Similarly, the African plain tiger butterfly (Danaus chrysippus) established its first population in southern Spain in 1980 and by the 1990s had established multiple, large metapopulations (Haeger 1999). Phenology Like all the organisms, insects cannot complete their life history cycles until experienceing enough effective accumulated temperature. Elevation in temperature reduces the time for heat accumulation and thereby advances the phenology of insects. With increasing spring temperature, 26 out of 35 butterfly species studied advanced their first and peak appearance by 2–10 days in England (Roy and Sparks 2000). A study in Spain indicated that, with 1–1.5°C elevation in March temperature, all the 17 butterfly species studied advanced their first appearance time (Stefanescu et al. 2003). In California, 70% of the 23 butterfly species studied advanced average first spring flight by 24 days (Forister and Shapiro 2003). During 1960–2004, the flight time of dragonfly in UK advanced by 3.08 days with the temperature increase of 1°C. There appears a tendency in Odonate species which display egg nondiapause to advance their phenology. This is well supported by a manipulation experiment in Inner Mongolia of China (Guo et al. 2009). Another manipulation experiment with heating pipe as warming facilities showed that the phenology of Auchenorrhyncha was significantly advanced with a winter temperature increase of 3°C (Masters et al. 1998). Species abundance Temperature variation has differential impacts on diverse insect species. A study in UK showed, with 3°C elevation of winter temperature, the abundance of nymph and adult of Heteropteran decreased (Roth and Masters 2000); whilst in central section of Japan, increased summer temperature raised the abundance of Leptocorisa chinensis, hence enhanced the harm to rice (Yokosuka


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2001). In Hungary, a research on Hempitera community indicated that increased temperature enhanced abundance of dominant species (Hufnagel and Gaal 2005). Voltinism As increased temperature can advance development and phenology, it is possible that climate warming may cause more generation of insects to occur within one year. Basing on the relationship of 28 Hemiptera species and temperature, Yamamura and Kiritani (1998) predicted that temperature increase by each 2–3°C would lead to occurrence of one more generation each year. In Rocky mountains of USA, with temperature increase, Dendroctonus ponderosae completes one generation in one year instead of two years, accompanying increased abundance (Logan et al. 2003). Synchrony of different trophic levels Having different life history traits, different species can have differential responses to climate change even in the same environments (Parmesan and Yohe 2003). The disruption of synchrony caused by climate change between insects and their host plants and between parasitoids and their host insects has been found to affect individuals more than that of climate change itself (Harrington et al. 1999; Parmesan 2006; Visser and Both 2005). Feeny (1970) found that because egg hatching of Opheroptera brumata couldn’t match the sprout of their host plant oak under conditions of climate warming, 90 percent larvae died of starvation. In another case, the larvae of Euphydryas editha began to develop when their host plant Plantago erecta experienced senescence under increased temperature conditions, and this mismatch led to high motality of Euphydryas editha (Singer 1972). However, there are exceptions to the above patterns. For example, in UK, Anthocharis cardaminers has shown phenology shift synchronous with that of their host plants (Sparks and Yates 1997). Population extinction Due to climate change, many species are losing their habitat. For those species that fail to expand new habitat or are not able to expand because of geographical barriers, they have high risk of extinction because of declining habitat. In the past 30 years, 16 butterfly species in Spain have lost one thirds of their habitats (Wilson et al. 2005). Southern boundary of 25% of European butterfly species shift northward by 35–50 km over 30–70 years. For some species whose northern boundary didn’t expand, they are likely to become extinct because of contraction of distribution (Parmesan et al. 1999). Community structure Increased temperature has different impacts on different trophic levels or different species of the same trophic level, resulting in alternation of community structure. A study on Hempitera species of UK showed that increased winter temperature affected the abundance and phenology of different species differentially, and thereby changed the community structure (Roth and Masters 2000). Distribution shift of insects can also result in changes in the local community structure (Andrew and Hughes 2005).

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A case study with three co-existing grasshopper species of Inner Mongolia grasslands Inner Mongolia grasslands host some 172 grasshopper species out of about 30000 in total worldwide (Li et al. 2007). Nearly all those Inner Mongolian grasshopper species have only one generation per annum, and occur in different seasons and form sequential development cohorts (Kang et al. 2007). They survive the winter as eggs in soil, and are categorized into early-, mid- and lateseason species based on timing of occurrence (Kang and Chen 1994). Complex life histories have evolved in insects to cope with seasonal fluctuations in climate, and to permit maximum exploitation of warmer seasons while cooler seasons are survived by more resistant forms such as diapause stage (Butterfield and Coulson 1997). The grasshoppers lay both diapause and nondiapause eggs, most of which can safely survive during the winter season, and the diapause eggs enter diapause in embryo stage 19 (Hao and Kang 2004a, b,c; Zhao et al. 2005 ). There are significant niche differentiation of the grasshopper species in temporal, spatial and food dimensions in Inner Mongolia (Kang and Chen 1994). Guo et al. (2009) conducted a comprehensive field manipulation experiment in the period 2006–2007 in Inner Mongolia, China, aiming to investigate the effects of artificially increased temperature and precipitation on three contrasting grasshopper species, which included the early-season species Dasyhippus barbipes (F.-W.), the mid-season species Oedaleus asiaticus (B.-Bienko), and the late-season species Chorthippus fallax (Zub.). In the study, infrared heaters were used for warming the ground surface by 1–2°C above the ambient condition and periodic irrigations were applied to simulate a 50% increase in annual precipitation. The use of suspension infrared heats mimics the conditions of climate warming by enhancing downward infrared radiation (Harte et al. 1995; Wan et al. 2002). Guo et al. (2009) found that warming advanced the timing for egg hatching and grasshopper eclosion in each of the three species. However, grasshopper diapause and increased precipitation appeared to offset the effect of warming on egg development (Fig. 11.1). Hatching and development were more strongly affected by warming in the mid-season O. asiaticus and the late-season C. fallax relative to the early-season D. barbipes (Guo et al. 2009). Warming by ~1.5 °C advanced the occurrence of the mid-season O. asiaticus by an average of 4.96 days; whilst warming and increased precipitation interactively affected the occurrence of the late-season C. fallax, which advanced by 5.53 days (Guo et al. 2009). The study of Guo et al (2009) suggest that most grasshopper species in the Inner Mongolian grassland are likely to extend their distribution northward with climate change and, because of the differential responses to warming, that different grasshopper species may aggregate toward the middle period of the growing season (Fig. 11.2), potentially increasing inter-specific competition and grazing pressure on grasslands. Concluding remark By reviewing experimental evidence in literature, we have demonstrated that


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Fig. 11.1 Developmental divergence of non- and diapause eggs in warming and ambient conditions (after Guo et al. 2009).

Fig. 11.2 Sketch map of phenology shift of three grasshopper species induced by warming.

climate warming imposes substantial effects on insects. The effects occur upon different organizational levels, i.e. from individuals to populations and communities through ecosystems. However, we have only examined a fraction of insects and ecological and physiological processes that are sensitive to climate warming. In fact, there are many groups of insects not being represented in the climate change research and the processes studied are highly selective. From the information available in literature, it appears that most of the researches to date have been focused on shift of phenology and distribution as means of insect responses to climate warming. Further studies on responses to climate warming

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by insects with contrasting life history and ecological traits are still needed to better understand the life of insects, and their interactions with other components of ecosystems, in future world of changing climate.

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CHAPTER 12

Biology and Reproductive Strategies in the Subterranean Termites (Isoptera: Rhinotermitidae) Xingping Hu

Abstract The subterranean termites in the Rhinotermitidae family have the most complex and plastic life style and fascinating reproductive systems. Some of these termites are the most economically and ecologically important species in many parts of the world. This review presents updates on their biological features, life cycle pathways and breeding structures from the aspects of organismal and molecular levels. Keywords Subterranean termites, biology, management, neotenic differentiation, reproductive mechanisms

12.1

Introduction

As with all other social insects, termites are characterized by the differentiation of colony members into either reproductive or sterile individuals. Termites are typically divided into the higher termites (Termitidea), containing 80% of all species, and the lower termites, represented by the remaining six families. Of the lower termites, the Rhinotermitidae family is the most derived with complex life style and reproductive systems (Inward et al. 2007) and contains the most economically and ecologically important termites in many parts of the world, especially temperate and subtropical regions (Eggleton 2000). Thus, understanding their basic biological features of the life history, reproductive systems can provide insights into the eu-social evolution and management. Although subterranean termites have attracted increasing attention from entomologists working at basic to molecular aspects, the diverse of these insects’ mating systems and reproductive strategies and how they are maintained remain mysterious. A significant amount of theoretical and empirical publications have Xingping Hu Department of Entomology and Plant Pathology, Auburn University Auburn, Alabama, 36849, USA E-mail: [email protected]


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Xingping Hu

documented termite life cycle and termite breeding systems including inbreeding, outbreeding, sexual and asexual reproductions. Colony social structures range from simple families consisting of a single queen and a single genetically unrelated king, and their progeny to a variety of more complex mixing families which have been explained by several mechanisms that are broadly divided into the those driven by alate reproductive strategies (multiple unrelated alate reproductives or adoption) and those driven by worker strategies (colony fusion or replacement reproductives) (Myle 1999; Deheer and Vargo 2004). Termites are unique among social insects because they undergo incomplete metamorphosis and display a remarkably complex and diversified caste polyphenism (Snyder 1926). The study of post-embryonic development in termites is extremely difficult and complex because of their hiding living style, plastic biology and behavior, a lengthy developmental time, polymorphism, different developmental pathways, and convertible reproductive mechanisms. Given the inherent difficulties in studying such cryptic, eusocial organisms, it is not surprising that the literature on their biology has failed to reach a consensus. Moreover, the ambiguity of colony architecture and the cryptic nest and feeding habits hinder our understanding of their evolutionary ecology of population structure (Thorne et al. 1999). In this chapter, I summarize current information from results of field and laboratory observations and experiments from the aspects of organism to molecular levels, to shed light on understanding of the biology and reproductive systems in Rhinotermitidae termites.

12.2

Termite castes

A termite colony is typically composed of individuals that have differentiated into different castes, typically including reproductives, workers, soldiers, nymphs, larvae, and eggs. As with other types of polyphenisms, termite castes differentiation is believed the result of the interaction of endogenouse (a complex genetic inheritance pattern) and exogenouse (environmental) factors during critical stages in development (Nijhout 1994, Hayashi et al 2007), whereas in early days it had been suggested that caste development is determined within the embryo (Snyder 1925). The terminology regarding caste is complicated and somewhat controversial (Myle 1999, Thorne et al. 1999, Forschler and Jenkins 1999) because there appears to be a considerable variation in the development pathways (Roisin 2000). The classical paradigm of life history is that colony reproduction occurs with the dispersal flights of fertile adults. These fertile adults are alates that are fully pigmented and sclerotized, and have 2 pairs of wings and a pair of compound eyes. Such alates are also named imagos, incapable of further molts. Alates are the only caste that leave parent colony by taking a dispersal flight and expose themselves to air. The alates breaks off their wings as soon as they alight, and the resultant dealate runs in apparently random directions until it encounters a potential partner to form a tandem running pair. Homoseusal tandems often occur

Biology and Reproductive Strategies in the Subterranean Termites

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when two consexual dealates encounter one another. The paired dealates locate a suitable nest site where moisture and cellulose materials are sufficient, mate and initiate a new colony. The female is the queen and the male is the king and they are both the primary reproductives. The primary reproductives are initially able to feed, but the ability is reduced after their first brood of offspring grows into workers, who then take over the duty of feeding the primary reproductives (Krishna 1989). The role of the primary productives is to produce offspring by laying eggs. Eggs hatch into larvae, namely the 1st and 2nd instars. Their mandibles are not strong enough to eat so they are initially tendered by primary reproductives and later by workers (Grasse 1949). At the 2nd molt, the larvae differentiate into either wingless worker pathway or wingpad nymph pathway (Fig. 12.5) (Hayashi et al. 2007). Wingless worker pathway: Workers go through varying numbers of instars depending on the species and environmental conditions. Buchli (1958) observed seven worker instars, excluding the first two larval instars, after which the workers continue to molt without change in size (so named stationary molt). Within a colony, the vast majority of the individuals (both male and female) enter the functionally sterile, irreversibly wingless worker pathway. Workers are the working force of the colony: foraging for food, feeding members of other castes, building nest and tunnels, tending the nursery and grooming nestmates. They can remain true workers, or at certain stages become presoldiers that molt into soldiers, or transform into a reproductive form named ergatoids. True workers are the later instars that are non-reproductive and have diverged irreversibly from the nymphal pathway. Workers are blind; their compound eyes and ocelli are either highly reduced or absent. Presoldiers (also referred to white soldiers or pseudosoliders) have soldier-like head and morphology but their heads are unpigmented and unscelrotized. Presoldiers molt into soliders. Soliders have a heavily sclerotized head and well developed mandibles that are modified as a weapon for defense. Both presoldiers and soldiers are not capable of feeding but rely on liquid diet through trophallaxis from workers. Presoldiers and soldiers are both blind and often significantly outnumbered by workers. Once the soldiers are formed, they are unable to regress to another caste. Ergatoids arewingless workerlike reproductives that are also referred to as apterous secondary reproductives, apterous neotenic reproductives, or the 3rd form reproductive. They show no wing development and retain many juvenile characteristics. All the castes are comprised of both sexes but only ergatoids are able to reproduce. Wingpad nymph pathway: Nymphs also go through varying numbers of stages depending on species and environmental conditions. They have a similar role as the workers. They are individuals with wingpads and develop either into alates with wings and eyes (imagoes) that disperse and become primary colony founders, or they develop into nymphoids which are also referred to as brachypterous secondary reproductives, brachypterous neotenic reproductives, or the 2nd form reproductive. Nymphoids have rudimentary wing buds, neither eyes

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nor ocelli, and cannot fly. They have no eyes either. Their role is to supplement or replace the primary reproductives within the colony. Traditionally the term neotenic has been used in reference to non-alate-derived reproductives, so do the terms secondary, supplementary, or replacement reproductives. At the loss of either or both primary reproductives, whether through death or colony fragmentation, one or more nymphs and/or workers are stimulated to transform into neotenic substitutes.

12.3

Mechanisms of neotenic differentiation

Although an inhibitory pheromone from primary reproductives is likely involved, but the genetic and physiological mechanisms influencing neotenic differentiation remain one of the great mysteries of termite biology (Myles 1999). Scharf et al (2005) identified genes that were differentially expressed in nymphs and reproductives. A gene coding for a hexamerin protein showed highest expression in nymphs and neotenic reproductives and is thus assumed to play a role in reproductive differentiation, possibly by modulating the availability of JH in the hemolymph. Following the general model of insect development, low JH titers during critical JH-sensitive periods in development almost certainly regulate differentiation into reproductives in subterranean termites, although the number and timing of these critical periods are likely to differ between development into primary reproductives and development into neotenics (Nijhout 1994).

12.4

Life cycle pathways

A variety of possible life cycles have been proposed to depict the developmental bifurcation pathways of Reticulitermes species. There are some disagreements over the number of molts that occur for development into different stages and the stages that differentiate into different castes. This is likely due to the difficulty in identifying individuals at different developmental stages. Four of the well cited diagrams are presented here. The first proposed life cycle (Fig. 12.1) by Snyder (1915) based on the work of Grassi and Sandias (1893). Snyder identified only one undifferentiated larval instar, with the 2nd instar differentiating into different pathways. Larvae with large heads develop along the “neutral worker-soldier pathway” while those with small heads develop along the “sexual line”, which then splits into neotenic and imagal pathways. The second life cycle (Fig. 12.2) was proposed by Grasse (1949). He confused the sexual and worker neutral pathways by showing a cross-over from sexual line (individuals developing from small-headed larvae) and the worker line (individuals developing from large-headed larvae). In his theory, ergatoid develops from the nymph line. Currently, this theory is considered incorrect. However, should not simply be disregarded as it may still play a role in improving our knowledge of rhinotermitid life cycle.


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Fig. 12.1 Life cycle of Reticulitermes species (Snyder 1915). L: larva; N: nymph; LB = larva with large head; EL = emergency reserve reproductive; W: worker; WL: worker larva; NL: nymph with long wing buds; NS: nymph with short wing buds; SL: emergency larval substitute; SN: emergency nymphal substitute; BN: brachypterous neotenic; PS: presoldier; S: soldier.

Fig. 12.2 Life cycle of Reticulitermes species (Grasse 1949). L: larva; N: nymph; LB = larva with large head; W: worker; S: soldier; LS: larva with small head; LE = emergency reserve reproductive; E: ergatoid; NL: nymph with long wing buds; NS: nymph with short wing buds; BN: brachypterous neotenic.

The third life cycle (Fig. 12.3) is from Vieau (1991), who believed ergatoid develops from the 6th or 7th instars and the soldiers from the 6th instar of the worker line. At the 6th instar of the sexual nymph line, nymphs with long wingpads develop into alates and those with short wingpads into brachypterous neotenic reproductives. The most recent theory (Fig. 12.4) is proposed by Hayashi et al. (2007).

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Fig. 12.3 Life cycle of Reticulitermes species (Vieau 1994). L: larva; N: nymph; W: worker; E: ergatoid; PS: presoldier; S: soldier; NL: nymph with long wing buds; NS: nymph with short wing buds; BN: brachypterous neotenic; P: pseudergate.

Fig. 12.4 Caste developmental pathways of Reticulitermes spp ((Hayashi et al. 2007). Termites are diploid and hemimetabolous; larvae of each sex follow either the worker pathway, in which individuals remain irreversibly wingless, or the nymphal pathway, leading to the alate form. Arrows indicate molts; dotted arrows indicate occasional molts to neotenic nymphoids or ergatoids, which may reproduce in the absence of the queen and/or king. Workers undergo stationary molts after w5. Soldiers are derived from w4 and w5. Ergatoids arise from all worker stages after w1.


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Utilizing biological and genetic analyses, they supported the hypothesis that a bifurcation occurs early after the 2nd instar in the development. The vast majority of larvae enter the functionally sterile and irreversibly wingless worker pathway. Alternatively some larvae develop wing buds and enter the nymph pathway, which leads to the alate caste. In the absence of the primary queen and/or king, some individuals develop into neotenics from the nymph and/or the worker pathways. In the nymph line, nymphoids may arise from the 4 – 6 nymph stages. In the worker line, ergatoids may arise from all worker stages after the 1st worker stage. Workers undergo stationary molt after 5th worker stages. Solders are derived from the 4th and 5th worker stages.

12.5 Sexual colony foundation and reproductive mechanisms Termite colonies are normally founded by a single pair of winged reproductives. They also form through fusing or splitting of existing colonies, or though parthenogenesis. The unique characteristic in termite colony foundation, compared with ants is that it usually involves both sexes (Wilson 1971). 12.5.1

Outbreeding

Colonies founded by primary reproductives (one queen and one king alate) are referred to as incipient colonies. In theory, the primary pair is unrelated, meaning that the female alate always searches and mates with a male alate from different colonies, This is known as outbreeding. Termites have evolved to swarm in great masses from many different colonies, a possibility to provide arenas for outbreeding. The males of Kalotermes flavocollis Kalotermitidae invariably leave the nest 2–3 hr after the females, a phenomenon as anything but a device to promote outbreeding. Recent studies of gene flow suggest that alate dispersal distances can be sufficient to promote outbreeding in some subterranean termite species (DeHeer and Vargo 2006), yet insufficient in others, leading to the probable pairing of related primary reproductives during colony founding (Vargo and Carlson 2006). Utilizing 13 and 12 microsatellite loci, DeHeer and Vargo (2004) found that the male and the female in the monogamous primary pair heading established mature colonies. These are nearly always unrelated to one another in some Reticulitermes species. However, the tandem pair formation in some Reticulitermes species appears to be random with respect to kinship (Kitade et al. 2006). DeHeer and Vargo (2006) reported that 26% of newly formed tandem-running pairs are significantly related to one another (sibling nestmates from the same colony) in R. flavieps, though only a few of these pairs successfully established colonies. The disparity of mating strategies revealed by genetic studies also suggest that tolerance for inbreeding is not a universal characteristic of termite breeding system (DeHeer and Vargo 2006). The lack of evidence

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supporting kin discrimination during colony founding is surprising considering the mounting evidence of negative effects of inbreeding in subterranean termite species (Fei and Henderson 2003; Husseneder et al. 2008), as well as in many plants and animals. The apparent variation in the tolerance to the potential effects of inbreeding in closely related sympatric species merits further investigation (Vargo and Husseneder 2009). 12.5.2

Inbreeding

Inbreeding is common within several eusocial groups, including naked mole-rats (Jarvis et al. 1994), social spiders (Aviles 1997), and termites (Myles 1999). Inbreeding has been considered favoring the evolution of termite eusociality in some conditions., Hamilton (1972) noted that termite ancestors are likely restricted to the specialized habitat of decaying wood, leading to further opportunity for and predispositions toward inbreeding. There are potential costs and benefits of inbreeding. Zootermopsis nevadensis Termopsidae shows inbreeding avoidance, suggesting that mating with relatives may sometimes have costs (Shellman-Reeve 2001). To the contrary, Z. angusticollis colonies founded by sibling pairs have lower mortality than nonsibling pairs during the incipient stages of colony foundation (Rosengaus and Traniello 1993). Inbreeding within termites, in general, can arise through one of the two processes: 1) pairing of related kings and queens (sibling-sibling) during colony foundation; 2) replacement of colony-founding primary reproductives with secondary neotenics, either apterous or, more commonly, brachypterous reproductives (parent-offspring, sibling-sibling, and cousin-cousin) from within the colony (Myles 1999; Thorne et al. 1999). Pairing with related nestmates is assumed to be a common incidence in Rhinotermitidae because of two reasons: 1) winged termites are purported to be very weak fliers capable only of short distances from their natal nest (Nutting 1969); and 2) unrelated mates are often not available due to asynchronous mating flights among different colonies. Therefore, colonies should be often founded by brother-sister pairs, a condition that increases relatedness among family members and discourages selfishness within kin groups (Bartz 1979; Wade and LaFage 1987). As aforesaid that genetic analysis has revealed significant relatedness of newly formed tandem-running pairs in Reticulitermes spp, lending credence to this prediction (DeHeer and Vargo 2006). However, the genetic structure from the pairs heading established mature colonies indicates otherwise: the male and female are nearly always unrelated to one another in some Reticulitermes species (Vargo 2003; DeHeer and Vargo 2004). Two hypotheses have been proposed to explain the disparity in relatedness patterns for colony-founding pairs and within established colonies. One is inbreeding depression (ID), which eliminate inbred colonies early in development. Another is inbreeding avoidance (IA), which may result in a late parting of the related pairs in the tandem-running phase or after it is completed. The IA implies either historical or contemporary ID, and these results


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therefore suggest that, either directly or indirectly, ID could be a more potent force in the evolution of termites mating systems than is generally appreciated. It has been recognized that although inbreeding increases an individual’s relatedness to its siblings and to their offspring, it also boosts relatedness between individuals and their own offspring (Bartz 1979). Thus, if inbreeding continues generation after generation, the extent to which inclusive fitness is increased by helping to raise siblings over offspring becomes limited, if present at all, and the selective force for such behavior is weakened (Bartz 1979; Lacy 1980). With respect to inbreeding producing secondary neotenic reprouductives within the colony is considered the most important reproductive strategy for sustaining established colonies founded by primary reproductives (Thorne et al. 1999; Myles and Nutting 1988). Several studies have shown relatively slow growth rates observed in Reticulitermtes sp, even under the best-case conditions (Thorne et al. 1997) and rarely seen primary reproductives in field mature colonies (Howard and Haerty 1980). As aforementioned, one or more secondary neotenics can arise from either nymph pathway (brachypterous nymphtoids) or worker pathway (apterous ergatoids) to replace the primary queen and/or king within the colony or an orphaning group (also termed colony budding). Thus the inbreeding occurs within the colony between parent-offspring, sibling-sibling, or cousin-cousin. The cumulative egg production in colonies headed by multiple neotenics is substantially greater than that of colonies containing a single primary queen (Pawson and Gold 1996). 12.5.3

Breeding structures

Bartz (1979) suggests that termite colonies may have undergone alternating phases of outbreeding and inbreeding. In this model, alate reproductives disperse from the parental colony to mate with genetically unrelated alates to form a new colony that generate genetic asymmetries favorable to social evolution. Male and female neotenics subsequently develop in the parental colony and produce new alates by inbreeding. If a new colony is founded by an unrelated queen and king but each comes from an inbred colony, their offspring will be relatively homogeneous and thus more closely related to one another than they would be to their own (outbred) offspring. If subsequent generations within the colony are inbred, progeny would be more closely related to siblings than to outbred offspring. The fundamental life-history aspects of the model are plausible, but the determination of the extent to which ancestral termites fit the premises of the hypothesis is a challenge. Recent population genetics studies using molecular methods seem to support this hypothesis. According to the summarized model by Vargo and Husseneder (2009), an incipient colony is genetically a simple family composed of a monogamous pair of unrelated primary reproductives and their offspring. Eventually one or both of the primary reproductives senesces or dies, and then is replaced by neotenics within the colony, producing an extended family colony.

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The number of neotenics in these extended family colonies can vary from a few to several dozen. These colonies undergo several generations of inbreeding. Another breeding structure is mixed-family, so far only been demonstrated in R. flavipes (Vargo and Carlson 2006) and R. grassei (Nobre et al. 2008). Several mechanisms driven by alates or workers have been proposed to account for the formation of mixed families. Worker driven strategies have been documented (DeHeer and Vargo 2004). Infiltration of unrelated populations by unrelated dealates, one of the alate-driven strategies, has been observed by Hu and Ding (unpublished).

12.6

Asexual facultative parthenogenesis

Facultative parthenogenesis, or condition-dependent alternation of sexual and asexual reproduction, is widespread in animals. Parthenogenesis enables females to produce offspring by themselves, preventing reproductive failure when they do not find a mate before dying (Cuellar 1977). Therefore, facultative parthenogenesis is likely to be advantageous in certain situations, even though it may be ultimately inferior to sexual reproduction in terms of long-term fitness. Among termites, parthenogenesis has been documented in the lower termites Zootermopsis angusticollies (Light 1944), Z. nevadensis (Light 1944), Kalotermes flavicollis (Grasse 1949), Reticulitermes virginicus (Howard et al. 1981), R. speratus (Matsuura and Nishida 2001), and Velocitermes sp. Termitida (Stansly and Korman 1993), and is suspected in R. lucifugus (Herfs 1951) and Bifiditermes beesoni Kalotermitidae (Chhotani 1962). Parthenogenesis has been well studied in Reticulitermes speratus Kolbe. In this species, a colony can be formed through three different means: 1) sexually with a primary queen and king; 2) asexually by two delate females cooperatively; or 3) asexually by neotenics (both ergatoids and nymphoids) (Hayashi et al. 2003; Matsuura and Nishida 2001; Matsuura et al. 2004). Female alates in this species always need a partner for colony foundation and so make sure they retain one, whether male or female, kin or nonkin. If a partner male is present, only one female will survive in colony, even though multiple females are introduced initially. In the absence of a partner male, two females, but never more than two, found a colony cooperatively. Female dealates that fail to pair with males form colonies cooperatively with partner females and reproduce by parthenogenesis (Matsuura et al. 2002). The cooperation between two females promotes their survival rates, much like a monogamous foundation. The mtDNA restriction fragment length polymorphism (RELP) analysis reveals that the two queens each produce the same number of offsprings within the first 100 day of colony foundation, though the number of offspring is fewer per capita than the only female in colonies founded by one queen and one king. Furthermore, initially smaller females gained significantly more weight than the initially larger females, indicating altruistic behaviors of the larger females towards the small ones. Interestingly, if one of the colony founders dies, the other will also die not long


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after. Most recently, Matsuura et al. (2009) censused 30 natural colonies and found only one colony has its primary queen replaced with 128 secondary queens. This colony contained a single primary king, the secondary queens were exclusively produced parthenogenetically by the founding primary queen, whereas workers and alates were produced by normal sexual reproduction. Interestingly, all parthenogenetic offsprings were female and all most all differentiated into nymphs that later become either alates or secondary reproductives (nymphoids with wing buds). In contract, offspring from different pairs diverged in both caste and sex ratios (Hayashi et al. 2007). It is known that parthenogenes are generally less viable than sexually produced offspring in various facultative parthenogenetic insect species (Roth and Willis 1956). In Z. angusticollis, parthenogenic larvae have low survival rates and are poor replacement reproductives (Light 1944). A female could utilize one generation of parthenogenic larvae to found the colony if the co-founder of the colony dies, but sexual reproduction is necessary to ensure long-term survival of the colony. Her first brood might be a mixture of asexually and sexually generated offspring, depending on whether the make was ready to copulate before she completed the first bout of oogenesis. In the R. speratus termites, the unmated diploid females produce diploid offspring via parthenogenesis, the same way as sexually reproduce offspring (Matsuura et al. 2004). Even if parthenogenesis is inferior to sexual reproduction in terms of fitness, facultative parthenogenesis may be still advantageous. The parthenogenetic eggs of R. speratus termite have a greater mortality and longer hatching period but a high viability and short preoviposition period. Therefore, parthenogenesis should not be explained as an accidental event but a “best of a bad job” strategy for termites.

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Section 3: Insect Toxicology and Insecticide Resistance Management

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CHAPTER 13

P450–mediated Insecticide Detoxification and Its Implication in Insecticide Efficacy Zhimou Wen, Xing Zhang and Yalin Zhang

Abstract Insecticides have been extensively used to control insect pests that transmit human diseases and damage our crops by feeding or indirectly by transmitting plant diseases. However, insect pests still pose serious challenges to human health and agricultural production today because they can avoid or reduce the harm of insecticides via various mechanisms, including P450–mediated insecticide detoxification. P450s are a very important enzymatic system capable of metabolizing structurally different compounds including insecticides and are thus called “the most versatile biological catalyst(s)” (Porter and Coon 1991). In this chapter we discuss how insect pests deal with the challenges posed by synthetic insecticides with a focus on P450–mediated insecticide detoxification and its implication in insecticide efficacy. P450–mediated insecticide detoxification is one of the bases for differential tolerance of insects to insecticides, i.e. selective toxicity of insecticides. The tolerance of insects to insecticides can be reduced or enhanced by inhibitors or inducers of P450s involved in insecticide detoxification. Constitutive overexpression of P450s involved in insecticide detoxification is one of the most important mechanisms for insecticide resistance. Concerted research efforts in this area may provide better insight in the development of specific strategies for effective use of currently available insecticides or in developing new insect control agents. Keywords P450, insecticide, detoxification, tolerance, resistance

Zhimou Wen Department of Entomology, University of Georgia, Athens, GA, 30602, USA E-mail: [email protected] Xing Zhang, Yalin Zhang College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China


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13.1

Zhimou Wen, Xing Zhang and Yalin Zhang

Insecticides

Insecticides are a subclass of pesticides that target insect pests. A pesticide is defined as any substance or mixture of substances intended for preventing, destroying, repelling, or mitigating any pest (EPA: http://www.epa.gov/ pesticides/about). Insecticides have been extensively used to control insect pests because vector insects transmit human diseases, e.g. malaria by mosquitoes, and plant–eating insects damage our crops directly by feeding and indirectly by transmitting plant diseases. It is painful to recognize that insect pests are still posing serious challenges to human health and agricultural production today. For example, 3 billion people in 109 countries and territories are at risk of malaria infection transmitted by mosquitoes with about 250 million annual cases and one million deaths (WHO 2008). DDT, as one of only 12 available synthetic insecticides recommended by World Health Organization (WHO) for mosquito control (WHO 2006, 2007), was ironically the backbone of the Global Malaria Eradication Program (GMEP) initiated by WHO during the 1950s and 1960s but banned in the 1970s because of environmental concerns. An obvious question is: "How do insect pests successfully deal with the challenges posed by synthetic insecticides – the very chemicals that man has invented for their destruction?" The answer to this question is very important because not only may it illuminate the fascinating phenomenon of rapid evolution and genetics of insecticide resistance (IR) but it also has practical implications in pest control. Only by in– depth understanding of insect interactions with insecticides can we develop a strategy for the effective use of currently available insecticides or for developing new control agents. Today, this understanding becomes ever more urgent. On one hand, synthetic insecticides will continue to be an integral part of pest management strategy in the foreseeable future and are the only option in case of a severe pest outbreak or emergency when pests cannot be controlled by other means (Committee on the Future Role of Pesticides et al. 2000; Zaim and Guillet 2002; Ware and Whitacre 2004). On the other hand, our insecticide repertoire is rapidly dwindling due to IR, government regulation and increasing difficulty in bringing new materials to the market (Casida and Quistad 1998; Scott 1999; Ware and Whitacre 2004). In theory, every physiological system can be a target for insect control. Not surprisingly, virtually all insect systems have been targeted in one way or another by a broad array of insecticides. For instance, insect growth regulators (IGRs) such as methoprene and tebufenozide target the endocrine system by acting as juvenile hormone mimic and ecdysone agonist, respectively. Boric acid and silica gel target the cuticle by causing dehydration and death through physical abrasion. Rotenone and mineral oil target the respiratory system by inhibiting NADH– ubiquinone oxidoreductase and blocking the openings of spiracles, respectively. However, the most widely used insecticides target two systems: the digestion system embodied by Bt endotoxins (Griffittsm and Aroian 2005), which are not the focus of this discussion, and the nervous system exemplified by the major

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groups of synthetic insecticides (Casida and Quistad 1998; Casida 2009). It is not surprising that the most widely used insecticides target the nervous system because they work very fast. DDT and pyrethroids affect propagation of action potentials along axons by targeting the voltage–gated sodium channels. Cyclodienes and phenylpyrazols target the γ–aminobutyric acid (GABA)–gated chloride channel. Organophosphates and carbamates inhibit acetylcholinesterase (AChE) which degrades the neurotransmitter acetylcholine right after it completes its function in synaptic neuronal signal transmission. Inhibition of AChE results in excessive accumulation of acetylcholine in the synaptic cleft leading to overstimulation of the nervous system. Neonicotinoid insecticides compete with acetylcholine in binding to the nicotinic acetylcholine receptor (nAChR) and therefore disrupt the normal signal transmission across the synaptic cleft. No matter what the specific target for an insecticide might be, molecules of the insecticide must reach the target site to be effective. Only a portion of insecticide molecules that enter insects can reach the target site because the remaining portion may be sequestered, metabolized or excreted. Metabolism is the most important factor responsible for the loss of insecticide molecules that are on their way to reach the target site. Insecticide metabolism is mainly accomplished by three enzyme systems that include carboxylesterases (COEs), glutathione transferases (GSTs) and P450s (Hemingway and Karunaratne 1998;Scott 1999; Hemingway et al. 2004; Enayati et al. 2005; Feyereisen 2005; Li et al. 2007) in a two–phase process. A functional group, for example, –OH, is usually added to a parent compound in phase 1 reactions, a process called functionalization. The modified compound is then covalently linked to another functional group in phase 2 reactions, a process called conjugation. COEs and P450s are major phase 1 enzymes and GSTs are phase 2 enzymes. Through these two–phase reactions, a portion of insecticide molecules that enter insects are transformed to less toxic metabolites and excreted. The focus of this chapter is on P450s involved in insecticide detoxification and how a better understanding of this process enables better decisions in pest management when insecticides are needed.

13.2

Insect P450s

Cytochrome P450 monooxygenases (P450 monooxygenases) are a very important enzymatic system capable of metabolizing structurally different compounds and thus called “the most versatile biological catalyst(s) known” (Porter and Coon 1991). P450 monooxygenase–mediated reactions can be generalized as a split of molecular oxygen with one oxygen atom incorporated into the substrate and the other oxygen atom reduced to H2O. The reactions require electrons provided by NADPH via redox partner(s) which are adrenodoxin and adrenodoxin reductase for mitochondria P450s and P450 reductase for microsomal P450s. The key component of this enzymatic system is

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cytochrome P450 (P450) which is a heme–thiolate protein (Mansuy 1998) with a characteristic absorbance peak at 450 nm when its reduced form is combined with carbon monoxide (Omura and Sato 1964). In P450 monooxygenase–mediated reactions, P450 binds substrate and acts as terminal oxidase. Because of its key role in the P450 monooxygenase system, the term P450 is often used as synonym for cytochrome P450 monooxygenase. In our discussion throughout this chapter, P450 represents cytochrome P450 monooxygenase when enzymatic activity is discussed and cytochrome P450 when the component of cytochrome P450 monooxygenase system is referred. Eukaryotes possess multiple P450s that are located in both endoplasmic reticulum and mitochondria (Omura 2006). Based on the annotation of several recently completed genome sequencing projects, the numbers of P450s in insects range from 47 in Apis mellifera (honey bee) to 164 in Aedes aegypti (yellow fever mosquito). The model organism Drosophila melanogaster has 84 and the malaria vector Anopheles gambiae 105 (Scott 2008). In contrast, humans have 57 P450s (Nelson et al. 2004). All of the current P450s are believed to have descended from a common ancestral gene by gene duplication and adaptive diversification (Gotoh 1993). The number of P450s appears to have increased rapidly during the past 400 million years, reflecting probably intense animal–plant warfare (Gonzalez and Nebert 1990). The current nomenclature system based on amino acid (AA) sequence was first introduced in 1987 (Nebert et al.1987) and has been under constant revision to reflect the rapid accumulation of new P450 sequences (Nelson et al. 1993, 1996 and Nelson P450 website). A P450 is named CYP followed by a number and a letter to assign the P450 to a specific family and subfamily respectively. Another number then follows the letter to designate the individual P450. For example, CYP6L1 which was isolated from Blattella germanica (German cockroach) (Wen and Scott 2001) is the first P450 named in subfamily L within family 6. P450s are generally grouped into the same family for members with AA identity > 40％ and the same subfamily for members with AA identity > 55％, though there are many exceptions to this rule (Feyereisen 2005). The overall three–dimensional structures are well conserved for all P450s though their primary AA sequences may differ significantly. Since the nomenclature system is based on overall AA identity and considering that a single AA change may dramatically alter substrate specificity (Linderberg and Negishi 1989; Wen et al. 2005), function cannot be predicted for a given P450 solely based on its name. An important feature of P450s is the complexity of their substrate specificity due to existence of multiple forms that may have broad and overlapping substrates. As a result, P450s not only catalyze biosynthetic reactions to produce juvenile and ecdysteroid hormones that have essential physiological functions (Feyereisen 2005; Rewitz et al. 2006) but also catalyze detoxifying/bioactivating reactions that bear significant ecological and toxicological consequences (Scott and Wen 2001; Berenbaum 2002; Feyereisen 2005; Li et al. 2007). While biosynthetic reactions are carried out by P450s coded by stable genes conserved


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for “core functions in development and physiology”, detoxification reactions are often catalyzed by P450s coded by unstable genes in response to “unstable environmental interactions” (Thomas 2007). Insecticides are destructive external environmental factors imposed on insects by human beings. In response to this destructive challenge, insects mobilize their “environmental response genes” which are defined by Berenbaum (2002) as “those encoding proteins involved in interactions external to the organism, including interactions among organisms and between the organism and its abiotic environment.” As an important group of the “environmental response genes,” insect P450s are involved in the detoxification of insecticides and therefore play important roles in the interactions of insects with insecticides, an embodiment of the “unstable environmental interactions.” The complexity of the P450 system has historically impeded functional definition for individual P450s, yet it is important because only by dissecting the roles of individual P450s can we have a better appreciation of the roles of the P450 system in insect interactions with insecticides. For practical consideration, characterization of individual insect P450s in insecticide detoxification can provide insight for the enhancement/ mitigation of insecticide toxicity.

13.3 Insect P450s and differential tolerance to insecticides among insect species Insecticides are meant to control insects. However, not all insecticides are equally toxic to a given insect species, neither is a given insecticide equally effective against all insect species. While less selective insecticides are useful for the control of a broad range of pests, highly selective insecticides are preferred when non–target organisms are of a concern. Reflection of the selective toxicity of insecticides to different insect species is the differential tolerance of insect species to insecticides. From the insect side, differences among insect species in many aspects including size, behavior, insecticide penetration, target sensitivity, excretion and metabolism can all contribute to differential tolerance to insecticides. In the case of insecticide detoxification, if pest A is more effective than pest B in detoxification of a given insecticide, with other factors being equal, it can be predicted that pest A will be more tolerant to this insecticide. As one of the most important mechanisms that affect the process of insecticide molecules reaching their target site, P450–mediated insecticide detoxification has a great impact on insecticide toxicity. Differences among insect species in their capacity for P450–mediated detoxification of insecticides are an important factor responsible for differential tolerance among insect species to insecticides, and therefore, an important basis for developing selective insecticides. Imidacloprid is an insecticide of the neonicotinoid class, the most important development for synthetic insecticides over the last 3 decades. While the target sites of imidacloprid, nAChRs, are fundamentally different between insects and

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mammals conferring the later preferential protection, their binding affinity to imidacloprid is very high across a broad range of insect species from many different Orders including Homoptera, Diptera, Blattaria, Lepidoptera and Orthoptera (Tomizawa and Casida 2003, 2005). Yet, imidacloprid is highly toxic only to sap–sucking insects such as aphids but only moderately toxic to B. germanica and Musca domestica (house flies). The major reason for the tolerance of these two pest species to imidacloprid is that P450s in these insects are highly active in imidacloprid detoxification. This is evidenced by the dramatic increase of toxicity when imidacloprid is applied together with piperonyl butoxide (PBO), a general P450 inhibitor (Liu et al. 1993; Wen and Scott 1997). Similar to imidacloprid, tau–fluvalinate, a pyrethroid insecticide highly toxic to varroa mites, is technically nontoxic to A. mellifera because honey bee P450s can efficiently detoxify tau–fluvalinate (Johnson et al. 2006). In fact, this selective tolerance has made tau–fluvalinate an ideal agent for controlling varroa mites which are a big threat to A. mellifera colonies (Williams 2000). Evidence implicating insect P450s in insecticide metabolism has been presented for a long time (See Agosin 1985 and Hodgson 1985 for review) and the first reports for the involvement of specific P450s in insecticide metabolism appeared in the 1990s showing that CYP6D1, CYP6A1 and CYP12A1 of M. domestica can detoxify insecticides: aldrin, heptachlor and diazinon by CYP6A1 (Andersen 1994; Sabourault et al. 2001), chlorpyrifos, chlorpyrifos oxon, deltamethrin and cypermethrin by CYP6D1 (Hatano and Scott 1993; Wheelock and Scott 1992; Zhang and Scott 1996; Korytko and Scott 1998) as well as aldrin, heptachlor and diazinon by CYP12A1 (Guzov et al. 1998) (Table 13.1). Since the start of the new millennium, about a dozen more have been added to the list of insect P450s whose activities in metabolizing insecticides have been unequivocally demonstrated (Table 13.1). From Table 13.1 we can see several patterns. Most of the individual P450s can metabolize multiple insecticides and P450s from the same species can have overlapping substrates. For example, CYP6B8, CYP6B27 and CYP321A1 from Helicoverpa zea can all detoxify insecticides from different classes: aldrin (cyclodiene), diazinon (organophosphate), carbaryl (carbamate) and α– cypermethrin (pyrethroid) (Li et al. 2004; Sasabe et al. 2004; Rupasinghe et al. 2007; Wen et al. 2009). Although some P450s have only one insecticide as substrate reported, we should not assume that these P450s have only one substrate, rather that other insecticides as substrates have yet to be determined. It is also clear that the definition of individual P450s in insecticide metabolism has focused mainly on model organisms and insect pests of significance in human health and agriculture. The pace in defining functions for individual P450s involved in metabolism of insecticides is accelerating, especially in recent years. However, considering the large numbers of insect P450s potentially involved in insecticide detoxification and the urgent need for such research, we have just tapped a tip of the P450 iceberg.

P450–mediated Insecticide Detoxification and Its Implication in Insecticide Efficacy Table 13.1

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Insect P450s involved in insecticide metabolism

Species

P450s

Insecticide substrates

References*

M. domestica CYP6A1

aldrin, heptachlor, diazinon

1, 2

CYP6D1

chlorpyrifos, chlorpyrifos oxon, deltamethrin, cypermethrin

3, 4, 5, 6

CYP12A1

aldrin, heptachlor, diazinon, azinphosmethyl, amitraz

7

D. melanogaster CYP6a2

DDT, aldrin, dieldrin, diazinon

8, 9

CYP6g1

DDT, imidacloprid

10

CYP6Z1

DDT, carbaryl

11

CYP6Z2

Carbaryl

11

CYP6P3

trans- and cis-permethrin, deltamethrin

12

CYP6AA3

deltamethrin

13

A. gambiae

A. minimus H. zea CYP6B8

aldrin, diazinon, carbaryl, α-cypermethrin

14, 15, 16

CYP6B27


16

CYP321A1


15, 16, 17

CYP9A12

esfenvalerate

18

CYP9A14

esfenvalerate

18

CYP6B

diazinon

14

H. armigera

P. polyxenes

*References: 1. Anderson et al. 1994; 2. Sabourault et al. 2001; 3. Wheelock and Scott 1992; 4. Hatano and Scott 1993; 5. Zhang and Scott 1996; 6. Korytko and Scott 1998; 7. Guzov et al. 1998; 8. Dunkov et al. 1997; 9. Amichot et al. 2004; 10. Joussen et al. 2008; 11. Chiu et al. 2008; 12. Muller et al. 2008; 13. Boonsuepsakul et al. 2008; 14. Li et al., 2004; 15. Rupasinghe et al. 2007; 16. Wen et al. 2009; 17. Sasabe et al., 2004; 18. Yang et al. 2008.

13.4 Regulation of P450s and insect tolerance to insecticides Besides their multiplicity and broad and overlapping substrate specificity, another amazing feature of insect P450s is their complicated regulation. P450s can be inhibited or induced. While inhibition decreases an enzyme’s activity, induction increases the activity. For those insect P450s involved in insecticide detoxification, decrease or increase in activities can have significant ecological consequences for insects, some to their benefit and others to their harm. Inhibition compromises P450’s capacity to detoxify insecticides, therefore chemicals with P450–inhibiting capacities have been employed as synergists to

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enhance insecticide toxicity. PBO, a well–known P450 inhibitor, has been extensively used not only commercially as a synergist to enhance the toxicity of pyrethroid and carbamate insecticides but also diagnostically as a tool in insecticide toxicology for determining the involvement of P450s in insecticide metabolism (Section 3) and IR (Section 5) (Hodgson and Levi 1998). PBO inhibits P450 activity by forming a stable and inhibitory P450–metabolite complex (Hodgson and Levi 1998). PBO is synthesized based on the structure of methylenedioxyphenyl (MDP) compounds such as sesamex and safrole found in many plant species. Since plants synthesize a variety of toxins for their defense against herbivores, the co–occurrence of MDP chemicals is hypothesized to enhance plant defense by synergizing the toxicity of plant toxins, an evolutionary response of plants to the breach of their first line of chemical defense by herbivores (Berenbaum 1985, 1990; Berenbaum and Neal 1985, 1987; Neal 1989; Neal and Berenbaum 1989; Wen et al. 2006). MDP compounds are not the only chemicals with P450 inhibiting capacity. Chemicals of various structures, natural or synthetic, can inhibit the activities of insect P450s (Neal and Wu 1994; Scott 1996; Baudry et al. 2003; Wen et al. 2006; Mclaughlin et al. 2008). With the establishment of heterologous expression systems for insect P450s (Andersen et al. 1994; Wen et al. 2003) and the development of monospecific antibody for individual insect P450s (Wheelock and Scott 1990), we have already been able to successfully screen for isoform specific inhibitors for insect P450s (Scott 1996; Baudry et al. 2003;Wen et al. 2006; Mclaughlin et al. 2008). Like in mammalian systems where identification of specific inhibitors for individual P450s and understanding their mechanisms are important for rational chemical manipulation of drug–drug interactions (Kalgutka et al. 2007), we can anticipate that identification of specific inhibitors for individual insect P450s would prove useful in rational development of insecticide synergists that block the activities of P450s involved in insecticide detoxification. In sharp contrast to inhibition, induction increases P450’s detoxification activity. Induction of P450s in herbivorous insects by plant allelochemicals is a common phenomenon and it allows insect herbivores to rapidly detoxify not only a wide variety of plant defensive toxins occurring in their hosts (Li et al. 2007) but a broad range of insecticides as well (Yu 1986). The consequence of inducing P450s involved in the detoxification of an insecticide is enhanced insect tolerance to the insecticide, i.e. reduced efficacy of the insecticide against the insect. Induction of P450s in herbivorous insects by plant allelochemicals conferring insect induced tolerance to insecticides was first recognized in the 1970s and 1980s when groups of the same insect species, feeding on different plant species or artificial diet in the presence or absence of a specific allelochemical, varied in their P450 activity and tolerance to insecticides (Yu 1986; Dominguez–Gil and Mcpherson 1999; Feyereisen 2005). These early studies provided great insight for insect P450 regulation by plant allelochemicals and for the control of herbivorous insect pests using insecticides (Yu 1982). Given the existence of multiple P450s in each insect species and the broad and overlapping substrates for


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some P450s, however, most of the reported P450 activities using model substrates in the early studies might be irrelevant to the P450(s) that detoxifies the insecticide or maybe they are the combined activities of several P450 enzymes. Since the publication of the first insect P450 sequence in 1989 (Feyereisen et al. 1989), many insect P450s have been cloned and their induction by foreign chemicals including plant allelochmicals has been determined to be mainly at the transcriptional level (Feyereisen 2005). The establishment of heterologous expression systems for insect P450s (Anderdson et al. 1994; Wen et al. 2003) has made it possible to determine if individual P450s are involved in induced insect tolerance to insecticides. CYP6B8, CYP6B27 and CYP321A1 are P450s cloned from H. zea, a polyphagous herbivore with more than 100 host plants (Li et al. 2000, 2002; Sasabe et al. 2004). Metabolic analyses using heterologously expressed proteins indicated that all three enzymes could metabolize insecticides of several classes including pyrethroids (α–cypermethrin), organophosphates (diazinon), carbamates (carbaryl) and cyclodienes (aldrin) (Sasabe et al. 2004; Li et al. 2004; Wen et al. 2009). The transcripts of these three P450s were differentially induced by a broad range of plant allelochemicals and synthetic chemicals (Li et al. 2000, 2002; Sasabe et al. 2004; Zeng et al. 2007; Wen et al. 2009). Among the plant allelochemicals, coumarin and rutin are frequently encountered by H. zea larvae, indole–3–carbinol (I3C) is occasionally encountered and, xanthotoxin and flavone are rarely encountered. All these five chemicals could significantly induce CYP321A1 which was constitutively expressed at low levels, but not CYP6B27 which was constitutively expressed at high levels in the fifth instar. Only xanthotoxin, coumarin and flavone could induce CYP6B8 which was constitutively expressed at a level between CYP321A1 and CYP27B1 (Li et al. 2002; Sasabe et al. 2004; Zeng et al. 2007; Wen et al. 2009). When caterpillars feeding on diet in the presence of one of these allelochemicals were challenged with α–cypermethrin, diazinon or carbaryl, they displayed a much higher tolerance to these insecticides with only one exception: addition of rutin in the diet did not significantly change the tolerance of caterpillars to carbaryl (Wen et al. 2009). Based on these results, it seems that CYP6B27 plays important role for the basal tolerance of H. zea larvae to insecticides while CYP321A1 and CYP6B8 are more important for induced tolerance of H. zea larvae to insecticides.

13.5

Insect P450s and insecticide resistance

Insecticide resistance (IR) is defined as the development of a heritable ability in a strain of some insects to survive a toxicant which would kill the majority of individuals in a normal population of the same species (WHO 1957). In a more practical sense, it is defined as “a heritable change in the sensitivity of a pest population that is reflected in the repeated failure of a product to achieve the expected level of control when used according to the label recommendation for

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that pest species” (IRAC: http://www.irac–online.org). IR is different from induced insect tolerance to insecticides in that IR is constitutive while induced tolerance only occurs in the presence of inducer(s). IR has severe consequences such as failure of control, increased application frequency, increased dosage, decreased yields, environmental contamination, and destruction of non–target organisms including natural enemy (Scott 1991). IR is so rampant that more than 500 species of insects and mites have been reported to have developed resistance to one or more insecticides since the introduction of DDT for insect control in the 1940s (Denholm et al. 2002). Among the major IR mechanisms (behavioral change, slowed penetration, target site insensitivity and increased metabolic detoxification), metabolic detoxification is the most important and is mainly accomplished by three different enzyme systems: COEs, GSTs and P450s (Scott 1999; Hemingway et al. 2004). P450–mediated IR has been found in many important insect pests and may confer high levels of resistance (Scott 1999; Feyereisen 2005). Due to the broad substrate spectra of some P450s, this mechanism may potentially affect several classes of insecticides and therefore confer cross–resistance to unrelated compounds (Hodgson 1985; Scott 1999). While P450s have long been associated with IR, identification of specific P450s responsible for IR has proven difficult. It has been a common practice to associate a P450 with IR if it is expressed at a higher level in resistant strains compared to susceptible strains. But this practice may be misleading as explicitly demonstrated in the title of a paper: “High expression of Cyp6g1, a cytochrome P450 gene, does not necessarily confer DDT resistance in Drosophila melanogaster” (Kuruganti et al. 2007). Just five years before the publication of this paper, overtranscription of Cyp6g1 was declared to be “both necessary and sufficient for resistance” of D. melanogaster to a broad range of insecticides (Daborn et al. 2002). Two criteria were therefore proposed to demonstrate a P450’s involvement in IR (Scott et al. 1998; Scott 1999): (1) the P450 must detoxify (or sequester) the insecticide to which a strain was claimed to have P450–mediated IR and (2) the P450 should have a greater amount in the resistant strain or the resistant P450 allele should have a greater catalytic activity (i.e. detoxification) compared to its susceptible allele. Based on these criteria, P450–mediated IR can be the result of increased catalytic activity of a P450 and/or increased expression in resistant strains of a P450 that detoxifies the insecticide. While many P450s have been shown to have higher expression levels in resistant strains, only a few can be unequivocally associated with IR because activities for most of these P450s in metabolizing relevant insecticides have not yet been characterized. Among the few P450s that meet the above criteria are CYP6D1 from M. domestica resistant to pyrethroid insecticides in the LPR strain (Scott 1999), CYP6A1 and CYP12A1 from M. domestica resistant to organophosphate insecticides in the Rutgers strain (Carino et al. 1994; Guzov et al. 1998; Sabourault et al. 2001), Cyp6a2 from D. melanogaster resistant to DDT in RRDTR strain (Brun et al. 1996; Amichot et al. 2004), CYP6AA3 from A. minimus resistant to pyrethroid in F19 and F25 populations (Rodpradit et al.


239

2005; Boonsuepsakul et al. 2008) and CYP9A2 and CYP9A14 from H. armigera resistant to pyrethroid insecticides in YGF strain (Yang et al. 2006, 2008). During the last decades, one of the most active and fascinating research areas concerning IR has been defining P450–mediated IR in disease–transmitting vectors in an attempt to find solutions to deal with the growing problem of IR in vectors. With the completion of genomic sequencing projects for several insect species including the major malaria vector A. gambiaeand dengue fever vector A. aegypti as well as the advancements in molecular biotechnology including microarray for profiling genome–wide gene transcription, proteomics for profiling genome–wide protein expression and heterologous expression of recombinant proteins for defining gene function, we have been moving forward at an unprecedentedly fast pace in identifying P450s involved in IR in recent years. In contrast to the conventional methods that examine the expression profiles of transcripts or proteins for one to several genes at a time, microarray and proteomics simultaneously examine the expression profiles of transcripts or proteins for a large number of genes. For example, a detoxification microarray chip was constructed for 230 genes covering COEs, GSTs, P450s and redox genes for A.gambiae and used to compare the expression of these genes between insecticide resistant and susceptible strains. Among the five genes that were strongly up–regulated in DDT resistant ZAN/N strain are two P450s: CYP6Z1 and CYP12F1 (David et al. 2005). The evidence for the involvement of CYP6Z1 in DDT resistance was finally presented when metabolic assay using CYP6Z1 coexpressed with P450 reductase and cytochrome b5 indicated that CYP6Z1 metabolized DDT (Chiu et al. 2008). Similarly, CYP6P3 is the culprit responsible for high levels of pyrethroid resistance in several populations of A. gambiae (Djouaka et al. 2008; Muller et al. 2008).

13.6

Conclusions

As one of the most important groups of “environmental response genes,” P450s play important roles in the interactions of insects with insecticides. However, the complexity of the P450 system has for a long time impeded our efforts to decipher the roles of individual P450s in these interactions. The publication of genome sequences for several insect species, recent advancements in modern biotechnonology including genomic and proteomic techniques, heterologous expression of recombinant proteins and RNAi technology have provided an excellent platform for studying gene functions, one of the principal goals for the postgenomic biology (Berenbaum 2002). Even though we have made unprecedented progress in the last decade, especially the last several years, in our understanding of the roles of individual P450s involved in insect interactions with insecticides, we still have a long way to go until we can fully appreciate the complexity of P450s’ involvement in these interactions considering the large numbers of insect P450s involved and their complicated regulation schemes. For example, we still know little about the mechanisms of inhibition and induction of P450s involved in

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insecticide detoxification in response to environmental factors including plant phytochemicals; only a few of the many P450s involved in insecticide detoxification and IR have been characterized. We believe concerted research efforts in these areas will provide better insight in the development of specific strategies for effective use of currently available insecticides or in developing new insect control agents. Acknowledgements We thank Dr. Arthur Zangerl at the University of Illinois (Urbana) and Dr. Andrew Nuss at the University of Georgia (Athens) for valuable comments on the manuscript.

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CHAPTER 14

House Fly Ctyochrome P450s: Their Role in Insecticide Resistance and Strategies in the Isolation and Characterization Nannan Liu and Fang Zhu

Abstract Insect cytochrome P450s are known to play an important role in detoxifying insecticides and plant toxins, resulting in the development of resistance to insecticides and facilitating the adaptation of insects to their plant hosts. Insect P450s are associated with enhanced metabolic detoxification of insecticides in insects, as evidenced by the increased levels of P450 proteins and P450 activities that result from constitutively transcriptional overexpression of P450 genes in insecticide resistant insects, and some insect P450 genes can be induced by exogenous and endogenous compounds. Both constitutively increased expression (overexpression) and induction of P450s are thought to be responsible for increased levels of detoxification of insecticides. This chapter reviews strategies for the isolation of cytochrome P450s from house flies and the characterization of their possible importance in insecticide resistance. Keywords Cytochrome P450, metabolic detoxification, insecticide, resistance

14.1

Introduction

Cytochrome P450s constitute one of the largest gene superfamilies in all living organisms, including mammals, fish, arthropods, fungi, plants, and bacteria, and possess a great diversity of physiological and biochemical functions. In insects, more than 1000 P450s have so far been identified, distributed over 150 subfamilies of 40 known P450 gene families (http://drnelson.utmem.edu/P450. stats.2006.htm). Insect cytochrome P450s are known to play an important role in

Nannan Liu Department of Entomology and Plant Pathology, Auburn University, Auburn, AL, 36849, USA E-mail: [email protected] Fang Zhu Department of Entomology, S-225 Agricultural Science Center North, University of Kentucky, Lexington, KY, 40546-0091, USA


House Fly Ctyochrome P450s

247

detoxifying exogenous compounds such as insecticides (Scott 1999; Feyereisen 2005) and plant toxins (Berenbaum 1991; Schuler 1996). Increased P450 protein levels and P450 activities resulting from transcriptional up-regulation of P450 genes have been shown to be involved in the enhanced metabolic detoxification of insecticides and plant toxins in insects, leading to the development of resistance to insecticides (Carino et al. 1994; Liu and Scott 1997, 1998; Kasai et al. 2000; Feyereisen 2005) and tolerance to plant toxins (Li et al. 2002; Wen et al. 2003). Insect P450s are also an important part of the biosynthesis and degradation pathways of endogenous compounds such as pheromones, 20hydroxyecdysone, and juvenile hormone (JH) (Reed et al. 1994; Sutherland et al. 1998; Winter et al. 1999; Gilbert 2004; Niwa et al. 2004) and thus play important roles in insect growth, development, and reproduction. However, although their importance in insect physiology and toxicology is widely recognized, there are enormous gaps in our knowledge of insect P450s. In particular, their precise role in the regulation processes that govern the evolution of insecticide resistance, which typically requires the interaction of multiple genes, has not yet been determined. As more P450 sequences become available, our understanding of the roles of P450s in physiological and toxicological processes should improve.

14.2

The house fly

The house fly, Musca domestica, is not only a serious pest affecting livestock and poultry facilities, but also poses a major threat to human health by transmitting the pathogens that cause many human diseases. House flies are the vectors of more than 100 human and animal intestinal diseases (Scott and Lettig 1962; Greenberg 1965; Keiding 1986; Scott et al. 2009). Consequently, there is a substantial effort devoted to controlling fly populations worldwide, primarily with insecticides such as pyrethroids. However, house flies have shown a remarkable ability to rapidly evolve resistance to each of the insecticides used against them (Keiding 1999; Liu and Yue 2000). In a series of studies designed to investigate how this resistance develops, a house fly strain, designated ALHF, was collected from a poultry farm in Alabama after a control failure with permethrin (a pyrethroid insecticide), and further selected with permethrin for 6 generations after collection in the laboratory to reach a 6,600-fold resistance (Liu and Yue 2000, 2001). Synergism studies found that permethrin resistance in ALHF was largely suppressed by piperonyl butoxide (PBO), indicating that P450 monooxygenase-mediated detoxification may be one of the major mechanisms involved in the development of pyrethroid resistance in ALHF (Liu and Yue 2000). Genetic linkage analysis provided further evidence pointing to the localization of PBO-suppressible-P450-mediated resistance on autosomes 1, 2, and 5 of ALHF (Liu and Yue 2001). Factors on autosome 5 are known to play a major role in P450-mediated resistance.

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Nannan Liu and Fang Zhu

14.3 Isolation of insect cytochrome P450 genes in ALHF house flies Several methods have been used to isolate insect P450 genes. One of the most common is to clone and isolate P450 genes by polymerase chain reaction (PCR) with degenerated primers (Snyder et al. 1996; Feyereisen 2005). In this method, the degenerated primers were designed according to the sequences of P450 conserved motifs PFxxGxRxCxG/A and GxE/DTT/S (Feyereisen 2005). A number of new P450s have been successfully cloned with the PCR method, including 17 CYP4 genes in Anopheles albimanus (Scott et al. 1994), 8 CYP genes from Helicoverpa armigera (Pittendrigh et al. 1997), 14 P450 fragments from Ceratitis capitata (Danielson et al. 1999), and 95 P450 sequences from 16 Drosophila species (Fogleman et al. 1998). Three strategies have been used for PCR cloning and sequencing P450 fragments from ALHF house flies in our laboratory. In strategy I, a primer pair consisting of a sense primer based on the P450 heme binding region, Flyh1, and an antisense 5’-anchored oligo(dT) primer (Tomita and Scott 1995), was used. In strategy II, the sense primer Flyc1, based on a conserved 13 amino acid region found in rat, human, and insect P450 sequences (Liu and Zhang 2004) was paired with an oligo(dT) antisense primer. In strategy III, one of degenerated sense primers based on the conserved region of insect P450 family 6, was used together with HemeR1, an antisense primer based on the complementary sequences of Flyh1 (Fig. 14.1, Table 14.1, Zhu and Liu unpublished data).

Fig. 14.1 Three strategies used to amplify P450 gene fragments from ALHF house flies. The blank box represents the P450 sequence. The small boxes with different patterns indicate the conserved regions from which the degenerated primers were designed. Degenerated or oligo(dT) primers were represented by arrows.

Using these 3 strategies, our group successfully cloned and sequenced 19 cDNA fragments from ALHF house flies (Table 14.2, Zhu and Liu, unpublished data). The P450 protein signature motif, FXXGXRXCXG, was present in all

House Fly Ctyochrome P450s Table 14.1

249

Degenerated primers used for amplifying P450 gene fragments in house flies

Strategies

Name of Primers

I

Sequence of Primers

Flyh1

5'-GGICCIAGIAACTGCATIGG-3'

II

Flyc1

5'-GGAAGTNGACACNTTYATGTT-3'

III

Heme R1

5'-CCIATGCAGTTICTIGGICC-3'

CYP6A1

5'-CYTTTGGCATTGARTGCARKAG-3'

CYP6AD1 CYP6D1

5'-CVTCNGSHAARATKAARHNNATG-3' 5'-GATCGYGGSVTBTAYGTKGAYG-3'

Table 14.2

Nineteen P450 cDNA fragments isolated from house flies (Zhu and Liu, unpublished)

Fragment Number

Sense Primer

Anti-sense Primer

Homologous Protein

Similarity (%)

Length (bp)

1

Flyh1

oligo(dT)

CYP6N3v3

57

308

2

Flyh1

oligo(dT)

CYP6G1

50

238

3

Flyh1

oligo(dT)

CYP9

51

273

4

Flyh1

oligo(dT)

CYP4D1

60

285

5

Flyh1

oligo(dT)

CYP4D14

75

289

6

Flyh1

oligo(dT)

CYP9

56

301

7

Flyh1

oligo(dT)

CYP4D14

69

405

8

Flyh1

oligo(dT)

CYP6A25

92

277

9

Flyh1

oligo(dT)

CYP6A5

98

274

10

Flyc1

oligo(dT)

CYP4D1

66

658

11

Flyc1

oligo(dT)

CYP4D8

57

874

12

Flyc1

oligo(dT)

CYP4D8

87

382

13

Flyc1

oligo(dT)

CYP4AC1

63

796

14

Flyc1

oligo(dT)

CYP4P3

53

975

15

CYP6AD1

Heme R1

CYP6A9

62

448

16

Flyh1

oligo(dT)

CYP6A9

66

231

17

CYP6A1

Heme R1

CYP6A24

96

976

18

CYP6AD1

Heme R1

CYP6A24

87

672

19

CYP6D1

Heme R1

CYP4G1

80

583

P450 fragments. In strategy I, with Flyh1 and oligo(dT) as the primers, PCR products of about 230–400 bp (including a variable 3’ UTR sequence) that coded for around 60 amino acids were isolated. In strategy II, with Flyc1 and oligo(dT) as the primers, fragments of about 650–950 bp (including a variable 3’ UTR sequence) were produced in all but one case, where the fragment of #12

250


(Table 14.2) had only 382 bp, possibly because of exon skipping in the 3’ end. In strategy III, with a degenerated primer based on the conserved region of insect P450 family 6 and the antisense primer Heme R1, ~ 450 – 970 bp PCR fragments were generated, encoding for about 150 to 320 amino acids in the middle region of P450 genes. The successful cloning of these 19 novel P450 fragments scattered in families 4, 6, and 9 from house flies using the PCR method indicates that the three strategies offer a useful approach for P450 cloning. Although most of these 19 P450s were cloned using the strategy I, the fragments cloned by this method tends to be very short and hard to obtain the full length sequences. The strategy II primarily produced CYP4 genes with fragments that were about half of the fulllength of P450 genes. Using the strategy III, CYP6 gene fragments were generated with the length about 440–970 bp, since the forward primer in strategy III was designed according to the conserved region of insect P450 family 6. Because it is not far from either 3’ or 5’ end of the sequence, the full length of each of these P450 genes can easily be obtained. Accordingly, if the forward primer was designed according to the conserved regions of other P450 gene families in the strategy III, new P450s scattered in those families could also be isolated. With the sequence information generated from these P450 fragments, the full-length of eight house fly's P450 genes, CYP4G13v1, CYP4D4v2, CYP4G2, CYP6A5, CYP6A5v2, CYP6A36, CYP6A37, CYP6A38, and CYP28B1 (Liu and Zhang 2002; Zhu and Liu 2008; Zhu et al. 2008a, b), have been isolated in our lab by 3’- and/or 5’-RACE. A phylogenic tree has been constructed to explore their phylogenic relationships with other house fly P450s previously reported (Fig. 14.2, Zhu and Liu unpublished data).

Fig. 14.2 Phylogeny of house fly P450 genes. The sequences of house fly P450 genes were processed by ClustalW2.


251

14.4 Constitutive overexpression and tissue specific expression patterns of P450 genes in house flies A significant characteristic of insect P450s that is associated with enhanced metabolic detoxification of insecticides is the increased levels of P450 proteins and P450 activities that result from constitutively transcriptional overexpression of P450 genes in insecticide resistant insects (Liu and Scott 1996, 1997, 1998; Feyereisen 2005; Scott 2008). Another important feature of insect P450 genes is that they may vary as to the tissues where they are expressed in response to physiological and environmental stimulators. In insects, the midgut and fat body tissues are generally considered to be the primary detoxification organs (Hodgson 1985) where most insect detoxification P450s are expressed (Scott et al. 1998). Thus, when characterizing the importance of the P450 genes in increasing metabolic detoxification of insecticides, it is necessary to evaluate the tissue specific expression of these genes in resistant insects. The differential expression patterns of all eight P450 genes, CYP4G13v1, CYP4D4v2, CYP4G2, CYP6A5, CYP6A5v2, CYP6A36, CYP6A37, CYP6A38, and CYP28B1 were characterized between resistant ALHF and susceptible CS house fly strains and among different tissues by the Northern blot analyses and quantitative real-time PCR (qRT-PCR) by our group. No significant difference in the expression of CYP4D4v2, CYP4G2, CYP4G13v1, CYP6A37, CYP6A38, and CYP28B1 between resistant ALHF and susceptible CS and aabys flies was observed, but CYP6A5v2 and CYP6A36 showed significant constitutive overexpression in the resistant ALHF strain. The expression of CYP6A36 was no of significant difference between the head + thorax and abdomen tissues of the CS strain; was lower in the head + thorax tissue and higher in the abdomen tissue of ALHF; and was significant high in both tissues of the ALHF strain compared with the CS strain. The expression of CYP6A5v2 was significantly higher in the abdominal tissue in both the susceptible CS and aabys flies and the resistant ALHF flies compared with their head + thorax tissues. Significant overexpression was more evident in both sets of tissues for ALHF flies compared with the tissues of CS and aabys flies. It has been proposed that constitutive overexpression of P450s is the insect's response as it adapts to changes in its environment (Daborn et al. 2002). In many cases, increased levels of P450 gene expression have resulted in increased levels of both total P450s and P450 activities and there is strong evidence to suggest that this is a major cause of insecticide resistance (Liu and Scott 1996, 1997, 1998; Feyereisen 2005). As midgut and most fat body components are located in the abdomen of insects and are known to be of primary importance in detoxificationrelated functions, the overexpression of the 2 P450 genes, CYP6A5v2 and CYP6A36, specifically in the abdomen tissues of ALHF suggest the importance of these genes in increasing the metabolic detoxification of insecticide in ALHF house flies, and provides further evidence supporting the important role of the P450 genes in insecticide resistance. It also seems highly likely that a significant

252


increase in an insect's resistance level is unlikely to be conferred by only one or two overexpressed genes and that multiple P450s will generally be involved in the development of resistance.

14.5 Co-up-regulation of P450 genes in response to permethrin exposure in house flies Another feature of some insect P450 genes is that their expression may be induced by exogenous and endogenous compounds (Feyereisen 2005), a phenomenon known as induction. It has been suggested that the induction of P450s and their activities in insects is involved in the adaptation of insects to their environment and the development of insecticide resistance (Terriere 1983, 1984). While all insects probably possess some capacity to detoxify insecticides and xenobiotics, the degree to which insects can metabolize and detoxify these highly toxic chemicals is of considerable importance to their survival in a chemically unfriendly environment (Terriere 1984). Insects may use various biochemical pathways to enable them to tolerate the lethal action of insecticides. For example, increased cytochrome P450 detoxification is known to play an important role in many insect species. Both constitutively increased expression (overexpression) and induction of P450s are thought to be responsible for increased levels of detoxification of insecticides. However, unlike constitutively overexpressed P450 genes, where the link with insecticide resistance has been extensively studied, the induction of P450s is less well characterized in insecticide resistance. In another study by our group, the expression patterns of eight P450 genes, CYP4G13v1, CYP4D4v2, CYP4G2, CYP6A5, CYP6A5v2, CYP6A36, CYP6A37, CYP6A38, and CYP28B1 in response to permethrin treatment in resistant ALHF and susceptible CS and aabys house flies were characterization by treating 2-day old adult house flies with permethrin. Dose range, time course, and P450 gene induction assays showed the ALHF P450 genes to have a time- and dosedependent response to permethrin (Zhu et al. 2008b). Thus, a LD50 dose and a 24 h time interval were chosen for further induction studies of P450 genes in both resistant ALHF and susceptible CS and aabys strains. Three P450 genes, CYP4D4v2, CYP4G2, and CYP6A38, were co-up-regulated by permethrin treatment in permethrin resistant ALHF house flies (Zhu et al. 2008b). No significant induction in the expression of the three P450 genes was found in susceptible CS and aabys house flies that had either been treated with acetone alone or with permethrin solution in acetone compared with untreated house flies. Similarly, no significant induction was obtained in acetone treated ALHF house flies compared with their untreated counterparts. However, these three genes were induced at a variety of levels in permethrin treated ALHF house flies compared with untreated or acetone treated flies; there was a marked induction of CYP4D4v2 and CYP6A38 mRNA in permethrin treated ALHF house flies, whereas a low level of induction for CYP4G2 was detected in the permethrin treated ALHF house flies (Zhu et al. 2008b). These results suggest that unlike


253

some P450s, where constitutive expression may play an important role in insecticide resistance, CYP4D4v2, CYP6A38, and/or CYP4G2 may be unique features in ALHF in response to the insecticide exposure through induction of their expression, which in turn enhances their capacity to detoxify the insecticide and leads to enhanced insecticide resistance. Induction and/or constitutive overexpression of P450s are thought to be linked to the adaptation of insects to their environment (Terriere 1983, 1984). Further, in many cases increased levels of P450 gene expression have resulted in increased levels of both total P450s and the activities of those P450s, strongly suggesting this to be a major cause of insecticide resistance (Carino et al. 1994; Liu and Scott 1997, 1998; Kasai et al. 2000; Feyereisen 2005; Scott 2008). The identification of 2 P450 genes that were constitutively overexpressed in ALHF house flies (discussed in Section 4) and the 3 P450 genes in this section that were found to be co-up-regulated in response to permethrin exposure in a single resistant house fly strain, i.e., ALHF, suggest that both constitutive overexpression and induction mechanisms participate in increasing P450-mediated metabolic detoxification of permethrin in resistant ALHF house flies and imply the importance of these genes in the evolution of insecticide resistance.

14.6 Linkage resistance

analysis

of

P450

genes

and insecticide

Genetic linkage between an overexpressed P450 gene or protein and insecticide resistance is an important step in establishing a causal link between a P450 gene and its role in the development of resistance (Carino et al. 1994; Liu and Scott 1996; Rose et al. 1997; Guzov et al. 1998; Maitra et al. 2000; Feyereisen 2005). Our synergism studies and genetic studies have linked the PBO-suppressibleP450-mediated resistance in resistant ALHF house flies to autosomes 1, 2, and 5 (Liu and Yue 2001), and factors on autosome 5 are thought to play a major role in P450-mediated resistance. The factors or autosomes involved in the constitutive overexpression of CYP6A5v2 and CYP6A36 and permethrin induced expression of CYP4D4v2, CYP4G2 and CYP6A38 were further determine by our group using BC1 house fly lines (A2345, A1345, A1245, A1235, and A1234) generated by reciprocal crosses of resistant ALHF and susceptible aabys flies (Liu and Yue 2001). The genotype of BC1 for each line was homozygous for the recessive mutant allele from aabys and heterozygous for the dominant wild-type alleles from ALHF (Fig. 14.3). Genetic mapping of each of the constitutively overexpressed and permethrin induced house fly P450 genes, CYP6A5v2, CYP6A36, CYP4D4v2, CYP4G2 and CYP6A38, was conducted by allele specific PCR (Liu and Scott 1995) using the cDNA from BC1 house fly lines (Zhu and Liu 2008; Zhu et al. 2008a, b). Sequence comparison of these 5 P450 genes between ALHF and aabys revealed the nucleotide polymorphisms present in the coding region or the 5’ flanking region of the P450 genes. Based on the sequence differences at the polymorphism

254


Fig. 14.3 Diagrammatic representation of genetic isolation of BC1 house fly lines by crossing the resistant ALHF and susceptible aabys strains. These lines are named according to the autosomes bearing wild-type markers from ALHF (Liu and Yue, 2001).

sites in these genes between ALHF and aabys, specific primer pairs were designed for allele specific PCR reactions by placing single mismatched bases at the 3’ ends of primers that permitted preferential amplification of ALHF alleles over aabys alleles (Zhu and Liu 2008; Zhu et al. 2008a, b). The ALHF allele-specific primer sets for CYP4G2 amplified specific DNA fragments only in BC1 flies having the autosome 3 wild type marker from ALHF, showing that CYP4G2 is located on autosome 3. Our earlier study indicated that although factors on autosome 3 were very important in the overall level of permethrin resistance in ALHF house flies (Liu and Yue 2001), the resistance in ALHF governed by them was not suppressed by PBO. Since PBO appears not to be a perfect inhibitor for some of the P450s responsible for resistance, the product of CYP4G2 with a ~1.5-fold level of induction may be too small to be detected by a synergism study. Whereas, the ALHF allele-specific primer sets for CYP4D4v2, CYP6A5v2, CYP6A36, and CYP6A38 amplified specific DNA fragments only in BC1 flies having the autosome 5 wild type marker from ALHF. These findings indicate that the 4 P450 genes are located on autosome 5. Genetic studies have linked the P450-mediated resistance in ALHF to autosomes 1, 2, and 5 (Liu and Yue 2001). The correlation of genetic linkage of CYP4D4v2, CYP6A5v2, CYP6A36, and CYP6A38 on autosome 5 with the development of resistance and/ or P450-mediated resistance in ALHF suggests the functional importance of these 4 P450 genes in the increased detoxification of insecticides in ALHF.

14.7

Conclusions

Insect cytochrome P450s are known to play an important role in detoxifying insecticides and plant toxins, resulting in the development of resistance to


255

insecticides and facilitating the adaptation of insects to their plant hosts. A significant characteristic of insect P450s that is associated with enhanced metabolic detoxification of insecticides in insects is the constitutively increased levels of P450 proteins and P450 activities that result from constitutively transcriptional overexpression of P450 genes in insecticide resistant insects. Another interesting characteristic of insect P450 genes is that the expression of some P450 genes can be induced by exogenous and endogenous compounds, a phenomenon known as induction. The induction of CYP4D4v2, CYP4G2, and CYP6A38 and the constitutive overexpression of CYP6A5v2 and CYP6A36 only in resistant ALHF house flies, taken together with the close correlation between the production of these genes and the development of resistance and/or P450mediated resistance in ALHF, provide overwhelming evidence that P450 induction and constitutive overexpression are co-responsible for the detoxification of insecticides, evolutionary insecticide selection, and the ability of insects to adapt to changing environments. Acknowledgements Authors thank Dr. Julia W. Pridgeon and Jan Szechi for their valuable comments and editorial assistance. The studies performed by the author’s group were supported by the U.S. Department of Agriculture National Research Initiative (USDA-NRI) Competitive Grants Program; AAES Foundation Grants; Auburn University Biogrants; the Hatch Project ALA08-029; and the Department of Entomology and Plant Pathology, Auburn University.

References Berenbaum M R. Coumarins. // Rosenthal, G.A., Berenbaum, M.R. (Eds.). Herbivores: Their Interaction with Secondary Plant Metabolites. New York: Academic Press, 1991: 221–249. Carino F A, Koener J F, Plapp F W Jr, Feyereisen R. Constitutive overexpression of the cytochrome P450 gene CYP6A1 in a house fly strain with metabolic resistance to insecticides. Insect Biochem. Mol. Biol., 1994, 24, 411–418. Danielson P B, Foster J L, Cooper S K, Fogleman J C. Diversity of expressed cytochrome P450 genes in the adult Mediterranean fruit fly, Ceratitis capitata. Insect Mol. Biol., 1999, 8: 149–159. Daborn P J, Yen J L, Bogwitz M R, et al. A single P450 allele associated with insecticide resistance in Drosophila. Science, 2002, 297: 2253–2256. Feyereisen R. Insect cytochrome P450. // Gilbert, L.I., Iatrou, K., Gill, S.S. (Eds.). Comprehensive Molecular Insect Science. Oxford: Elsevier, 2005: 1–77. Fogleman J C, Danielson P B, MacLntyre R J. The molecular basis of adaptation in Drosophila: the role of cytochrome P450s. // Hecht, M., MacIntyre, R., Clegg, M. Evolutionary Biology, vol. 30. New York: Plenum, 1998: 15–77. Gilbert L I. Halloween genes encode P450 enzymes that mediate steroid hormone biosynthesis in Drosophila melanogaster. Mol. Cell Endocrinol., 2004, 215: 1–10. Greenberg B. Flies and disease. Sci. Am., 1965, 213: 92–99. Guzov V M, Unnithan G C, Chernogolov A A, Feyereisen R. CYP12A1, a mitochondrial cytochrome P450 from the house fly. Arch. Biochem. Biophys., 1998, 359: 231–240. Hodgson E. Mocrosomal mono-oxygenases. // Kerkut, G.A., Gilbert, L.I. (Eds.). Comprehensive Insect Physiology, Biochemistry and Pharmacology, vol. 11. Oxford: Pergamon, 1985: 225–321.

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Kasai S, Weerashinghe I S, Shono T, Yamakawa M. Molecular cloning, nucleotide sequence and gene expression of a cytochrome P450 (CYP6F1) from the pyrethroid-resistant mosquito, Culex quinquefasciatus Say. Insect Biochem. Mol. Biol., 2000, 30: 163–171. Keiding J. The house fly: biology and control. WHO Vector Control Series, 1986, 63. Keiding J. Review of the global status and recent development of insecticide resistance in field populations of the housefly, Musca domestica (Diptera: Muscidae). Bull. Entomol. Res., 1999, 89: S7–S67. Li W, Petersen R A, Schuler M A, Berenbaum M R. CYP6B cytochrome P450 monooxygenases from Papilio canadensis and Papilio glaucus: potential contributions of sequence divergence to host plant associations. Insect Mol. Biol., 2002, 11: 543–551. Liu N, Scott J G. Genetics of resistance to pyrethroid insecticides in the house fly, Musca domestica. Pestic. Biochem. Physiol., 1995, 52: 116–124. Liu N, Scott J G. Genetic analysis of factors controlling elevated cytochrome P450, CYP6D1, cytochrome b5, P450 reductase and monooxyenase activities in LPR house flies, Musca domestica. Biochem. Genet., 1996, 34: 133–148. Liu N, Scott J G. Inheritance of CYP6D1-mediated pyrethroid resistance in house fly (Diptera: Muscidae). J. Econ. Entomol., 1997, 90: 1478–1481. Liu N, Scott J G. Increased transcription of CYP6D1 causes cytochrome P450-mediated insecticide resistance in house fly. Insect Biochem. Mol. Biol., 1998, 28: 531–535. Liu N, Yue X. Insecticide resistance and cross-resistance in the house fly (Diptera: Muscidae). J. Econ. Entomol., 2000, 93: 1269–1275. Liu N, Yue X. Genetics of pyrethroid resistance in a strain (ALHF) of house flies (Diptera: Muscidae). Pestic. Biochem. Physiol., 2001, 70: 151–158. Liu N, Zhang L. Identification of two new cytochrome P450 genes and their 5’-flanking regions from the housefly, Musca domestica. Insect Biochem. Mol. Biol., 2002, 32: 755–764. Liu N, Zhang L. CYP4AB1, CYP4AS1 and Gp-9 overexpression in workers of the red imported fire ant, Solenopsis invicta. Gene, 2004, 327: 81–87. Maitra S, Dombrowski S M, Basu M, et al. Factors on the third chromosome affect the level of Cyp6a2 and Cyp6a8expression in Drosophila melanogaster. Gene, 2000, 248: 147–156. Niwa R, Matsuda T, Yoshiyama T, et al. CYP306A1, a cytochrome P450 enzyme, is essential for ecdysteroid biosynthesis in the prothoracic glands of Bombyx and Drosophila. J. Biol. Chem., 2004, 279: 35942–35949. Pittendrigh B, Aronstein K, Zinkovsky E, et al. Cytochrome P450 genes from Helicoverpa armigera: expression in a pyrethroidsusceptible and-resistant strain. Insect Biochem. Mol. Biol., 1997, 27: 507– 512. Reed J R, Vanderwel D, Choi S, et al. Unusual mechanism of hydrocarbon formation in the housefly: cytochrome P450 converts aldehyde to the sex pheromone component (Z)-9-tricosene and CO2. Proc. Natl. Acad. Sci. USA, 1994, 91: 10000–10004. Rose R L, Goh D, Thompson D M, et al. Cytochrome P450 (CYP) 9A1 in Heliothis virescens: the first member of a new CYP family. Insect Biochem. Mol. Biol., 1997, 27: 605–615. Scott H G, Lettig K S. Flies of public health importance and their control. US Government Printing Office, Washington, DC, 1962. Schuler M A. The role of cytochrome P450 monooxygenases in plant-insect interactions. Plant Physiol., 1996, 112: 1411–1419.


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Scott J A, Collins F H, Feyereisen R. Diversity of cytochrome P450 genes in the mosquito, Anopheles albimanus. Biochem. Biophys. Res. Commun., 1994, 205: 1452–1459. Scott J G. Cytochromes P450 and insecticide resistance. Insect Biochem. Mol. Biol., 1999, 29: 757– 777. Scott J G. Insect Cytochrome P450s: thinking beyond detoxification. // Liu, N. Recent Advances in Insect Physiology, Toxicology and Molecular Biology. Kerala: Research Signpost, 2008: 117– 124. Scott J G, Liu N, Kristensen M, Clark A G. A case for sequencing the genome of the house fly, Musca domestica (Diptera: Muscidae). J. Med. Entomol., 2009, 46: 175–82. Scott J G, Liu N, Wen Z. Insect cytochrome P450: diversity, insecticide resistance and tolerance to plant toxins. Comp. Biochem. Physiol., 1998, 121C: 147–155. Sutherland T D, Unnithan G C, Andersen J F, et al. A cytochrome P450 terpenoid hydroxylase linked to the suppression of insect juvenile hormone synthesis. Proc. Natl. Acad. Sci. USA, 1998, 95: 12884–12889. Snyder M J, Scott J A, Andersen J F, Feyereisen R. Sampling P450 diversity by cloning polymerase chain reaction products obtained with degenerate primers. Methods Enzymol., 1996, 272: 304– 312. Terriere L C. Enzyme induction, gene amplification, and insect resistance to insecticides. // Georghiou, G.P., Saito, T. Pest Resistance to Pesticides. New York: Plenum Press, 1983: 265– 297. Terriere L C. Induction of detoxication enzymes in insects. Ann. Rev. Entomol., 1984, 29: 71–88. Tomita T, Scott J G. cDNA and deduced protein sequence of CYP6D1: the putative gene for a cytochrome P450 responsible for pyrethroid resistance in house fly. Insect Biochem. Mol. Biol., 1995, 25: 275–283. Wen Z, Pan L, Berenbaum M B, Schuler M A. Metabolism of linear and angular furanocoumarins by Papilio polyxenes CYP6B1 co-expressed with NADPH cytochrome P450 reductase. Insect Biochem. Mol. Biol., 2003, 33: 937–947. Winter J, Bilbe G, Richener H, et al. Cloning of a cDNA encoding a novel cytochrome P450 from the insect Locusta migratoria: CYP6H1, a putative ecdysone 20- hydroxylase. Biochem. Biophys. Res. Commun., 1999, 259: 305–310. Zhu F, Liu N. Differential expression of CYP6A5 and CYP6A5v2 in pyrethroid-resistant house flies, Musca domestica. Arch. Insect Biochem. Physiol., 2008, 67: 107–119. Zhu F, Feng J, Zhang L, Liu N. Characterization of two novel cytochrome P450 genes in insecticideresistant house-flies. Insect Mol. Biology, 2008a, 17: 27–37. Zhu F, Li T, Zhang L, Liu N. Co-up-regulation of three P450 genes in response to permethrin exposure in permethrin resistant house flies, Musca domestica. BMC Physiol., 2008b, 8: 18.

CHAPTER 15

Metamorphosis of Cisgenic Insect Resistance Research in the Transgenic Crop Era Xinzhi Ni, Xianchun Li, Yigen Chen, Fuzhen Guo, Jinian Feng and Huiyan Zhao

Abstract The biotechnological revolution has forever changed agricultural research and crop production worldwide. Commercial agriculture now includes plants that produce enhanced yield and quality, survival in hostile environmental conditions, manufacture and express defensive toxins, and yield grains with greatly enhanced nutrient fortification. Unprecedented opportunities and ethical challenges created by this new technology are transforming the field of plant protection may be compared with the dramatic changes of an insect undergoes during metamorphosis. In particular, the technology has brought enormous exciting opportunities as well as great challenges that are transforming the research fields of innate (or cisgenic) insect resistance and insect-host plant interactions. Here, we provide a review of the historic milestones in both cisgenic (insertion of genes from sexually compatible stock) and transgenic (insertion of genes from sexually incompatible stock) crop production in agriculture. Biological significance of transgenic technology and the critical role of basic research in crop and human protection are discussed. In addition, three research frontiers that utilize basic and applied research approaches and synergistically combine cisgenes with transgenic technology for broad crop protection (including both yield and quality improvements) are also discussed.

Xinzhi Ni Crop Genetics and Breeding Research Unit, USDA-ARS, Tifton, GA, 31793, USA E-mail: [email protected] Xianchun Li Department of Entomology and BIO5 Institute, University of Arizona, Tucson, AZ, 85721, USA Yigen Chen Department of Entomology, Michigan State University, East Lansing, MI, 48824, USA Fuzhen Guo, Jinian Feng, Huiyan Zhao College of Plant Protection, Northwest A&F University, Yangling, Shaanxi, 712100, China


Metamorphosis of Cisgenic Insect Resistance Research in the Transgenic Crop Era

259

Keywords cisgenic insect resistance, insect susceptibility, transgenic crop, metamorphosis, biotechnology

15.1

Introduction

Although the progress in agricultural science and technology has come in leaps and bounds in recent centuries, insects, diseases, and weeds are still formidable impediments to consistent crop yield and quality. The key pests and the goals for crop protection are intimately associated with other significant agricultural development, such as adoption of farming machinery, introduction of new crop hybrids, and practice of integrated pest management (IPM) worldwide. Integration of less intrusive crop protection tactics with development and availability of new crop genotypes have contributed significantly to utilization of innate crop traits to reduce insect damage and maximize crop yield and quality. The crop plants developed utilizing innate crop genes have also been known as cisgenic crops (Schouten et al. 2006; Schouten and Jacobsen 2008). Cisgenic crop plants are defined as plants containing only genes from within the same species or from sexually compatible species, and plant cisgenes have existed for thousands of years and have been utilized by traditional plant breeders (Schouten and Jacobsen 2008). In contrast, Schouten and Jacobsen (2008) define intragenesis as a novel combination of functional parts of genes from a gene pool otherwise inaccessible to the species to be modified. The novel combinations do not exist in nature, nor is likely that they would arise in nature, or as result of traditional breeding. For example, RNA interference (RNAi) constructs belong to intragenesis, not to cisgenesis. Nevertheless, Schouten and Jacobsen (2008) describe cisgenesis and intragensis as sister approaches in innovative plant breeding. Although the definitions of cisgenesis (natural breeding in general), intragenesis (breeding within a genus either naturally or artificially), and famigenesis (breeding within a family) are still being debated at present (see Schouten and Jacobsen 2008, and Rommens 2008 for details), we will use the definitions by Schouten and Jacobsen (2008) throughout this chapter. One of the most dramatic recent advancements in host plant resistance and crop protection in general was the birth of pest-resistant transgenic corn and cotton expressing insecticidal proteins from the bacterium, Bacillus thuringiensis (Bt) Berliner, in the 1980’s (Patlak 1998). While ordinary domesticated crops depend solely on limited cisgenic traits for their protection against insect pests and diseases, transgenic crops possess cisgenic and/or intragenic insect resistance, along with transgenes from unrelated organisms to protect from insect pests, diseases, and herbicides. Because of their general safety and specificity, a variety of non-native genes that confer insect, disease, and/or herbicide resistance, and other favorable quality enhancement traits have been successfully incorporated into major agricultural crop plants to improve the efficiency of agricultural production (Graff and Bradford 2005). Development and deployment of transgenic crops have increased rapidly

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Xinzhi Ni et al.

worldwide. Since the concept of transgenic crop was initiated in the 1980’s, and since 1996 when the transgenic crops were deployed, 398 transgenic specialty crops have been developed in 14 crop categories in the U.S. (Graff and Bradford 2005). Like other great advancements in agriculture, medicine, and other scientific disciplines, this unprecedented revolutionary breakthrough in crop breeding simultaneously and paradoxically presents us with enormous opportunities and great ecological and sociological challenges. Just as metamorphosis transforms life stages of an insect, introduction of transgenic crop is rapidly transforming the fields of cisgenic insect resistance and crop breeding, and ultimately across the disciplines of biology in general. To embrace the opportunities and meet the challenges presented by Bt and other transgenic crops, we have to learn from the history to devise appropriate strategic plans for the challenges of the expanding future. In this chapter, we intend to: 1) provide a brief review of cisgenic insect resistance research to emphasize the achievements of this discipline in agriculture; 2) highlight the milestones of multidisciplinary research advancements that have contributed to transgenic Btcrop development to demonstrate the importance of long-term basic research; 3) discuss the significance of transgenic crop introduction in biological research to further emphasize the role of long-term basic research in agricultural and medical research; and 4) outline three new research frontiers that synergistically combine basic and applied research strategies to solve practical insect problems and mycotoxin reduction in the transgenic crop era.

15.2

Historical milestones in cisgenic insect resistance research

Insect-resistant cultivars have been developed in most, if not all crops worldwide and they have had a vast positive impact. For example, the economic value of resistance against insect pests of wheat is estimated at over US$250 million annually (Smith et al. 1999). The identification and planting of the Hessian fly [Mayetiola destructor (Say) (Diptera: Cecidomyiidae)] resistant wheat cultivar ‘Underhill’ was noted as early as in 1782 by Havens (Smith 1989; Panda and Khush 1995). Three best-known early crop-insect systems that successfully demonstrated the essential role of host plant resistance covering multiple agricultural crops were: wheat-Hessian fly in 1781, apple-woolly aphid [Eriosoma lanigerum (Hausmann) (Homoptera: Aphididae)] in 1831, and grape-grape phylloxera [Daktulosphaira (Phylloxera) vitifoliae (Fitch)] in 1890 (Panda and Khush 1995) (Table 15.1). In addition to the impact on food supplies and economics at the time, these systems had a remarkable impact on multiple agricultural research disciplines, such as, entomology, agronomy, horticulture, and viticulture. A series of Hessian fly-resistant wheat cultivars has secured the wheat production and critical food supplies ever since, and the successful research on the Hessian fly has been of worldwide importance in the history of wheat production. Similarly, apple woolly aphid and grape phylloxera resistance have been of great importance in the history of horticultural crops. Grape

Metamorphosis of Cisgenic Insect Resistance Research in the Transgenic Crop Era Table 15.1 Year

261

Milestones in Cisgenic Insect Resistance Research Event

Reference

ca 1776

Hessian fly is reported as an important wheat pest in the Northeastern USA

Painter (1951)

1782

Hessian fly-resistant wheat cultivar ‘Underhill’ is reported in the U.S.

Panda and Khush (1995)

1785

Wheat resistance to the Hessian fly is discussed

1817

C. V. Riley reported that sorghum is more resistant to grasshopper than corn


1831

Apple cultivar ‘Winter Majetin’ resistance to woolly aphid is utilized in the U.K.


1861

Introduction of grape phylloxera in France


1871

The Hessian fly caused $5-25 million loss in Kansas


1890

Successfully control of grape phylloxera by grafting grapes with American rootstocks


1900’s

Selection of ‘U4’ cotton for leafhopper resistance in Africa


1905

Sir Roland Biffen has speculated and demonstrated wheat resistance to rust infection is genetically inherited

Patlak (1998)

1951

The book Insect Resistance in Crop Plants by Painter is published

Smith (1989)

1964

Over 60 wheat cultivars resistant to the Hessian fly have been developed and grown on 8.5 million acres in 34 states of the U.S.


1974

Over 30 wheat cultivars resistant to 9 Hessian fly biotypes have released and planted on 20 million acres in the U.S.


1995

Russian wheat aphid-resistant wheat cultivar ‘Halt’ is released in the U.S. after the discovery of the aphid in 1986 in Texas and western states of the U.S.

Clement and Quisenberry (1999)

Painter (1951)

phylloxera has been a formidable pest worldwide not only in the history of entomology but also in viticulture and oenology. The plague of D. vitifoliae on grapevines can be traced back to 1850 in Europe (Granett et al. 2001). American grape cultivars were resistant to this native pest, while the French grape cultivars were extremely susceptible. The French wine industry was devastated by 1884. However, the subsequent grafting of rootstocks from the United States to French scions saved the French wine industry (Panda and Khush 1995). Some California vineyards (at least 20,000 ha) still suffer injury from D. vitifoliae. Costs for clearing, fumigating, and replanting these vineyards have been estimated as high as US$2 billion (Hill 1997; Granett et al. 2001; Quisenberry and Ni 2007). A number of historical successes in developing new cereal cultivars for insect resistance have derived from extensive field screenings. Porter et al. (1999) reported that after the screening of over 23,000 barley germplasm entries, 121 of

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them showed resistance to the Russian wheat aphid, Diuraphis noxia (Mordvilko) (Hemiptera: Aphididae), and 23 of over 8,800 barley germplasm entries have been identified as resistant to cereal leaf beetle, Oulema melanopus (L.) (Coleoptera: Chrysomelidae), and 33 of 5,100 barley germplasm entries were also identified as Hessian fly resistant. The entries used for wheat screenings at various research laboratories through the last century could be much higher than the number of entries used for barley screenings. The wide utilization of over 60 Hessian fly-resistant wheat cultivars has resulted in the removal of the Hessian fly from the list of major insect pests in wheat production in the U.S. (Panda and Khush 1995). A number of other crop cultivars have been developed to reduce the damage of pests arising from new introductions, resurgence, or development of new biotypes. A good example focusing on an introduced insect pest is the development and release of new wheat cultivar ‘Halt’ in 1995, and other cultivars that confer resistance to the Russian wheat aphid [Diuraphis noxia (Mordvilko) (Hemiptera: Aphididae)]. The research efforts of multiple institutions on host plant resistance have effectively reduced wheat crop losses caused by the Russian wheat aphid in the western region of the U.S., especially in the Great Plains region (Quisenberry and Peairs 1998; Clement and Quisenberry 1999). Biotype development of more virulent insect pests is one of the most important and dynamic challenges for the insect-resistant cultivar development. In wheat pests, 11 biotypes (A-K) have been identified for the greenbug (Burd and Porter 2006), 9 for the Hessian fly (Panda and Khush 1995), and 5 for the Russian wheat aphid (Shufran and Payton 2009). A number of wheat cultivars have been developed against various biotypes of insect pests. For example, over 30 wheat cultivars have been released against various Hessian fly biotypes (Panda and Khush 1995). These great recent successes of host plant resistance to insects have shared the same origins of massive screens of available germplasm once a new or emerging pest biotype outbreak occurs. The treatise by Painter (1951) is considered as a historical monument for the research field of host plant resistance to insects, although the utilization of insectresistant crop cultivars started long before publication of Painter’s book as indicated in Table 15.1. A renewal of interest in host plant resistance in the 1950’s was motivated by the ever-increasing insecticide resistance of insect pests and negative ecological impact of synthetic insecticides on agricultural and natural ecosystems. Seven books on host plant resistance to insect pests have been published since 1951, which have documented the chronological advancements of host plant resistance research in detail. A subsequent book Breeding Plants Resistant to Insects by Maxwell and Jennings (1980), has provided a thorough review of host plant resistance concept and summarized the accomplishments on eight crops in agriculture and forestry ecosystems. Three later books (Smith 1989; Smith et al. 1994; and Smith 2005) documented the dynamic changes in the research technologies before and after the introduction of transgenic Bt crops worldwide. Two additional books were published in the 1990’s: Host Plant


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Resistance to Insects by Panda and Khush (1995), and Global Plant Genetic Resources for Insect-Resistant Crops by Clement and Quisenberry (1999). These two books have documented global research advancement and genetic resources related to insect resistance in major crops. Contrary to the previous publications that only covered agricultural crops, a book by Fritz and Simms (1992) entitled Plant Resistance to Herbivores and Pathogens – Ecology, Evolution, and Genetics presented a broad ecological perspective on host plant resistance to insect herbivores and phytopathogens in both managed and natural ecosystems. These are all valuable intellectual resources for understanding and utilizing crop plant cisgenes as well as transgenes to improve yield and quality protection of agricultural crops in the coming years.

15.3

Historical highlights in transgenic crop development

While the development of insect-resistant crop cultivars followed the effective centuries old repetitive pattern of massive screening for insect resistance among diverse crop genetic resources (Table 15.1), the history of transgenic crop development is comparatively short. According to the Merriam-Webster Dictionary, the concept of the transgenic crop production was first being recognized only less than three decades ago with the coinage of the new word “transgenic” in biological lexicons in 1982 (http://www.merriam-webster.com/). However, the fundamental research in multiple disciplines related to the concept was initiated long before the arrival of transgenic crops. The critical basic research related to the development of Bt-transgenic crops in entomology can be traced back to as early as the beginning of the 20th century when the insecticidal property of B. thuringiensis was discovered in 1901 (Tanada and Kaya 1993). In addition to advances in crop genetics and breeding, the development of Bttransgenic crops was the result of the century-long etiological research from entomology and plant pathology, [i.e., the identification of δ-endotoxin genes in B. thuringiensis as the insecticidal factor in entomology (Tanada and Kaya 1993; Patlak 1998)], and the discovery of the transferred DNA (T-DNA) region of the tumor-inducing (Ti) plasmid of Agrobacterium tumefaciens (At) in the nuclear genome of host plant cells (Escobar and Dandekar 2003). We herein intend to highlight the milestones in the meandering history of etiological research on Bt and At, respectively, which has led to the conceptual formation of Bt transgenic crop in crop protection. 15.3.1

Breakthroughs in Bacillus thuringiensis research

The discovery and extensive work on the insecticidal toxins produced by the entomopathogenic bacterium B. thuringiensis comprise one of the critical milestones in transgenic crop development. Key events are summarized in Table 15.2. The renowned microbiologist Luis Pasteur might have been one of the first researchers who encountered B. thuringiensis infection during his study

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of the silkworm diseases between 1865 and 1870 (Steinhaus 1960; Tanada and Kaya 1993). However, the entomopathogenic bacterium was not isolated from the silkworm until 1901 by Ishiwata in Japan (Table 15.2). Formal isolation of the bacterium did not occur until 1911 when Berliner in Germany isolated it from the Mediterranean flour moth, Anagasta kuehniella (Zeller) (Lepidoptera: Pyralidae). He named the bacterium after the provincial name Thuringia, Germany, where the infested flour was obtained for his research. It is worth noting that although Berliner and Mattes examined the bacterium extensively afterwards in the early 1920’s, neither of them was able to correlate high pathogenicity with the toxin production at that time. The identification of the proteinous Bt-toxin did not occur until the 1950’s (Tanada and Kaya 1993) (Table 15.2). Table 15.2 Date

Milestones in Bacillus thuringiensis Research Event

Reference

1860’s

Luis Pasteur encountered Bacillus thuringiensis (Bt) Steinhaus (1960) infection in silkworm rearing facilities Tanada and Kaya (1993)

1901

Bacteriologist Ishiwata Shigetane describes a bacterial Tanada and Kaya (1993) disease outbreak in silkworms as a new species of bacteria Patlak (1998)

1907

Description of Bt on Mediterranean flour moth by Berliner Tanada and Kaya (1993)

1927

Mattes re-isolates Bt and confirms Berliner’s description on Tanada and Kaya (1993) high pathogenicity

1938

Commercial use of Bt insecticide in France

1942

After having obtained a culture of Bt originally established Tanada and Kaya (1993) by Mattes, Steinhaus has demonstrated the potential of Bt as microbial insecticide in the North America

1953

Hannay describes the parasporal body staining properties, Tanada and Kaya (1993) and soluability in alkali, but not in organic solvent

1955

Hannay and Fitz-James describe that the proteinous of the Tanada and Kaya (1993) crystal has at least 17 amino acids

1954–1956

Angus published three reports describing the parasporal Tanada and Kaya (1993) body as the source of Bt toxin, that confirms the report by Japanese scientists

1981

Helen Whiteley and Ernest Schnepf (University of Washington) have cloned a Bt toxin gene

Patlak (1998) Frankenhuyzen (2009)

2009

174 holotype Bt toxin genes belonging to 55 Cry and 2 Cyt families have been tested against 163 species of insects (mainly from Lepidoptera, Coleoptera, and Diptera)

Frankenhuyzen (2009)

Patlak (1998)

The first field trials with Bt were conducted against the European corn borer, Ostrinia nubilalis (Hübner) [Lepidoptera: Pyralidae (Crambidae)] in 1929, and the first commercialized insecticide containing Bt was sold in 1938 in France (Tanada and Kaya 1993; Patlak 1998). In North America, Steinhaus recognized the significance of Bt and the value in commercializing it as a biological control agent for agricultural pests in 1942, a decade before the protein toxins, most


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notably δ-endotoxin, were identified (Tanada and Kaya 1993) (Table 15.2). Because major insect pests were developing insecticide resistance to synthetic insecticides in the 1950’s, Bt insecticides garnered increasing interest because of their narrow spectrum of insect targets (Steinhaus 1960). Further details of chronological milestones in research on B. thuringiensis, particularly the δendotoxin, can be found in five reviews published in the 1980’s (Carlton and González 1985; Aronson et al. 1986; Whiteley and Schnepf 1986; Andrews et al. 1987; and Höfte and Whiteley 1989). Frankenhuyzen (2009) recently reviewed the research on molecular genetics of Bt toxin genes. In total, 174 Bt toxin genes (from 5 Cry and 2 Cyt families) that affect 163 insect species in the orders Lepidoptera, Coleoptera, and Diptera have been examined (Frankenhuyzen 2009) (Table 15.2). 15.3.2

Milestones in Agrobacterium tumefaciens research

While scientists in the entomological field were making steady progress on Bt research, gradual progress was being made with phytopathological research on the crown gall disease on crop plants, including fruit trees, grape vines, and berry canes (Patlak 1998; Stafford 2000). The most significant initial progress on the crown gall disease was the identification of the bacterium Agrobacterium tumefaciens (or At) (E. F. Smith & Townsend) as the etiological factor for this disease in 1907 (Escobar and Dandekar 2003) (Table 15.3). The discovery that A. tumefaciens inflicts prolific growth of the infected plant cells to form a tumor was contradictory to the scientific dogma at that time- all bacterial phytopathogens were assumed to cause plant tissues to cease growth, e.g., death, wilt, or discoloration. No further research was conducted on At until 40 years later when Armin Braun of the Rockefeller Institute for Medical Research examined the underlying mechanisms of the prolific growth on the stems caused by this soilborne bacterium (Patlak 1998; Escobar and Dandekar 2003) (Table 15.3). Braun then conducted pioneering research to demonstrate that plant cells were permanently programmed into tumor cells by the tumor-inducing principle (TIP) introduced by the pathogen-A. tumefaciens (Patlak 1998; Escobar and Dandekar 2003). In the 1950s and 1960s, scientists made numerous breakthroughs toward understanding DNA and its role in genetics of all living organisms. Armin Braun started by searching for the bacterium’s DNA for evidence of a TIP. It turned out that the TIP did not reside in the chromosome equivalent structure in the bacterium, but in a large tumor-inducing (Ti) plasmid that was identified in 1974 by Jeff Schell and Marc van Montagu (Patlak 1998; Escobar and Dandekar 2003) (Table 15.3). The identification of the transferred DNA (T-DNA) region of the Tiplasmid to the chromosomes of crown gall tumor cells was reported in 1977 by microbiologists Eugene Nester, Milton Gordon, and Mary-Dell Chilton from the University of Washington. By the end of the 1970’s, it was clearly established that crown galls were caused by the transfer of the T-DNA into the nuclear DNA of plant cells (Escobar and Dandekar 2003) (Table 15.3). In the late 1970’s and

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Table 15.3 Year

Milestones in Etiological Research on the Crown Gall Disease Event

Reference

1850’s

Discovery of crown gall disease in grapevines

Stafford (2000) Escobar and Dandekar (2003)

1907

Erwin Smith and C. O. Townsend from USDA-ARS discover that the cause of crown gall in Paris daisy is a bacterium called Agrobacterium tumefaciens (At)

Patlak (1998) Zupan et al. (2000) Escobar and Dandekar (2003)

1947

Armin Braun shows that A. tumefaciens introduces a tumor-inducing principle (TIP) into plant cells that permanently transforms the plant cells into tumor cells


1956–1971

Opines (nitrogenous compound with low molecular weight) are identified exclusively in tumor cells; At loses its virulence at 37°C; and the TIP is transferable between virulent and non-virulent At

Escobar and Dandekar (2003)

1974

The discovery by Jeff Schell and Marc van Montagu, University of Gent, Belgium, has identified the tumor-inducing (Ti) plasmid as the TIP


1977

Eugene Nester, Milton Gordon, and Mary-Dell Chilton further identified that the transferred DNA (T-DNA) region] on the Ti plasmid is the TIP


1983

First plant is transformed with a recombinant gene using At as a vector


1984

T-DNA oncogenes that mediate overproduction of auxin and cytokinin are identified


1985–2003

vir-gene regulatory system identification; identification of plant genes involved in At transformation; extension of At host range for transforming monocots; and At (C58) genome sequence


early 1980’s, it became possible to remove tumor-inducing T-DNA of the Tiplasmid and introduce desirable foreign genes into crop plant genomes using At as a vector. At the same time, new techniques were developed to allow researchers to manipulate DNA in a variety of ways to meet their preselected specifications. Much more extensive research has also been conducted on molecular genetics of At and genome sequencing during the last decade (Escobar and Dandekar 2003) (Table 15.3). These advancements and new technologies have allowed scientists to efficiently insert foreign genes into plant genomes. Detailed information about historical events in At research as a phytopathogen and a vector of genetic transformation can be found in two reviews by Zupan et al. (2000) and Escobar and Dandekar (2003). 15.3.3

Concept formation and expansion for transgenic crops

Developing transgenic organisms was a nascent event in the 1980’s (Patlak 1998) (Table 15.4) compared to the more protracted century-long meandering history of

Metamorphosis of Cisgenic Insect Resistance Research in the Transgenic Crop Era Table 15.4

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Historical Highlights in Transgenic Crop Development

Year

Event

Reference

1982

Introduction of “transgenic” in Merriam-Webster Dictionary

http://www.merriam-webster.com/

1983

First At plasmid vectors are developed

Patlak (1998)

1983

Scientists at Monsanto introduce foreign genes into plants by using A. tumefaciens plasmid vectors

Patlak (1998)

1983

Kary Mullis and colleagues have developed and patented polymerase chain reaction (PCR) method for in vitro amplification of DNA; subsequently, Mullis and Michael Smith won the Nobel Prize in 1993

Rabinow (1996)

Mid 1980’s

A number of scientists have proposed that the previous advancements could be combined to modify plants, and develop self-protecting plants

Patlak (1998)

1986

Roger Beachy shows that plants bioengineered to produce a viral coat protein are protected from infection with the virus

Patlak (1998)

1990

Field trials show that transgenic Bt-cotton cultivars confer resistance to cotton bollworm and tobacco budworm

Patlak (1998)

1996

Genetically-engineered virus-resistant squash seeds reached the market

Patlak (1998)

1996

Bt cotton reached the market

Patlak (1998)

1998

Herbicide-resistant soybean, cotton, and canola cultivars, and corn hybrids reached the market

Patlak (1998)

2005

398 specialty transgenic crops developed in 14 crop categories in the U.S. alone

Graff and Bradford (2005)

2009

Transgenic corn with fortified multiple vitamins is developed

Naqvia et al. (2009)

At and Bt. Transgenic advancements have been made relatively rapidly in 14 agricultural crops, involving at least 398 transgenic events have been added to 14 agricultural crops (Graff and Bradford 2005) (Table 15.4). Two factors have contributed to the unprecedented progress in transgenic crop research and development in recent decades. First, public interest and demand for improving crop yield and/or quality has increased exponentially since the establishment of the concept of transgenesis in 1982 as shown in Table 15.4. Second, previous basic and applied research achievements in crop protection set the stage for this technology to become a reality. The century-long basic research at both private and public institutions should not be overlooked, although the key recent technological innovations have been contributed mainly by private companies such as Monsanto Company (Patlak 1998) (Table 15.4). Two of the latest reports describe new transgenic corn with fortified nutrients (i.e., multivitamin levels) targeted for developing African countries (Zhu et al. 2008; Naqvia et al. 2009).

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This is another significant expansion of utilizing transgenic technology to improve crop nutritional quality, which is far beyond the development of pestprotected transgenic Bt crops first deployed over a decade ago. Transgenic technology has permitted these extraordinary achievements in such a short period of time – from self-protecting (i.e., Bt) to quality improving (i.e., nutrientenriched) transgenic crop development. Many intensive research programs are also examining and developing transgenic animals (e.g., fish and livestocks) (Liu et al. 1990; Naylor and Burke 2005; Miller 2008), edible vaccines (Pascual 2007), and biofuel production using transgenic crop and/or plants (Sticklen 2008). In contrast to the crops and animals, the progress in the other extreme of the biological gamut –the microbial world is also exciting. The most recent development of two bioengineered species of yeast (fission yeast and baker’s yeast) to artificially produce vanillin (the dominant flavor compound of vanilla) is truly refreshing (Li and Rosazza 2000; Röling et al. 2001; Hansen et al. 2009). The diversity and intensity of current transgenic crop research are still increasing rapidly, which strongly suggest that the tremendous value of transgenic crops are far from being fully realized.

15.4

Significance of transgenic crop development

15.4.1

Conceptual revolution in biology

Crop protection has enjoyed a long and rich history accompanying human civilization for at least 6,000 years (Chou 1990). However, transgenic Bt crop development was the first time that human manipulations have introduced a foreign gene that profoundly altered the genetic makeup of a crop plant, and instantly extends the definition of crop breeding from inter- and/or intra-species genetic improvements to inter-phylum genetic modification. While the traditional crop breeding methods have always focused on selection of intra- and interspecies variations, and never manipulated inter-phylum gene flow, the transgenic crop is essentially a recombination of genetic makeup of two unrelated organisms from different phyla for the purpose of improving yield and/or quality of the crops. We believe that the transgenic Bt crops with pest-protected traits represent the dawn of a new era of agricultural production with unprecedented opportunities. It is truly exciting to see that, in addition to a number of crops that have been developed against insects and diseases, many valuable plant traits could be manipulated with transgenes, ranging from salt and drought tolerance in cucumber and rice (Graff and Bradford 2005) to fortified multiple vitamins in corn (Naqvia et al. 2009). Furthermore, the development of transgenic animals, microbes, edible vaccines, and biofuel production, as indicated in the previous section, is also exciting for both agricultural and medical research purposes.


15.4.2

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Critical role of long-term basic research

The development of Bt transgenic crops is definitely a historical opportunity for us to compare crop and human protection in term of utilizing opposite research approaches. Of many realistic factors, low financial support for agriculture and ethic challenges for medical research, the two disciplines have long been known to utilize paradoxical (i.e., basic versus applied) research approaches to solve practical problems. While medical research is designed to find cures to human diseases, and save everyone in human populations, host plant resistance has been focused on selecting individuals in a population that confers resistance to pests. Such contradictory tactics have achieved similar remarkable successes, and established a series of milestones in the history of crop and human protection research. In general, development of insect-resistant crop cultivars starts from the massive screenings of crop germplasm against a given insect pest, while the treatment of a human disease usually starts from basic etiological research. The basic research (e.g., etiology) in medical history has repeatedly enabled man to achieve scientific triumphs from medical tragedies. For instance, the first report of highly pathogenic avian influenza (HPAI) occurred in 1878, but HPAI was not separated from fowl cholera until two years later (Lupiani and Reddy 2009). Although many early scientists believed HPAI was caused by a Bacillus spp., the pathogen was identified as a virus in 1901 (Bruce 1890; Lupiani and Reddy 2009). Such misunderstanding was caused by the limited scientific knowledge of microbiology at the time. HPAI spread between 1901 and the1930’s and was responsible for the major human pandemic in 1918 that killed 100 million people (Barry 2005). Although recognition that influenza was caused by a virus occurred in 1901, the first influenza virus was not isolated for another 30 years (Lupiani and Reddy 2009). The basic research work in the 20th century has thus been focused on a variety of influenza virus strains and their detailed basic biological characteristics that are critical for vaccine research and development (Lupiani and Reddy 2009). A similar historical path of etiological research on human malaria has also been described by Prescott et al. (1993). The name Mal’aria originates from the 1600’s and is translates from Italian to mean “bad air.” The name suggested that the disease was associated with the bad smell of the swamps near Rome (Prescott et al. 1993). The etiological breakthrough did not occur until the 1800’s by the identification of Plasmodium in blood and mosquito vectors (Prescott et al. 1993). Basic research on influenza, human malaria, and small pox in the 19th and early 20th centruries are similar to 21st century challenges like global warming, bioterrorism, and biosecurity (Pennington 2003; Khardori 2006; Mills et al. 2008). Science and medicine have synergistically merged together because scientific principles and methodologies are an essential part of routine clinical practice, diagnostics and therapy. It is common practice and imperative to identify the causal agent of a new disease to design effective treatments and vaccinations and other preventative protocols, by restrictively following the century-old

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Koch-Henle postulates (or Koch’s postulates in short) (Inglis 2007). As technology advances, new state-of-art test of causality test of diseases, such as “molecular Koch’s postulates” still need to be finalized (Inglis 2007). From a historical perspective, the above cited research examples based on human influenza, human malaria, and crown gall diseases of plants demonstrate that basic etiological research is critically important in long term, but intricate and laborious in short term. We hope that our parallel comparison between human and crop protection (in particular, host plant resistance research) can stimulate the interest in synergistically utilizing basic and applied research in the field of crop protection, in particular, developing new crop cultivars with both cisgenes and transgenes that confer resistance to insect pests.

15.5

Metamorphosis of host plant resistance research

The introductions of transgenic crops is transforming crop breeding for host plant resistance to insects, diseases, and other research disciplines of agriculture. Given the relatively short period since the introduction of the Bt transgenic cotton crop into agricultural ecosystems, the long-term impact of the introduction of Bt transgenic crops is not yet clear. Continued long-term ecological monitoring of risks and benefits on large scales are imperative for the field of crop protection, although an array of important ecological questions (such as target insect resistance to Bt toxin, effect of Bt on non-target organisms, and on tri-trophic interactions) have become the focus of significant research efforts in recent years. Since Losey et al. (1999) reported the effects of Bt-corn pollen ingestion on larvae of a non-target organism, the monarch butterfly, Danaus plexippus (L.) (Lepidoptera: Danaidae), several reports have examined the impact of transgenic crops on non-target organisms, including arthropod natural enemies (Wolfenbarger and Phifer 2000; Hellmich et al. 2001; Zangerl et al. 2001; Romeis et al. 2006; Marvier et al. 2007; Chen et al. 2008; Wolfenbarger et al. 2008; Lövei et al. 2009). The British Farm Scale Evaluations have reported that in transgenic oilseed rape and beet crop fields, biological diversity indices were lower than those in conventional varieties (Giles 2003). The evaluations examined the weeds and their seeds in crop fields, and insects, birds, and small animals that feed on these plant populations. Giles (2003) also suggested that the reduced biodiversity in weeds, seeds and insects in the farmlands could have a “knock-on effect” (also known as bottom-up effect) on birds and small animals feeding off these populations. A review by Pilson and Prendeville (2004) discussed the potential consequences of transgene escape on ecosystems in general. In addition, the development of Bt-endotoxin resistance in insect pests has been examined extensively in recent years. Reports by Tabashnik et al. (2008) and Gassmann et al. (2009) summarized the latest developments in this research area. After examining the monitoring data for the Cry1A toxin resistance in H. zea and other pests over a decade, Tabashnik et al. (2008) concluded that the refuge strategy is working in delaying Bt resistance in major agricultural crops. In addition, the


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fitness cost of Bt-resistant insects in the absence of Bt-toxins demonstrates the importance of refuges in delaying of Bt resistance in insects (Gassmann et al. 2009). To address the synergism between basic and applied research approaches in crop protection, numerous research opportunities are emerging to integrate transgenes and cisgenes and sustain the fidelity of highly efficient of agricultural production in the transgenic crop era worldwide. Three stimulating research frontiers with great potential are highlighted in this section to combine basic and applied research synergistically and enhance host plant resistance to insect pests by utilizing both cisgenes and transgenes. 15.5.1

Pyramiding cisgenes and transgenes

Integration of pyramided transgenes and native crop plant genes that confer multiple insect pest resistance is a logic priority for research to reduce (or delay) Bt- resistance in pest populations, thereby preserving and augmenting the efficacy of crop-pest-natural enemy tritrophic interactions. Recent reports utilizing RNAi technology in addition to Bt transgenes to control lepidopteran and coleopteran pests (Baum et al. 2007; Gordon and Waterhouse 2007; Mao et al. 2007) also point to exciting research frontiers for managing Bt-toxin resistance in pest populations. Improving host plant resistance by understanding reciprocal molecular interactions between crop plants and their insect pests also has great potential (See Chapter One by Li and Ni in this book for further details). Interest in pyramiding transgenes and integrating plant cisgenes with transgenes is rapidly gaining momentum. All of the significant recent research progress could be summarized in three ways: 1) Synergism among multiple transgenes (Monsanto 2007); 2) Integration and synergism between transgenes and plant cisgenes that confer insect resistance; and 3) Identification of novel insecticidal genes for developing new transgenic crop germplasm (Kaur 2006). Despite a few examples of collaborative research between public and private sectors, profound hurdles still exist between the publically funded research institutions and private companies; most notably, the intellectual property rights (and profits) of the Bt transgenic crops belongs to the private companies. However, the alliance of two private companies, the Monsanto and the Dow AgroSciences, is encouraging (Monsanto 2007). These two companies have established a collaborative research effort to pyramid eight existing transgenes (trade named Smart-Stax) into new corn hybrids to be released in the near future (2010) by both companies (Monsanto 2007). The eight genes will target multiple insect and weed issues in corn production (Monsanto 2007). For aboveground insect control, two genes of YieldGard VTPro from Monsanto and one gene of Herculex 1 from Dow AgroSciences are used. For root-feeding insect control, however, one gene of YieldGard VT Rootworm/Roundup Ready 2 from Monsanto, and two genes of Herculex Rootworm from Dow AgroSciences are utilized. In addition, the two weed control genes are the Roundup Ready 2 Technology from Monsanto, and Liberty Link Herbicide Resistance from Dow

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AgroSciences. The target pests include four insects identified as primary pests, and four insects that are secondary pests (Monsanto 2007). The four primary pests to be controlled by this technology are the European corn borer, Ostrinia nubilalis (Hübner), southwestern corn borer, Diatraea grandiosella Dyar, northern corn rootworm, Diabrotica barberi Smith and Lawrence, and western corn rootworm, and D. virgifera virgifera LeConte. The four secondary pests to be controlled are black cutworm, Agrotis ipsilon (Hufnagel), corn earworm, Helicoverpa zea (Boddie), fall armyworm, Spodoptera frugiperda (J.E. Smith), and western bean cutworm, Richia albicosta (Smith). Such collaborative research efforts using multiple transgene-encoded traits are encouraging for mitigating Bttoxin resistance in insect pest population for long-term and sustainable transgenic crop production worldwide. However, the pyramiding and/or integration of cisgenes and transgenes, and identification of novel insecticidal transgenes are still lagging compared with the pyramiding of transgenes. 15.5.2

Understanding susceptibility to improve resistance

In the history of research on host plant resistance to insect pests, the main goal has been to select crop plants that are either not preferred (antixenotic), antibiotic, or tolerant to insect feeding. These three types of insect-crop plant interactions are the classic mechanisms of host plant resistance (Painter 1951; Smith 1989; Smith et al. 1994; Panda and Khush 1995). However, both susceptible plants and their underlying mechanisms have both been long overlooked, and not been further examined in the continuous screening processes. In contrary, in the history of the development of Bt transgenic crops, a critical aspect is the disentanglement of the etiology of the crown gall disease in crop plants. In particular, identification of factors relating to crown gall formation is one of the three critical aspects of transgenic Bt-crop development. The T-DNA from Ti-plasmid of A. tumefaciens causes host plants to produce hormones abnormally, initiates abnormal prolific growth, and forms tumors (Table 15.3). This critical basic research introduced a new dimension in agricultural research that was previously unknown at that time. It is possible that insect and/or arthropod herbivory might elicit similar or even more interesting responses in susceptible host plants than that of phytopathogens. Information garnered from a better understanding of programmed prolific cell growth and programmed cell death in crop plants might be useful for understanding prolific cell growth associated with some cancers and programmed cell death or apoptosis. Quisenberry and Ni (2007) made similar speculation in their discussion regarding aphid-plant interactions. Likewise, new discoveries in the medical field related to the molecular interactions between pathogen and human host might benefit crop protection research. Continued research of host plant resistance/susceptibility to insects and insecthost plant interactions will lead to novel pest management tactics without any doubt. In particular, in parallel with the three classic mechanisms for host plant resistance to insects, antixenosis, antibiosis, and tolerance, there are three corresponding mechanisms of susceptibility, that is, xenosis, probiosis, and


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intolerance. Deciphering of xenosis, probiosis, and intolerance of host plants to insect herbivory will provide new insights that have great potential to develop a range of novel pest management strategies. For example, identification of oviposition cues of females from the susceptible hosts could lead to the development of kairomone-based traps to attract and kill the female adults to effectively suppress a pest population. Similarly, deciphering of sap-feeding insect-elicited intolerance (e.g., necrosis, chlorosis, and death) of host plants could possibly lead to the development of bio-herbicides. At the same time, we could reduce susceptibility (e.g., probiosis) and improve resistance by inactivating (or silencing) susceptible genes using molecular techniques. The fast growing basic research literature, ranging from nutrient regulation of herbivores (Behmer 2009) to plant immunity to insect herbivores (Howe and Jander 2008), also has great potential to synergistically improve the efficacy of host plant resistance to insect pests research. The outlook for development of new crop cultivars with diverse cisgene- and transgene-encoded traits that confer insect resistance is promising. 15.5.3

Unraveling insect-plant-microbe interactions to reduce mycotoxins

The three-way (multitrophic) interactions among host plants, herbivorous insects, and microbes are facultative components of the ecosystems in general. The roles of microbes range from being beneficial (e.g., nitrogen fixation, and alkaloid production) to being detrimental to plants (e.g., plant pathogens, and mycotoxin contaminations). Although microbial mediation in insect-plant interactions has been examined extensively before the commercial introduction of Bt transgenic crops (Quisenberry and Joost 1990; Barbosa et al. 1991), the introduction of Bt crop in 1996 and increased ethanol production caused by rising energy demand have rejuvenated interest in enhancement of beneficial microbes for crop plants, and mitigation and/or elimination of parasitic and toxigenic microbes in crop production. A gamut of beneficial microbe-plant associations, ranging from bipartite to tetrapartite associations for host plants to cope with nutrient deficient environments, have been described (Prescott et al. 1993). The term bipartite describes the host and single microbe association, such as between a nitrogenfixing Rhizobiae and its legume host, while the term tripartite association was among Rhizobiae and Mycorrhizae with a host plant. The tetrapartite association describes three microbial groups (e.g., endomycorrhizae, ectomycorrhizae, and Frankia) with a host plant (Prescott et al. 1993). The influence of beneficial microbes on either root- or leaf-feeding insect pests is worth further detailed examination. In particular, the possibility of using beneficial microorganisms as vectors for developing new transgenic crops is intriguing. In a similar manner, symbiotic and/or parasitic microbes of insect pests, and their ramifications in insect transgenesis would be another emerging and exciting research area for multi-disciplinary researchers to explore in the coming decades.

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Reduction and elimination of mycotoxin contaminations in grain crops has long been examined extensively in both tropical and warm temperate regions worldwide. It is well documented that, in comparison with the raw corn, ethanol production process caused up to a 3-fold increase in aflatoxin concentration in dried distillers grains (DDGs) (Wu and Munkvold 2008). Such an increase in aflatoxin concentration imposes a high risk when DDGs are used as animal feed. While detoxification and other biochemical treatments have been examined, the antagonistic microbial interactions should be explored as a more efficient approach to reduce mycotoxin production. Plant allelochemical-producing and/ or-transforming microbes for insect pest control in combination with transgenic technology has great potential in designing next generation of transgenic crops. Such synergistic combination is likely to impose less negative ecological consequences than the transgenes alone. Bacon et al. (2007) presented a valuable study documenting control of mycotoxin-producing fungus Fusariumverticillioides using Bacillus mojavensis via host plants. Fusariumverticillioides is known to produce fumonisins, a group of mycotoxins, which are important food and feed safety problems. The fungus detoxifies hydroxamic acids, such as, benzoxazolin-2(3H)-one (BOA) in maize and wheat plants, and produces 2-aminophenol (AP), which is then converted to the less toxic N-(2-hydroxyphenyl) malonamic acid (HPMA) and 2acetamidophenol (HPAA) (Bacon et al. 2007). Bacon et al. (2007) also reported that, in vitro, B.mojavensis could transform hydroxamic acids (also known as BOA) and produce a red pigment accumulation only on BOA-amended media when it is being cultured with wild type and the progeny of genetic crosses of F. verticillioides. The pigment has been identified as 2-amino-3H-phenoxazin-3one (APO), which is a stable product. The results suggest that B. mojavensis disrupts the normal transformation of AP to the nontoxic HPMA by the fungus, resulting in the accumulation of higher amount of APO than when the fungus is cultured alone. The APO is also highly toxic to F. verticillioides (Bacon et al. 2007). Such interactions could be utilized as a biological control agent for reduction and/or elimination of mycotoxin production in corn and other cereal crops that are known to produce hydroxamic acids. At the same time, utilization of the plant allelochemicals for the biosynthesis of anti-microbial compounds is intriguing, which might be used as one of the effective tactics to design new transgenic crops and eliminate or reduce mycotoxin contaminations. Similarly, the effect of the antimicrobial compounds, e.g., APO, on various key insect pests of crop plants could be further examined. Unraveling of the details in antagonistic interactions between a plant allelochemical-transforming bacterium and its capacity to eliminate mycotoxin-producing fungus is intriguing, because the transgenes from plant allelochemical-transforming microbes are likely to be benign to the nontarget organisms in agricultural and natural ecosystems compared to the insecticidal Bt-endotoxin-producing transgenes.


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Conclusion

Scientists repeatedly achieve breakthrough discoveries by seeking the mechanistic truth underlying root causes of natural phenomena and/or tragedies. Ironically, many times these breakthroughs require disproof of the current dogma ranging from “simultaneous generation of maggots in meat” to “Bacillus is the etiological agent of influenza.” These stimulating historical facts and lessons in this review have demonstrated the dynamic nature of the never-ending quest to uncover the truth in scientific research, irrespective of its utilization of either basic or applied research approach. Gupt (1990) used an elegant quotation to summarize the dynamic and successive nature of scientific research: “yesterday’s dogma is to-day’s anathema is to-morrow’s hypothesis revised.” While most of the historical achievements in host plant resistance to insects have demonstrated the critical role of applied research to solve practical research problems, the development of Bt-transgenic crops has shown the enormous long-term impact of basic research in multiple disciplines. The research on pyramiding (or integrating) cisgenic and transgenic traits in crop plants (e.g., insect resistance, and quality improvement) has the potential of leading us to a new generation of transgenic crops with improved yield and quality and no (or minimal) negative impact on agricultural and natural ecosystems. The wide deployment and success of Bt and other transgenic crops has also splendidly demonstrated that basic research can lead to revolutionary changes on multiple disciplines far beyond the imagination of an investigator conducting the research. It is imperative for us to cautiously embrace the transgenic technology, and seize the historical opportunities to transform crop protection in this new agricultural era. The cautious actions should be taken to minimize possible risks by performing long-term monitoring of ecological impact of transgenic crops on a large scale from both agricultural and natural ecosystems. At the same time, we must actively and synergistically utilize cisgenes and transgenes to improve yield by reducing insect and disease damage, and enhancing quality with fortified nutrients and reduced mycotoxin contaminations. Acknowledgments The authors would like to express their gratitude to G. D. Buntin (Department of Entomology, University of Georgia-Griffin), J. R. Ruberson (Department of Entomology, University of Georgia-Tifton), A. M. Simmons (USDA-ARS, Vegetable Research Laboratory, Charleston, SC), M. D. Toews (Department of Entomology, University of Georgia-Tifton) and Y. Chu (Department of Horticulture/ NESPAL, University of Georgia, Tifton, GA) for their thorough and critical review of the earlier versions of this chapter. This article reports the results of research only. Mention of trade names or commercial products in this article is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U. S. Department of Agriculture.

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CHAPTER 16

Time for a New Look at the Relationship Between Bt Plants and Insect Natural Enemies Mao Chen and Anthony M. Shelton

Abstract Insect-resistant Genetically Modified (IRGM) plants have become an important component of integrated pest management (IPM) programs worldwide. The currently available IRGM plants express insecticidal proteins from Bt (Bacillus thuringiensis), however additional insecticidal molecules are being investigated. In 2008, Bt crops were grown on 46 million ha, up from 42.1 million ha in 2007. However, the ecological safety of Bt crops continues to be debated. Much of the debate has focused on non-target organisms, especially predators and parasitoids that help control populations of pest insects in many crops. Thirteen years of commercial use of Bt plants in different parts of the world has documented their effectiveness in controlling target insects and delaying resistance development. The existing "high dose/refuge" insecticide resistance management (IRM) practices certainly contributed to delaying resistance development. However, natural enemies may have played an important role in diminishing the likelihood of resistance development, but this is an area that has not been explored to a great degree. Numerous individual case studies and recent meta-analysis studies of Bt plants and natural enemies indicate that the existing Bt plants have little or no adverse effects on predators, especially compared with conventional insecticides. However, negative effects were found on parasitoids in a few studies but such effects were most likely caused by poor prey/host quality. Using a unique system consisting of a strain of the insect pest, Plutella xylostella (herbivore), resistant to Cry1C that was allowed to feed on Cry1C broccoli plants and then become parasitized by Diadegma insulare, an important endoparasitoid of P. xylostella, our results indicated the parasitoid was exposed to toxic insecticidal protein residues while in the host but was not harmed by such exposure. Parallel studies conducted with several commonly used insecticides indicated they significantly reduced parasitism rates on strains of P. xylostella resistant to these insecticides. These results provide the first clear evidence of the Mao Chen, Anthony M. Shelton Department of Entomology, Cornell University, NYSAES, Geneva, New York, 14456, USA E-mail: [email protected]


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lack of hazard to a parasitoid by Bt plants, compared to traditional insecticides, and help provide new insight on how to rigorously evaluate the ecological risks of Bt plants. Furthermore, these results also suggest that it is time to take a new look at the relationship between Bt plants and insect natural enemies. Bt plants can be a friend to natural enemies, rather than a foe. The potential role of natural enemies in regulating insect resistance development to Bt plants is discussed. Keywords Bt plants, Plant-insect interactions, integrated pest management, insecticide resistance management

16.1

Introduction

The most recent survey on growing genetically modified (GM) crops indicates an accumulated benefit of $ 44 billion for the period of 1996 to 2007, including a savings of 359, 000 tons of active ingredient of pesticides (James 2008; Brookes and Barfoot 2009). GM crops have not only improved the income and quality of life of farmers, but provided tremendous benefits to the environment and agroecosystem. Insect-resistant genetically modified (IRGM) plants expressing insecticidal proteins (Cry toxins) from Bacillus thuringiensis (referred to as Bt plants hereafter) were first commercialized in 1996 in Australia, Mexico, and the US. The adoption and use of Bt plants has offered an alternative to traditional synthetic insecticides for control of important agricultural pests (Shelton et al. 2002). For instance, as of 2008 Bt corn and Bt cotton have been adopted by farmers in 22 countries to control lepidopteran pests such as corn borers (mainly Ostrinia nubilalis, Lepidoptera: Crambidae) in maize and the budworm/ bollworm complex (Heliothis virescens, Lepidoptera: Noctuidae and Helicoverpa zea and armigera, Lepidoptera: Noctuidae and Pectinophora gossypiella Lepidoptera: Gelechiidae) in cotton (Shelton et al. 2002; James 2008). The use of Bt plants has resulted in economic benefits to growers and reduced the use of conventional insecticides (Brookes and Barfoot 2008). However, the continual and season-long expression of Cry toxins in Bt plants has raised concerns about the evolution of resistance in target insects (Bates et al. 2005; Ferré et al. 2008), potential impact on natural enemies (O’Callaghan et al. 2005; Romies et al. 2006; Wolfenbarger et al. 2008; Naranjo et al. 2009), and other ecological risks (Shelton et al. 2002; O’Callaghan et al. 2005). From an IPM perspective, one of the greatest challenges for Bt plants is to delay resistance development in target insects and protect insect natural enemies as biological control agents. For IRM, the US and several other countries require the use of the high dose/refuge strategy (Ferré et al. 2008). This strategy requires the expression of the toxin by the plant to be at a sufficient level to control the heterozygotes within a population (prior to resistance occurring, the resistance allele will be most dominant in that genotype) and a refuge of non-Bt plants to serve as a source for susceptible alleles in the population. This strategy has been considered very successful. In comparison with synthetic insecticides, to date,

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there is only one verified case of an insect that has developed resistance to a Bt plant in the field (Spodoptera frugiperda, Lepidoptera: Noctuidae, on Bt maize in Puerto Rico) since the introduction of Bt plants in 1996 (Ferré et al. 2008). The “high dose/refuge” strategy is one of the major contributors to such success, however the role of insect natural enemies (predators and parasitoids) in regulating target insect resistance development to Bt plants has not garnered much attention. Potential impact of Bt plants on predators and parasitoids has been extensively evaluated based on individual studies (O’Callaghan et al. 2005; Chen et al. 2006; Romeis et al. 2006) and meta-analyses (Marvier et al. 2007; Wolfenbarger et al. 2008; Naranjo 2009). For this chapter, we summarized the impact of Bt plants on insect natural enemies from individual and meta-analysis studies in an effort to take a new look at the relationship between Bt plants and natural enemies.

16.2

Bt plants R&D

The insecticidal activity of Bt has been known for more than a century and commercial Bt products with effective insecticidal activity on targeted pests have been available for 70 years. Bt products account for>90% of the bio-pesticide market (Glare and O’Callaghan 2000; Naranjo 2009), although the bio-pesticide market is a small fraction of the global pesticide market. As a result of rapid development of modern DNA recombination technology, genes that encode for the insecticidal crystal proteins (Cry toxins) produced by Bt have been genetically engineered into several crop species, such as cotton and maize, to generate genetically modified (GM) crops. Because of the consistent economic, environmental and welfare benefits of growing GM crops, in 2008 the number of countries planting GM crops reached 25 and the total global area of GM crops reached 125 million ha. This is a remarkable growth since only 6 countries planted GM crops in 1996, their first year of commercial use. Within this total area of GM crops, Bt plants (including an insect resistant trait only and stacked traits for herbicide tolerance and insect resistance) were grown on 46.0 million ha (or 37% of total GM crop area) in 22 countries (James 2008). The majority of this total was Bt maize expressing the proteins Cry1Ab, Cry1Ac or Cry9C to protect it against O. nubilalis and Sesamia nonagriodes (Lepidoptera: Noctuidae, Mediterranean corn borer), Cry1F to protect it against S. frugiperda, and Cry3Bb, and the binary toxin Cry34Ab and Cry35Ab to protect it against rootworms in the genus Diabrotica (Coleoptera: Chrysomelidae) (Hellmich et al. 2008). Bt maize was grown on 31.6 million ha in 17 countries in 2008. Bt cotton with the genes cry1Ac, cry2Ab2 and cry1F (in China, the Cowpea trypsin inhibitor (CpTI) is also utilized) was grown on 14.5 million ha in 10 countries in 2008 (James 2008). Bt cotton is highly active against the lepidopterans that feed on cotton bolls: H. virescens, P. gossypiella, and reasonably effective against H. armigera and H. zea (Naranjo et al. 2008).


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Bt potatoes were first commercially produced in the US in 1995; however issues with cost, the narrow spectrum of activity, and the arrival of the broader spectrum neonicitinoid insecticides and consumer acceptance led to their retraction from the market after 5 years. However, Bt potatoes are currently under consideration for introduction to the markets in Asia, Africa and Eastern Europe (Grafus and Douches 2008; Naranjo 2009). Although Bt cotton and Bt maize are the only Bt crops currently available for commercial production in many countries, other potential Bt crops with high market values are being actively investigated. For instance, Bt rice lines with high levels of resistance against the rice striped stem borer, Chilo suppressalis (Walker) (Lepidoptera: Crambidae), yellow stem borer, Scirpophaga incertulas (Walker) (Lepidoptera: Pyralidae), and pink stem borer, Sesamia inferens (Walker) (Lepidoptera: Noctuidae) and the leaffolder, Cnaphalocrocis medinalis (Gueneé) (Lepidoptera: Pyralidae) have been field tested since 1998 and, consequently, there are several elite events that have entered the final stage before commercialization in China (Chen et al. 2006; Cohen et al. 2008). Under the aegis of the Collaboration on Insect Management for Brassicas in Asia and Africa (CIMBAA 2008), development of dual-gene Bt transgenic cauliflower and cabbage was undertaken as a sustainable contribution for the management of Plutella xylostella (Lepidoptera: Plutellidae) and other key lepidopterans with the aim of it replacing the use of broad spectrum, traditional insecticides in India and Africa (Shelton et al. 2008). In addition, Bt eggplant expressing cry1Ac with resistance to Leucinodes orbonalis (Lepidoptera: Pyralidae) was produced in 2000 by the Maharashtra Hybrid Seeds Company Limited (Mahyco) under a collaborative agreement with Monsanto. The elite event was first tested in a contained trial in 2002 in India and is on the verge of commercialization (Shelton et al. 2008).

16.3 New opportunities and challenges to managing insect pest complexes with Bt plants Thirteen consecutive years of experience with commercial production of Bt cotton and Bt maize has proven that the Bt crops can provide high levels of protection against some of the target pest species. Before the introduction of Bt plants, the control of corn insect pests, such as European corn borer and corn rootworms, and cotton insect pests, such as cotton bollworm/budworm complexes, primarily relied on synthetic insecticides, which in turn led to the rapid resistance development in pests and negative impacts on insect natural enemies (Wu and Guo 2005; Hellmich et al. 2008; Naranjo et al. 2008). After the introduction of Bt cotton and Bt maize in 1996, Brookes and Barfoot (2008) estimated that between 1996 and 2006 the deployment of Bt cotton world-widely led to a 22.9% reduction of insecticides (i.e. reduced the total volume of insecticide active ingredient by 128.4 million kg), provided economic benefit of $9.6 billion, and reduced the environmental impact quotient (EIQ, a measure of a

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pesticide’s harm to the environment, Kovach et al. 2002) by 24.6%. During this same period, use of Bt maize led to a 5.0% reduction of insecticides (i.e. reduced the total volume of insecticide active ingredient by 8.2 million kg), provided economic benefit of $3.6 billion and reduced the EIQ by 5.3%. Although Bt rice has not been commercially released, a field study conducted in China indicated that Bt rice reduced insecticide use by 80%, in comparison with conventional varieties (Huang et al. 2005). Furthermore, numerous research and review papers have indicated that Bt plants are safer to natural enemies in comparison with synthetic insecticides because of the lack of direct toxicity of Bt toxins and widescale reductions in conventional synthetic insecticide use in Bt fields, which brings new opportunities to enhance biological control practices. Similar to conventionally bred plants that are resistant to an insect or pathogen, Bt plants can provide a foundation for suppressing targeted pests with little additional management input. However, likewise to conventional bred host plant resistance (HPR), the continual expression of Bt toxins exerts a high selection pressure on targeted pests which, along with the wide adoption of Bt plants, raises concern of resistance development in targeted pests. Furthermore, both conventional and genetic engineered HPR may have similar potential side effects on non-target organisms (such as non-target herbivorous pests and insect natural enemies) and these issues warrant investigations within a proper risk assessment framework. In maize and cotton production systems, a wide range of additional pests, such as aphids, mites, and mirids are not targeted by Bt toxins and can be yield limiting if left uncontrolled. Thus, while Bt crops represent an important tactic for managing key target pests, they must be integrated into a more comprehensive IPM program in order to attain successful management of the entire pest complex (Naranjo 2009). This presents challenges for the sustainable use of Bt crops, especially in developing countries where regulations on IRM strategies may not be followed and where setting aside land for a refuge is more difficult. For example, in 2008, 90% of GM crops growers were small, resource-poor farmers in developing countries (James 2008), who were in great need of education on agricultural biotechnology and reasonable use of insecticides for control of non-target secondary pests in Bt plants.

16.4 From case study to meta-analysis: evaluate the impact of Bt plants on insect natural enemies 16.4.1

Lessons from synthetic insecticides

In the late 1940’s and 50’s, synthetic insecticides were first introduced to agricultural production. With the introduction of synthetic insecticides, it became possible to achieve high levels of insect control easily and inexpensively. Because of the power and promise of insecticides, agricultural entomologists focused heavily on the development and use of chemical controls (Kennedy 2008). Approximately 5 million tons of pesticides are applied annually in the world, of


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which about 70% is used for agriculture (Tardiff 1992). Despite early concerns about the risks associated with near-exclusive reliance on insecticides for pest control, the prophylactic use of insecticides grew until an array of serious problems became apparent. For instance, an organochlorine pesticide DDT (dichlor-diphenyl-trichlorethylene), caused eggshell thinning in several high trophic level avian species and sufficiently impacted reproduction to result in population declines (Risebrough 1986). Hamburg and Guest (1997) reported that the number of cotton bollworm predators found in a plot without synthetic insecticide spray could reach ca. 110/16 plants, while < 5 predators/16 plants were found in insecticide sprayed plots. In the absence of insecticides, an average daily predation rate of 67% of cotton bollworms (eggs and larvae) was found. In addition, as a secondary effect of chemical sprays,>200 red spider mites/leaf were found in the sprayed treatment, while < 10 red spider mites were found in the control plots (Hamburg and Guest 1997). In addition, exclusive use of synthetic insecticides has also caused outbreaks of secondary pests and resurgence of target pest populations following destruction of beneficial arthropods; dramatic control failures following the development of insecticide resistance; hazards to pesticide applicators, consumers, and wildlife; and a general simplification of the biotic component of the agroecosystem (Smith 1970; Kennedy 2008). Lessons from synthetic insecticides remind us of the need for a wise and sustainable use of any new strategy (including Bt plants) for pest control within an overall IPM program. 16.4.2

Impact of Bt plants on insect natural enemies

Since the first introduction of Bt plants in 1996, controversy about the benefits and ecological risks of transgenic crops has been long debated (Shelton and Sears 2001). Potential impact, especially on non-target organisms (predators and parasitoids) and compatibility with biological control, was a major concern with the deployment of transgenic crops (Romeis et al. 2006). During the past 13 years of commercialization of transgenic crops, dozens of research articles and news relating to biosafety of transgenic crops were published monthly, especially after the “Pusztai case” and monarch butterfly reports which purported to indicate unintended hazards of GM crops (Dorcey and Serrano 2002). Two major groups of insect natural enemies are predators and parasitoids. Predators usually feed on several different species of insects during their lifetime, thus exposing themselves to several sources of the toxin. The situation with parasitoids is fundamentally different since they generally complete their development within a single host. Thus, if their host feeds on Bt plants it is likely that the parasitoid will become exposed to the Cry toxin. Negative impacts of Bt toxins on non-target parasitoids have been reported in some plantherbivore-parasitoid (tritrophic) studies with Bt plants and susceptible herbivores that were fed Bt plant tissue (Bernal et al. 2002; Baur and Boethel 2003; Meissle et al. 2004; Prütz and Dettner 2004; Vojtech et al. 2005), although the negative

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impacts on parasitoids most likely were mediated by poor host-quality (Romeis et al. 2006; Worfenbarger et al. 2008; Naranjo 2009). A large number of case studies on the impact of Bt plants on non-target organisms (including predators and parasitoids) have been published. A series of individual field studies conducted in the US and Australia that focused on the longer-term assessment of potential non-target effects of Bt cotton and Bt maize were published in Environmental Entomology (Naranjo et al. 2005). Bt plants producing one or more of a possible five insecticidal proteins were evaluated over multiple years. All 13 individual studies covered 772 non-target taxon-times in total (spiders, insect predators, parasitoids, herbivores, detritivores, and Collembola). The results documented that no negative impacts were found on the wide taxonomic breadth of non-target arthropods. In contrast, adverse impacts were found where conventional synthetic insecticides were applied. Romeis et al. (2006) compiled data from more than 100 peer-reviewed studies worldwide on biosafety of transgenic insecticidal crops and found no indication of direct toxic effects of Bt plants on natural enemies under laboratory and greenhouse conditions, unless Bt-susceptible, sublethally damaged herbivores were used as prey or host. Simultaneously, field studies confirmed that the abundance and activity of biological control agents were similar in Bt and non-Bt crops. In contrast, conventional chemical insecticide applications usually reduced biological control. Chen et al. (2006) analyzed a large number of individual studies on the impact of Bt rice on non-target organisms and did not find any direct toxic effects of Bt toxins on predators and parasitoids. However, in a very controversial paper, Hilbeck et al. (1998a) reported that Chrysoperla carnea Stephens (Neuroptera:Chrysopidae) larvae suffered a fitness cost when fed on Bt maize-reared-lepidopteran larvae and claimed it was associated with the Cry1Ab protein and that Cry1Ab is toxic to C. carnae. However, subsequent studies clearly indicated that Cry1A proteins are not toxic to C. carnae larvae (Romeis et al. 2004; Rodrigo-Simón et al. 2006; Lawo and Romeis 2008) and that these proteins do not bind to the midgut of C. carnea, a prerequisite for toxicity (Rodrigo-Simón et al. 2006). These results strongly indicate that the effects observed by Hilbeck et al. (1998a) were due to C. carnea feeding on poor quality (sick or dying) lepidopteran prey. The C. carnae story also applies to other individual reports on the claimed negative impact of Bt plants on parasitoids, i.e. negative effects are host quality driven but not due to direct toxicity of Bt toxins (Romeis et al. 2006; Chen et al. 2008; Wolfenbarger et al. 2008; Naranjo 2009). Meta-analysis is a statistical approach that combines the results of several studies that address a set of related research hypotheses through identification of a common measure of effect size using meta-regression, which can take into account the variability, sample size and the magnitude of differences in comparative studies (Egger et al. 1997). Different from the quasi-quantitative (vote-counting) approach, a meta-analysis can be used to overcome the weakness of vote-counting syntheses by quantitatively combining the results of multiple studies. In early 2006, Marvier and colleagues collated the extant literature on the


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non-target effects of Bt crops on invertebrates (http://delphi.nceas.ucsb.edu/ btcrops) and published the first general meta-analyses of 42 field-based non-target studies focused on Bt maize and cotton (Marvier et al. 2007). Marvier et al. (2007) found that non-target invertebrates (including predators and parasitoids) are generally more abundant in Bt cotton and Bt maize fields than in non-Bt fields managed with insecticides. Subsequently, Wolfenbarger et al (2008) conducted a meta-analysis of 45 field studies dividing the taxa into ecological function guilds and found that predators were slightly less abundant in Bt cotton compared with non-Bt cotton when both did not receive insecticide treatment; parasitoids in Bt maize were much less abundant in Bt maize compared with unsprayed non-Bt maize, which were entirely caused by the reduced abundance of Macrocentris grandii, a specialist exotic parasitoid of the primary Bt maize target, the European corn borer (Wolfenbarger et al. 2008; Naranjo 2009). Cloutier et al (2008) found that predator populations were slighter higher in Bt potatoes compared with untreated non-Bt potatoes using meta-analyses. Recently, Naranjo (2009) conducted extensive meta-analyses of 135 laboratory studies on 9 Bt crops and 22 different Bt Cry protein or protein combinations from 17 countries. He also used meta-analysis from 63 field studies on 5 Bt crops and 13 Bt proteins from 13 countries in comparison with the meta-analyses of laboratory studies in order to rigorously evaluate the impact of Bt plants on predators, parasitoids, and other ecological function guilds. Meta-analyses of laboratory studies indicated that development rate of predators was slightly reduced when the predators were exposed to Bt proteins directly (through plant tissues or pure proteins in artificial diets) compared with non-Bt controls. However, Bt proteins had no effects on survival or reproduction of either predators or parasitoids. At tritrophic levels, analyses revealed a clear and significant impact of host quality on the performance of parasitoids, i.e. developmental rates, reproduction and survivor of parasitoids as a group were reduced when they were provided with hosts that had been compromised by exposure to Bt proteins. However parasitoid development and survival were equivalent or even better on good quality hosts exposed to Bt proteins compared with the hosts non-exposed to Bt proteins (Naranjo 2009). As regarding predators, life history parameters were mostly unaffected by Bt proteins regardless of prey quality (Naranjo 2009). Metaanalyses of field studies indicated that predator abundance was slightly lower in Bt cotton but not in Bt maize, Bt rice or Bt eggplant and parasitoid abundance was much lower in Bt maize but not in Bt cotton, Bt rice or Bt eggplant. The lack of consistent effect of Bt plants on predators and parasitoids in laboratory and field studies indicates that other variables are involved in these interactions, most notably density and quality reductions in target prey or hosts (Naranjo 2009). In April 2009, Lovei et al (2009) published a paper in Environmental Entomology to analyze the impact of Bt proteins and other insecticidal proteins (proteinase inhibitors and lectins) on a total of 32 species of natural enemies as evaluated in 82 laboratory studies (55 on Bt proteins and 27 on others). In their paper, the authors summarized multiple life history parameters and used a

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quasi-quantitative (vote-counting) approach to categorize statistical significance based on author-reported P-values. They concluded that 30% of studies for predators and nearly 40% of studies for parasitoids reported significant negative effects on multiple life history parameters. However, their report has been strongly criticized in a 15-authored paper based on a series of mistaken assumptions, improper analytical methods and lack of ecological context (Shelton et al. 2009). For example, Lovei et al. ignored the the non-independence of multiple traits measured on given species in the same study and lumped together the effects of Bt proteins, proteinase inhibitors and lectins, even though each has a different mode of action and spectrum of activity. Most importantly, they failed to differentiate the direct effect of Bt protein and indirect effect of host quality. In a subsequent paper, these effects were teased out and showed that when there was an effect it was due to host quality (Naranjo 2009). Publication of the Lovei et al paper (2009) has caused deep concern in the scientific community that works on risk assessments of GM plants that their results may be used in regulatory decision-making. Based on all available literature on the impact of Bt plants on natural enemies, it appears that host quality was the real cause of reported negative effects, although direct evidence of such a cause is hard to come by. Several studies used resistant lepidopteran hosts to evaluate the impact of Bt plants on natural enemies in an attempt to avoid host quality issues. While they found no direct toxic effect on natural enemies (Schuler et al. 2001, 2003, 2004), they either did not confirm the exposure of the natural enemies tested to Bt protein or did not include a parallel comparison with synthetic insecticides. In order to fill this knowledge gap, we used a novel paradigm consisting of a strain of the insect pest, Plutella xylostella (herbivore), resistant to Cry1C and allowed it to feed on Bt plants and then become parasitized by Diadegma insulare, an important endoparasitoid of P. xylostella. Our results indicated that the parasitoid was exposed to a biologically active form of the Cy1C protein while in the host but was not harmed by such exposure (Fig. 16.1). Parallel studies conducted with several commonly used insecticides indicated they significantly reduced parasitism rates on strains of P. xylostella resistant to these insecticides (Fig. 16.1, Chen et al. 2008). These results provided the first clear evidence of the lack of hazard to a parasitoid by a Bt plant, compared to traditional insecticides, and described a test to rigorously evaluate the risks Bt plants pose to predators and parasitoids.

16.5 Can natural enemies regulate resistance evolution in target pests to Bt plants? As it has been demonstrated in so many individual and meta-analysis studies, Bt plants are safer to predators and parasitoids in comparison with synthetic insecticides. Numerous studies have also shown the importance of predators and parasitoids in the regulation of pest populations in agroecosystems (Chambers et al. 1983; Croft 1990). Especially early in the deployment of Bt plants, the


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Fig. 16.1 The number of Diadegma insulare emerged from different Plutella xylostella strains treated with formulated insecticide, Bt toxin or left untreated.Means (SE) marked with different capital letters are significantly different based on Fisher’s LSD mean separation test (P < 0.05).

frequency of resistance alleles is usually very low (Ferré et al. 2008). Consequently, there are very low numbers of targeted insect pests that can survive in Bt plant fields (possible RR individuals with < 1% RS that survived high dose expression of Bt toxins). The existence of natural enemies in the field may provide good control of surviving individuals, which in turn may reduce the numbers of resistant alleles (and perhaps their frequency) and eventually delay resistance evolution in targeted pests to Bt plants. Thus, it is appropriate to take a focused approach to evaluate the relationship between Bt plants and natural enemies. Synthetic insecticides have been generally treated as a foe to natural enemies due to the acute toxicity and non-selective killing of non-target organisms by many conventional insecticides. Bt plants may be viewed as a friend to predators and parasitoids because of the lack of direct toxic effects on predators and parasitoids. In a conceptual and mathematical model on tritrophic interactions of a plant, an herbivore, and a natural enemy, Gould et al. (1991) concluded that natural enemies that increase differential fitness between susceptible and resistant phenotypes on host plants will accelerate the evolution of resistance; those that decrease the differential will delay resistance. To evaluate these hypotheses, Gould and his colleagues performed a series of experiments using Bt tobacco and potato. Johnson and Gould (1992) conducted field experiments to examine interactions of the tobacco budworm, H. virescens, its natural enemies, and Bt tobacco plants considered partially resistant to H. virescens. They then calibrated a model to study the influence of natural enemies on the evolution of resistance to transgenic tobacco. Simulation results indicated that biological control could

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accelerate the evolution by pests to Bt plants. Conversely, Johnson et al. (1997a, b) carried out controlled studies of a parasitoid species and a pathogenic fungus that attack H. virescens on tobacco. They concluded that the parasitoid would likely delay the evolution of resistance to transgenic tobacco. Arpaia et al. (1997) investigated predation on the Colorado potato beetle, Leptinotarsa decemlineata, on Bt potato plants in greenhouse and field studies. They included predation rates in a mathematical model to simulate impact of natural enemies on the evolution of resistance by L. decemlineata to Bt potato. Simulations also included refuges of conventional potato plants. Their results showed that predation could decrease the rate of resistance evolution. Similarly, Gassmann et al. (2006) demonstrated the effectiveness of entomopathogenic nematodes for reducing the relative fitness of resistant Pectinophora gossypiella on cotton and concluded that nematodes could delay resistance in P. gossypiella to Bt cotton. Heimpel et al. (2005) extended the abstract work of Gould et al. (1991) and modeled the influence of egg mortality on the high-dose/refuge strategy for IRM. They modeled various levels and forms of pest egg mortality: density independence, positive density dependence, and inverse density dependence. Resistance was modeled as a single locus with a fully recessive allele that confers complete resistance with no fitness cost. Their results indicated that both the magnitude and form of egg mortality can influence the rate of resistance evolution, but the importance of egg mortality depends on other ecological processes in the pest population. Chilcutt and Tabashnik (1999) simulated a model of the interactions of a microbial insecticide containing Bt and a parasitoid in the control of P. xylostella. They also modeled the population genetics of P. xylostella and its evolution of resistance to B. thuringiensis. They concluded the use of parasitoids could slow the evolution of resistance to the Bt by decreasing the number of generations in which insecticide treatment is required, but this would not directly apply to Bt plants with continuous expression. As this summary indicates, the literature contains examples of natural enemies delaying resistance as well as accelerating the evolution of resistance, depending whether there is a differential impact on susceptible or resistant phenotypes. Those natural enemies that increase differential fitness between susceptible and resistant phenotypes on host plants will accelerate the evolution of resistance while those that decrease the differential will delay resistance. This general finding is valuable, however it should be evaluated further by comparing traditional foliar insecticides to Bt plants, and use representatives of two types of natural enemies: parasitoid and predator. Currently, we are evaluating the regulation effect of a parasitoid, Diadegma insulare and a predator Coleomegilla maculata on resistance development in P. xylostella to Bt plants in comparison to sprays of two commonly used conventional insecticides, spinosad and a pyrethroid. In our testing systems, we also take into account the behavioral differences of the predator and parasitoid to different genotypes of prey, plant types (non-Bt, Bt or insecticide treated plants), and refuges with or without insecticide treatment. With extensive tests and comparison with modeling studies,


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we hope to provide experiment-based evidence to reconsider the relationship between Bt plants and natural enemies. The information will have relevance to other cropping systems in which biological control can be an important component. Our ultimate goal is to understand better the tritrophic interactions of Bt plants, target pests and natural enemies and thereby improve IPM.

16.6

Concluding remarks

The global area of Bt plants has increased to 46 million ha in 2008 and Bt plants were grown in 22 countries in 2008 (James 2008). Resistance to Bt plants and the potential impact on non-target organisms are prime concerns for the sustainable use of Bt plants. Various strategies have been developed and implemented to reduce the likelihood of resistance development in targeted pests (Gould 1998; Bates et al. 2005; Ferré et al. 2008). The current high dose/refuge strategy, in conjunction with stacked gene strategies, have proven effective in delaying resistance evolution in targeted pests since there is only one verified case of an insect that has developed resistance to a Bt plant in the field (S. frugiperda on Bt maize in Puerto Rico (Ferré et al. 2008)). Numerous individual case studies and meta-analysis studies on the impact of Bt plants on insect natural enemies have indicated that Bt plants are safer to natural enemies than conventional synthetic insecticides. Natural enemies, as an important ecological service provider in agricultural production systems through regulating pest populations, should be given more attention for their potential effect on regulating resistance development in Bt plant systems. For sustainable use of IRGM crops, it will be necessary to integrate the effect of natural enemies into an overall IRM strategy within the larger picture of IPM. More attention should be given to this valuable research area. Acknowledgements We thank H. L. Collins for helpful comments on an earlier draft of the chapter and Drs. J. Z. Zhao, X. X. Liu and Y. H. Li for comments and suggestions. We gratefully acknowledge the financial support of USAID's Program for Biosafety Systems.

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CHAPTER 17

The Development of Pyrethroid Resistance in the Mosquito Culex quinquefasciatus Qiang Xu and Nannan Liu

Abstract Mosquito-borne diseases are the number-one killers of humans worldwide. A major obstacle in controlling these diseases is that mosquitoes have developed resistance to insecticides, including pyrethroids, which are currently the most widely used insecticides for the indoor control of mosquitoes and are the only chemicals recommended for the treatment of mosquito nets, the main tool for preventing malaria in Africa. A large number of studies have shown that multiple, complex resistance mechanisms or genes are likely to be responsible for insecticide resistance and gene overexpression, amplification, and structural mutations have frequently been linked to insecticide resistance in mosquitoes. Among them, the two major mechanisms involved in the development of insecticide resistance are: (1) increased metabolic detoxification of the insecticides and (2) decreased sensitivity of the target sites to the insecticides (i.e., target site insensitivity). In this chapter, we will summarize our current research and knowledge on the molecular mechanisms governing insecticide resistance development in mosquitoes. Keywords Mosquito, pyrethroids, metabolic detoxification, insecticide resistance, metabolic gene, target gene

17.1

Introduction

The repeated blood feedings of mosquitoes have made them the ideal transmitter of a wide variety of disease agents, including Plasmodium falciparum, the most deadly of the malaria parasites, filarial nematodes, and both the yellow fever and Qiang Xu Department of Biology, Abilene Christian University, Abilene, TX, 79699, USA E-mail: [email protected] Nannan Liu Department of Entomology and Plant Pathology, Auburn University, Auburn, AL, 36849, USA


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Qiang Xu and Nannan Liu

dengue viruses. Mosquito-borne diseases are the number one killers of humans (Marshall, 2000). There are an estimated 300–500 million cases of malaria per year, leading to over 1 million deaths annually (World Health Organization [WHO], 1999). Mosquito control is a very important strategy for the control of mosquito-borne disease (WHO 2000) and insecticide application plays an important role in this effort. However, mosquito-borne diseases are now reemerging as a significant threat to public health worldwide, increasing in incidence in areas where they were previously thought to be under control and expanding into new geographic regions (Gubler 2002). This is largely due to the difficulty of controlling mosquito vectors that have developed resistance to insecticides (Hemingway et al. 2002; Liu et al. 2004a; Enayati and Hemingway 2006).

17.2 Culex quinquefasciatus and pyrethroid resistance in Alabama Culex quinquefasciatus (Say) is an important disease vector throughout the wet tropics. In the southeastern U.S.A., this species is moderately competent as a vector of West Nile virus (WNV) (Sardelis et al. 2001), and is a primary vector of Saint Louis encephalitis virus (SLEV) in many urban settings (Jones et al. 2002). Cx. quinquefasciatus is the predominant mosquito species in urban areas of the southeastern state of Alabama, especially around Mobile and Huntsville. Current approaches to control mosquitoes, both in Alabama and elsewhere in the U.S., rely mainly on insecticides such as pyrethroids, which are currently the most widely used insecticides for the indoor control of mosquitoes and are the only chemicals recommended for the treatment of mosquito nets, the main tool for preventing malaria in Africa (Zaim et al. 2000). However, controlling mosquitoes with pyrethroids (e.g., permethrin) has proven difficult due to the development of insecticide resistance in mosquitoes. In 2002, two mosquito strains of Cx. quinquefasciatus, HAmCqG0 and MAmCqG0, were collected from Huntsville and Mobile, Alabama, respectively and established in our laboratory (Liu et al. 2004a). Bioassay studies indicated that both strains had high levels of resistance to permethrin and resmethrin (to which the mosquito populations had been exposed prior to collection) (Fig. 17.1) when compared with the susceptible SLab strain (Fig. 17.1; Liu et al. 2004a). Although the MAmCqG0 strain had been exposed to resmethrin for more than 15 years, it had only been treated with permethrin for one year before collection. Thus, while the possibility of the development of resistance to permethrin in the MAmCqG0 strain exists, the likelihood of the high level of resistance to permethrin in MAmCqG0 resulted from cross resistance to resmethrin must be considered. Both HAmCqG0 and MAmCqG0 also had elevated levels of crossresistance to deltamethrin even though this insecticide had not been used against the mosquito populations before collection (Fig. 17.1; Liu et al. 2004a). These results suggest that pyrethroid resistance is common in Culex mosquitoes in Alabama.

The Development of Pyrethroid Resistance in the Mosquito Culex quinquefasciatus

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Fig. 17.1 Insecticide resistance ratios (RRs) in HAmCqG0, MAmCqG0, and S-Lab strains of Cx. quinquefasciatus. RRs were calculated by dividing LC50 of resistant strains by LC50 of S-Lab.

17.3 The effect of laboratory permethrin selection on mosquitoes To gain information on the mechanisms of permethrin resistance in mosquitoes and establish a resistant mosquito strain that is more homozygous for resistance genes, HAmCqG0 mosquitoes, which had 100-fold resistance to permethrin, were further selected for 8 generations with gradually increasing concentrations of permethrin, producing the strains denoted HAmCqG1 to HAmCqG8. Fig. 17.2 shows the procedures adopted for permethrin selection, bioassay, and storage for each generation of HAmCq mosquitoes. Eight generations of selection with permethrin increased the level of resistance in HAmCqG8 3100-fold compared with the S-Lab strain (Xu et al. 2005, 2006a). The changes in permethrin resistance between parental (100-fold) and selected mosquitoes (3100-fold) suggest that resistance to permethrin in Cx. quinquefasciatus populations, or at least in these two strains, is developed rapidly. Such an ability of resistance development has also been found in house flies (Liu and Yue 2000). Interestingly, permethrin selection and bioassay studies on another mosquito species, Aedes albopictus, have revealed different results to those for Culex mosquitoes. The Ae. albopictus mosquito strain, HAmAal, collected at the same time and from the same locations in Alabama showed a normal susceptibility to pyrethroids when compared with laboratory susceptible mosquitoes (Liu et al. 2004b). HAmAal was further selected with permethrin for 5 generations in the laboratory and generated the HAmAalG5. No significant difference was detected in the level of susceptibility to permethrin in HAmAalG5 compared with their parental strain, HAmAalG0 (Liu et al. 2006). This result is different from the

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Fig. 17.2 The selection processes used for breeding successive generations of the HAmCq mosquito.

studies on Cx.quinquefasciatus, in which resistance to permethrin was dramatically increased following permethrin selection (Xu et al. 2005; Xu et al. 2006a). Although the reason for this difference in the ability to develop resistance between Ae. albopictus and Cx. quinquefasciatus has not yet been determined, it may result from lacking certain mechanisms that involved in the development of resistance as those in Culex mosquitoes.

17.4

Molecular mechanisms of pyrethroid resistance

The two major mechanisms involved in the development of insecticide resistance are: (1) increased metabolic detoxification of the insecticides and (2) decreased sensitivity of the target sites to the insecticides (i.e., target site insensitivity) (Taylor and Feyereisen 1996; Scott 1999). A target site is the protein on which an insecticide acts. The resistance mechanism of increased sequestration or detoxification contributes to a decrease in the effective dose of insecticides available at the target site, while decreased target site sensitivity contributes to the ineffective binding of a given dose (Scott 1990; Feyereisen 1995; Pasteur and Raymond 1996). The products of three gene families, cytochrome P450 monooxygenases (cytochrome P450s), esterases, and glutathione transferases (GSTs), are primarily implicated in the detoxification of insecticides, whereas three proteins, sodium channels, acetylcholinesterase, and GABA receptors, are known to be heavily involved in decreased target site sensitivity in insect nervous systems (Feyereisen 1995). The mechanisms conferring pyrethroid resistance in


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mosquitoes can thus be conveniently divided into two major groups, namely metabolic detoxification and target site insensitivity of sodium channels. 17.4.1 Role of metabolic detoxification in permethrin resistance of HAmCq mosquitoes

As noted above, the detoxification of insecticides in the mosquito involves 3 major metabolic detoxification gene families: cytochrome P450s (P450s), esterases, and glutathione S-transferases (GSTs) (Feyereisen 2005; Ranson and Hemingway 2005; Oakeshott et al. 2005). Transcriptional up-regulation of P450s results in increased levels of P450 proteins and in increased P450 activities, both of which promote the development of insecticide resistance (Carino et al. 1994; Liu and Scott 1997, 1998; Feyereisen 2005). The process is not straightforward, however; although an increase in the DNA amplification of esterases is often associated with esterase-mediated insecticide resistance (Hemingway and Karunaratne 1998; Small and Hemingway 2000), non-elevated levels of esterases have also been identified in mosquitoes exhibiting insecticide resistance (Whyard et al. 1995) and GST gene up-regulation has been identified in the mosquito Anopheles gambiae, which is resistant to pyrethroids (Ortelli et al. 2003). Synergists have long been exploited as tools to diagnose particular insecticide resistance mechanisms based on their ability to inhibit specific metabolic pathways (Scott 1999; Liu and Scott 1996; Liu and Yue 2000, 2001; Pridgeon et al. 2003). When HAmCq and MAmCq mosquitoes were treated with PBO, an inhibitor of cytochrome P450 monooxygenases, resistance to permethrin in these mosquitoes were dramatically decreased (Xu et al. 2005), suggesting that P450 monooxygenase-mediated detoxification contributes to permethrin resistance in both HAmCq and MAmCq mosquitoes. DEF, an inhibitor of esterases, and DEM, an inhibitor of GST, also can partially decreased permethrin resistance in HAmCq and/or MAmCq mosquitoes (Xu et al. 2005), reveling a minor contribution of esterase- and GST-mediated metabolisms to resistance. However, since neither PBO and DEF, nor DEM completely abolished resistance to permethrin in HAmCq and MAmCq mosquitoes, one or more additional mechanisms other than metabolic detoxification are likely to be involved in overall permethrin resistance. 17.4.2

Role of sodium channel insensitivity in pyrethroid resistance

The target sites in insects involve 3 major target proteins: acetylcholinesterases (AChEs), sodium channels, and GABA receptors (Oppenoorth 1985; Soderlund and Knipple 2003; Buckingham and Sattelle 2005). Target-site insensitivity results from structural modification or mutation (point mutation) of the target proteins acted on by insecticides. Point mutations in the target proteins cause a reduction in the nervous system’s response to insecticides (or a reduction in the binding affinity of the protein to the insecticides) (Narahashi 1988), which in turn results in the development of insecticide resistance. In insects, sodium channels

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are the targets for insecticides such as DDT and pyrethroids. After bonding with the sodium channels, insecticides cause the insect’s nervous system to repetitively discharge and its nerve membranes to depolarize (Narahashi 1988), resulting in the insect’s death. Insects are known to develop resistance to pyrethroids and DDT through structural modifications of their sodium channels that reduce their sensitivity to insecticides. The term “knockdown resistance” (kdr) is used to describe cases of resistance to DDT and pyrethroid insecticides in insects and other arthropods, resulting from the reduced sensitivity of the sodium channels. It has been found that a single mutation of leucine to phenylalanine (L to F), termed kdr mutation, in the sodium-channel gene has been involved in pyrethroid resistance of insects, including mosquitoes (Brengues et al. 2003; Martinez-Torres et al. 1998, 1999; Enayati et al. 2003). The L to F mutation (the kdr mutation) resulting from a single nucleotide polymorphism (adenine to thymine, A to T) in the sodium channel of Culex mosquitoes was found to cause insensitivity of the sodium channel to pyrethroids and, in turn, to lead to pyrethroid resistance in mosquitoes (Martinez-Torres et al. 1999). In addition, a substitution of the same leucine to serine (L1014S) has been found in Anopheles gambiae (Ranson et al. 2000). It has been suggested that the L to S substitution is associated with a high level of resistance to DDT combined with a low level of resistance to pyrethroid, while the L to F substitution is associated with high resistance to both pyrethroid and DDT (Martinez-Torres et al. 1999; Ranson et al. 2000). The L to F substitution was identified in both the HAmCq and MAmCq Culex mosquito strains (Xu et al. 2005). The frequency of alleles (A or T) at the kdr locus at the RNA level in HAmCqG0, and HAmCqG8 mosquitoes was investigated. A strong correlation between the frequency of the kdr allele expression at the RNA level and the level of resistance was identified: 100% of susceptible S-Lab strain expressed the susceptible A allele and 93% of HAmCqG8 expressed the resistance T allele (Fig. 17.3; Xu et al. 2006a). A pattern of allelic variation at the kdr locus in the individuals of HAmCqG0 (the field collected strain) was identified, with frequencies ranging from 0.2 for individuals that expressed only the susceptible allele A, 0.43 for those that expressed both alleles A and T, and 0.37 for those that expressed the kdr allele T (Fig. 17.3). The allelic variation pattern in this field collected HAmCqG0 mosquito population strongly correlated with its intermediate levels of resistance (Liu et al. 2004a) and the proportions of both susceptible and resistant individuals in this field population. Recently, Liu et al. (2006) conducted a similar study on characterization of the correlation of the kdr allelic expression and the levels of the resistance in HAmAalG0 and HAmAalG5 strains of Ae. Albopictus. Every individual in HAmAalG0 and HAmAalG5 mosquitoes expressed a codon of CTA at the L-to-F kdr site encoding Leu, strongly corresponding to their susceptibility to insecticides (Section; Liu et al. 2006). Xu et al. (2006b) also conducted studies of kdr allelic expression within and among resistant and susceptible house fly and German cockroach populations. A strong correlation between kdr allelic


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Fig. 17.3 Allele-specific transcription for two alleles in the kdr locus in mosquitoes Cx. quinquefasciatus. Thirty mosquitoes in each population were used for a total of 30 SNP determinations, with each of the 10 SNP determinations being repeated three times.

expressions and levels of insecticide resistance was found in both insect species (Xu et al. 2006b). These findings (Xu et al. 2006a, b; Liu et al. 2006) highlight the extraordinary ability of insects to adapt to evolutionary selection by regulating a target site with different transcripts and document the crucial role that the L-to-F kdr mutation plays in pyrethroid resistance in insects. However, although we know that the L-to-F kdr mutation is very important in the evolution of resistance development, it is unlikely to be the sole mechanism responsible for such a widespread phenomenon as kdr-mediated resistance. In particular, more than 20 sodium channel mutations have already been identified as being involved in reducing channel sensitivity to insecticides or neurotoxins (Soderlund and Knipple 2003). Very recently, several novel mutations have also been identified in the sodium channel gene of the mosquito Cx. quinquefasciatus (Xu et al. unpublished data). Further investigation of the role of these novel mutations will provide a more comprehensive picture of the mechanisms involved in sodium channel mediated pyrethroid resistance in mosquitoes. 17.4.3

Genes involved in mosquito resistance

Results from many previous studies (Liu et al. 2004a; Xu et al. 2005, 2006a, b; Liu et al. 2007; Liu and Scott 1996; Liu and Yue 2000, 2001; Pridgeon et al. 2003) suggest that the interaction of multiple insecticide resistance mechanisms or genes is responsible for insecticide resistance. To identify the genes involved in resistance beside the sodium channels we therefore compared the gene expression profiles of the HAmCqG6 (the last permethrin-selected generation at the time of the study) and S-Lab strains using an SSH/cDNA array technique to characterize the genes differentially expressed in HAmCqG6 and S-Lab by constructing a HAmCqG6-S-Lab subtractive library (Liu et al. 2007). Twenty two unique genes

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were identified that were over-expressed in HAmCqG6, 16 of which were functionally interpretable as being vital for molecular metabolism, signal transduction and regulation, molecular transport, cytoskeletal networks, protein biosynthesis, and ubiquitination, although most had not previously been implicated in insecticide resistance in mosquitoes (Liu et al. 2007). Two cytochrome P450s showed significantly increased expression in the resistant mosquito strain HAmCqG6 (Liu et al. 2007) and P450s are known to be centrally involved in the metabolic detoxification of xenobiotics, including insecticides. This study thus supports the findings of the previous synergistic study (Section 4.1) that P450-mediated detoxification was one of the major resistance mechanisms involved in pyrethroid resistance in Culex mosquitoes (Xu et al. 2005). Characterizing the function of the overexpressed cytochrome P450s in mosquito resistance will shed new light on the mechanisms of resistance in mosquitoes and provides fundamental information that increases our understanding of metabolic-mediated resistance in mosquitoes. Apart from these two P450 genes, most of the up-regulated genes had not previously been reported in the literature on insecticide resistance. For example, two of the overexpressed genes are Rhodopsin and Arrestin genes, both of which play a crucial role in signal transduction systems. Rhodopsin is an archetypal class A G-protein-coupled receptor (GPCR) (Filipek et al. 2003). GPCRs act as transducers for a range of different sensory, chemotactic, hormonal, and neuronal signals, and are involved in many essential physiological functions of organisms. Arrestins are a gene family of regulatory proteins that, along with G-proteincoupled receptor kinases (GRKs) and other co-factors, regulate the signaling and trafficking of G-protein-coupled receptors by virtue of their preferential binding to the phosphorylated active form of the receptor. Up-regulation of these two genes in resistant mosquitoes may indicate that the neuronal signaling is affected in the resistant mosquitoes. The resistance-specific overexpression of arrestins and rhodopsin in resistant mosquitoes was, for the first time, identified in a study (Liu et al. 2007) highlighting the functional importance of the signal transduction system in the regulation of insecticide resistance. A similar study by Vontas et al. (2006) indentified multiple genes that are up-regulated in DDT-resistant Anopheles gambiae mosquitoes. The data reported by Liu et al. (2007) and Vontas et al. (2006) suggest that pyrethroid resistance in mosquitoes involves multiple genes and multiple complex mechanisms.

17.5

Conclusions

Insecticide application is the most important component in the global mosquito vector control effort, and pyrethroids are currently widely used in this capacity. A better understanding of the mechanisms governing resistance development is crucial for developing novel strategies to circumvent and/or delay the development of resistance, control resistant mosquitoes and, ultimately, reduce the prevalence of mosquito-borne diseases. Characterization of the genes and


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mechanisms involved in resistance may lead to new approaches to study insecticide resistance. For example, a better understanding of the regulation mechanisms or genes controlling pyrethroid resistance is likely to open up new aspects of insect resistance research, possibly representing radically new approaches for studying the network structure. These could include research into the specific functions of these genes in resistance, the development of a comprehensive map of the resistant gene interaction network, and an in-depth examination of the biochemical pathways by which such interactions modulate the physiological properties in insecticide resistant systems. Identification of the genes responsible for resistance will also lead to novel strategies for controlling mosquitoes. Once the insecticide resistance genes have been identified, these might offer new targets for insecticides. These newly identified resistance genes are also likely to provide a foundation for studies that seek to pinpoint the genetic markers of insecticide resistance in mosquitoes, as well as for studies that monitor and predict the development of resistance in field mosquito populations. Acknowledgements Authors thank Drs. Lin He, James Nichols, Patricia Hernandez, Tom Lee and Tom Pirtle for their valuable comments on the manuscript. The studies were supported by Auburn University Biogrant 2008-Bio-Liu-03-08 to N.L.; AAES Foundation Grants to N.L.; AAES Hatch/Multistate Grants ALA08-045 to N.L.; and Hatch Project ALA08-029 to N.L.

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CHAPTER 18

Resistance to Transgenic Bacillus thuringiensis Crops in Target Insect Pests: Current Status and Prospect Fangneng Huang

Abstract The ability to transfer foreign genes among unrelated species represents a major technological advance in modern agriculture. Since commercialized in 1996, transgenic Bacillus thuringiensis (Bt) crops (e.g. corn and cotton) have been widely used for managing insect pests. During 2008, a total of >40 million ha of Bt corn and Bt cotton were planted in 22 countries worldwide. Bt crops have been excellent in controlling their target insect pests. Growers of both industrial and developing countries have recognized great economic, ecological, and social benefits by planting Bt crops. Resistance development is considered to be a major concern to the long-term success of Bt crops. Major resistance genes capable of survival on commercial Bt crops have been identified in several target species. However, frequency of major Bt resistance alleles in field insect populations has been estimated to be low in most target insect species. In most cases, resistance to Bt crops is incomplete and controlled by a single recessive or partially recessive gene. Bt resistance is often associated with fitness costs in absence of selection pressures. The most common mechanism of Bt resistance may be due to a reduction in binding of toxins to cadherin receptors in the midgut. Field resistance development has been suspected in four target species. However, up to date, field control failures or reduced control efficacies due to resistance development have been documented in only two isolated cases, the fall armyworm, Spodoptera frugiperda, to Cry1F corn in Puerto Rico in 2006 and the African stem borer, Busseola fusca, to Cry1Ab corn in South Africa in 2007. Mandatory Bt resistance management plans have been successfully implemented in the United States, Canada, and Australia. Because of the substantial benefits achieved from planting Bt crops, it is widely expected that adoption of Bt crops will consistently increase rapidly, especially in the developing countries. Bt resistance management, therefore, will Fangneng Huang Department of Entomology, Louisiana State University Agricultural Center, Baton Rouge, LA, 70803, USA E-mail: [email protected]


Resistance to Transgenic Bacillus thuringiensis Crops in Target Insect Pests

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continue to be a big challenge for the future success of Bt crop technologies. Keywords Transgenic crop, Bt, resistance, target insect pests, Bt corn, Bt cotton

18.1

Introduction

Transgenic crop technology represents a major technological advance in modern agriculture. Transgenic insect-resistant crops expressing Bacillus thuringiensis (Bt) proteins were first commercially planted during 1996 in the United States and four other countries in the world. Since then, Bt crops have been rapidly adopted for insect pest management worldwide (James 2008). The field performance of Bt crop varieties has been outstanding in controlling their target insect pests (Huang et al. 1999a; James 2008). Growers in both industrial and developing nations have recognized substantial economic, environmental, and social benefits by planting Bt crops (James 2008). On the other hand, the rapid adoption of transgenic Bt crops has been considered to be a major threaten to the long-term success of this technology because intensive selection pressure can accelerate resistance development in pest populations (US EPA 2001). For this reason, insecticide resistance management (IRM) for transgenic Bt crops has been one of the most active research areas in entomology during the last decade and a remarkable number of related papers have been published in various journals. This chapter will first present a summary of the current global adoption of Bt crops. Most of the review will focus on the recently published information on field resistance to Bt crops, resistance mechanisms, monitoring, and management. This review will concentrate on only commercialized transgenic Bt crops (e.g. Bt corn and Bt cotton) and their target insect species. Readers who are interested in Bt resistance in general can read several other related journal reviews and book chapters (Whalon and McGaughey 1993, 1998; Tabashnik 1994; Huang et al. 1999a; Ferré and van Rie 2002; Ferré et al. 2008; Gassmann et al. 2009).

18.2

Current global adoption of Bt crops

When transgenic crops were first commercialized in 1996, very limited hectares of Bt crops were planted in five countries including the United States, Canada, China, Australia, and Mexico. By 2008, the number of countries planting Bt crops increased to 22 with a total of >40 million ha (Table 18.1). Corn and cotton have been the predominant commercialized Bt crops. Very small hectares of Bt potato, Bt rice, and Bt poplar have been planted in the United States, Iran, and China, respectively (James 2007, 2008). Among the 40 million ha of Bt crops planted during 2008, 26.4 million ha (65%) were planted with Bt corn in 16 countries and 14.5 million ha (36%) were planted with Bt cotton in 10 countries. Since the commercial use, the United States has been the leading country in planting Bt crops, accounting for >60% of the world’s total accumulated

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Fangneng Huang

hectares. In spite of the rapid increase in planting Bt crops in other countries, the United States still accounted for 55% of the world total Bt crop hectares in 2008. In the United States, during 1995, one year before the officially commercial use of Bt crops, approximately 0.15 million ha of corn were planted with Bt corn. In 1996, 1.6 million ha of Bt corn (5% of corn hectarage) and 0.87 million ha (15% of cotton hectarage) of Bt cotton were planted in the country (Fig. 18.1). By 2008, the United States alone planted >20.1 million ha of Bt corn and 2.4 million ha of Bt cotton, which accounted for approximately 57% and 64% of its total hectarage of corn and cotton, respectively (NASS 2008). The annual increase rate in adoption of Bt crops during the past thirteen years in the United States was 23.5% for Bt corn and 8.8% for Bt cotton. Other countries including India, China, Argentina, Brazil, and South Africa planted >1 million ha of Bt crops during 2008 (Table 18.1).

Fig. 18.1 Percentage and hectares of transgenic Bt corn and Bt cotton planted in the United States from 1995–2008. Percentage of a Bt crop in a year was calculated based on the hectares planted with Bt varieties divided by the total hectares of that crop in the year. Most data were from the annual crop acreage reports by U. S. National Agricultural Statistics Service at http:// usda.mannlib.cornell.edu/usda/current/Acre/Acre-06-30-2008.pdf. [Last accessed 29 April 2009].


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The use of Bt plants for managing corn and cotton pests has increased dramatically in both hectarage planted and number of countries in the developing nations in recent years. For example, India first commercialized Bt cotton with a total of 0.05 million ha in 2002. Its Bt cotton area increased to 3.8 million ha in 2006. India passed China and the United States in 2006 to become the leading country in planting Bt cotton in the world (James 2006). It took only two years for India to double its Bt cotton hectarage of 2006 to 7.6 million ha in 2008. Based on the area planted in 2002, the annual increase rate in planting Bt cotton in India has been 273% during the last 5 years. In addition, Bt corn also has been highly Table 18.1

Global adoption of transgenic Bt corn and Bt cotton in 2008a Corn

Country

United Statesb India

Cotton

Total Bt crop area (million hectares)

Area (million hectares)

% of the total corn hectares

Area (million hectares)

% of the total cotton hectares

20.14

57

2.39

64

22.53

—

—

7.60

82

7.60

—

—

3.80

68

3.80

Argentina

2.30

58

0.16

40

2.65

Brazil

1.30

10

0.25

26

1.55

China

South Africa

1.34

52

0.01

84

1.35

Canada

0.84

53

—

—

0.84

Philippines

b

0.28

10

—

—

0.28

Australia

—

—

0.14

85

0.14

Uruguay

0.11

73

—

—

0.11

Spain

0.08

32

—

—

0.08

—

—

0.07

82

0.07

Colombia

—

—

0.03

100

0.03

Portugal

< 0.01

4

—

—

< 0.01

Czech Republic

< 0.01

3

—

—

< 0.01

Germany

< 0.01

6

—

—

< 0.01

Honduras

< 0.01

2

—

—

< 0.01

Poland

< 0.01

0.4

—

—

< 0.01

Romania

< 0.01

< 0.1

—

—

< 0.01

Slovakia

< 0.01

0.8

—

—

< 0.01

—

—

< 0.01

2

< 0.001

Egypt

< 0.01

0.1

—

—

< 0.001

Total

26.43

—

14.46

—

40.89

Mexico

Burkina Faso

a) Iran planted Bt rice in a total of 6000 hectares during 2004–2005 and data were not available for 2006–2008. A small area of Bt poplar has been planted in China. b) Data for the United States were from NASS (2008) and for Canada were based on the report from Canadian Corn Pest Coalition (2007). All others were from James (2008).

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adopted in Uruguay (77%), Canada (53%), and Spain (32%) and Bt cotton has been widely planted in Australia (85%), Mexico (82%), and Colombia (100%), although the absolute hectarage is not very large in these countries. Several European and Asian countries recently commercialized Bt corn with a limited hectarage (Table 18.1). The potential marketing of Bt crops in the two continents is expected to increase rapidly in the future (James 2008). In addition, transgenic Bt rice was first commercialized in Iran during 2004 with a total of 2000 hectares and about another 4000 hectares were planted during 2005 (James 2006, 2007).

18.3 Bt traits of commonly used transgenic Bt corn and Bt cotton Majority of the first generation of Bt corn (e.g. YieldGrad® corn) and Bt cotton (Bollgrad® cotton) expresses only a single Bt Cry protein and thus usually have a relatively narrow insecticidal spectrum (Table 18.2). For example, the YieldGard® Bt corn was first commercialized in 1996 and has been the most commonly used Bt corn technology worldwide. It expresses only the Cry1Ab protein and is effective against lepidopteran stalk borers, such as the European corn borer, Ostrinia nubilalis and southwestern corn borer, Diatraea grandiosella. In contrast, the YieldGard Plus® corn, released in 2003, contains two Cry genes, Cry1Ab for controlling Lepidoptera and Cry3Bb1 for controlling Coleoptera and thus is effective for managing both stalk borers and rootworms. The most recently released Bt corn, Herculex XTRA by Dow AgroSciences (Indianapolis, Indiana, USA) expresses three Bt proteins including Cry34Ab1, Cry35Ab1 and Cry1F. This Bt corn, in addition to be effective to the primary corn borer and rootworm species, also has significantly insecticidal activities against some secondary corn insect pests such as earworms, armyworms, and cutworms. Similarly, the first commercialized Bt cotton, Bollgard®, expresses only Cry1Ac protein and is effective to the tobacco budworm, Heliothis virescens, pink bollworm, Pectinophora gossypiella, and cotton bollworm, Helicoverpa armigera. The later released Bollgrad II® and WideStrike® cotton both express two Bt Cry proteins. Compared to the single-gene-expressed Bollgard®, Bollard II® is more effective against bollworm, Helicoverpa zea, and also has some insecticidal activities against loopers and armyworms. The WideStrike® also has a broader range of targets compared to Bollgard® cotton (Table 18.2). Transferring two or more different foreign genes into a same plant for different targets or purposes is often called “gene-stacking” (e.g. YieldGard Plus®), while it is termed “genepyramiding” if these foreign genes are for controlling the same target species (e.g. Bollard II®). Recently, a novel gene transfer technology called Vector-stack Transformation (VecTran technology) has been used in developing transgenic crops (Monsanto 2007). This technology allows combining two or more traits using a single DNAinsertion process to produce stacked/ pyramided traits in a plant (e.g. YieldGard


311

Common Bt corn and Bt cotton that have been commercialized since 1996

Table 18.2 Bt corps

a

Year of release

Bt Genes

Major target insect pests

YieldGard corn

1996

Cry1Ab

stalk borers, H. zea

b

Knockout corn

1996

Cry1Ab

stalk borers, H. zea

Bollgard cotton or Ingardc

1996

Cry1Ac

H. virescens, P. gossypiella, H. zea, H armigera

Bt Cotton/CAASd

1996

Cry1Ac

H. armigera, P. gossypiella

Herculex I corn

2001

Cry1F

stalk borers, cutworms, armyworms, H. zea

Bollgard II cotton

2002

Cry1Ac + Cry2Ab2

H. virescens, P. gossypiella, H. Zea, loopers, armyworms

YieldGard Rootworm corn

2003

Cry3Bb1

rootworms

YieldGard Plus corn

2003

Cry1Ab + Cry3Bb1

stalk borers, rootworms

WideStrike cotton

2004

Cry1Ac + Cry1F

H. virescens, P. gossypiella, H. zea, loopers, armyworms

Herculex RW corn

2005

Cry34Ab1 and Cry35Ab1 rootworms

Hexculex XTRA corn

2005

Cry34Ab1 + Cry35Ab1 + Cry1F

stalk borers, rootworms, H. Zea, armyworms, cutworms

Event 1 cottone

2006

Cry1Ac

H. armigera

2006

e

Cry1Ab and Cry1Ac

H. armigera

Agrisure CB corn

n/a

Cry1Ab

stalk borers, H. zea, armyworms

Agrisure RW corn

n/a

Cry3Aa

rootworms

Agrisure CB/RW corn

n/a

Cry1Ab plus Cry3Aa

stalk borers, rootworms, H. zea, armyworms

YieldGard VT Rootworm corn n/a

Cry3Bb1

rootworms

YieldGard VT Triple corn

2007

Cry1Ab + Cry3Bb1

stalk borers, rootworms, H. zea, armyworms, cutworm

VipCotTM cotton

2008

Vip3A + Cry1Ab

H. virescens, P. gossypiella, H. zea, armyworms, loopers

YieldGard VT PRO

2009

Cry1A.105 + Cry2Ab2

stalk borers, armyworms, black cutworm, H. zea

YieldGard VT Triple PRO

2009

Cry1A.105 + Cry2Ab2 + Cry3Bb1

H. virescens, P. gossypiella, H. zea, armyworms, loopers

Genuity SmartStax

2010 (proposed)

Cry1A.105, Cry2Ab2, Cry1F, Cry3Bb1, Cry34Ab1, Cry35Ab1

stalk borers, rootworms, H. zea, armyworms, cutworm

GFM cotton

a) Corn stalk borer species may include Ostrinia nubilalis, Diatraea grandiosella, D. crambidoides, D. saccharalis and S. nonagrioides. Corn rootworm species may include Diabrotica virgifea virifera, D. barberi, and D. virgifera zeae. Armyworm species may include S. frugiperda, S. ornithogalli, and S. exigua. Cutworm species may include Corimelaena pulicaria and Richia albicosta. Looper species may include Pseudoplusia includens and Trichoplusia ni b) non-longer available in the United States c) a marketing name of Bollgrad cotton in Australia d) available in China only e) available in India only

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Fangneng Huang

VT Triple® corn). Transgenic Bt corn developed using VecTran technology (e.g. YieldGard VT pro®) is more consistent in expressing Bt proteins compared to Bt corn products that are produced using traditional gene transfer procedures (Schilling 2007). In addition, pyramiding two or more Bt genes with different insecticidal mechanisms into one plant for controlling a same target species is expected to be able to delay resistance development. Currently, both Bt crops expressing single or multiple Bt genes are planted in the United States and Canada. With an extensive search, I could not find the data on the proportions of each type of the Bt crops being planted, but it is apparent that the hectarage of stacked/pyramided gene varieties has increased significantly in recent years. In Australia, single Bt gene cotton varieties were already phased out and replaced with pyramided-gene varieties (e.g. Bollgard II®) during 2004/2005 growing season (Farrell 2006). However, majority Bt crops planted in other countries express only a single Bt Cry protein (Table 18.2). It is expected that the use of pyramided-gene varieties will be increased dramatically and single-Bt-gene varieties will be completely phased out in the near future (Johnson 2007; Wu 2007).

18.4

Current status of resistance to Bt crops

Up to date, field control failures or reduced control efficacies of Bt crops due to resistance development have been documented in only two target species of Bt corn, the fall armyworm, Spodoptera frugiperda, to Cry1F corn in Puerto Rico in 2006 (Matten et al. 2008) and the African stem borer, Busseola fusca to Cry1Ab corn in South Africa in 2007 (Van Rensburg 2007). Recently, field resistance to Cry1Ac cotton has also been suspected in H. armigera in north China (Xu et al. 2009) and H. zea in the mid-southern region of the United States (Tabashnik et al. 2008). Resistance genes capable of survival on commercial Bt crops have been found in two other target species of Bt cotton, H. virescens and P. gossypiella and one target species of Bt corn, the sugarcane borer, Diatraea saccharalis (Huang et al. 2007a). In addition, a recent study showed that a resistant strain of the western corn rootworm, Diabrotica virgifera virgifera, selected with Bt corn in greenhouse has demonstrated a high survival rate on Cry3Bb1corn plants (Meihls et al. 2008). These findings further demonstrate that effective resistance management is crucial for the long-term success of transgenic Bt crop technologies. 18.4.1

Resistance to Cry1F Bt corn in S. frugiperda in Puerto Rico

Cry1F Bt corn was first commercialized in the Puerto Rico Island for controlling S. frugiperda during 2003. Unexpected field control failures were observed in Puerto Rico during 2006 growing season (Matten et al. 2008). The field control failures of Cry1F corn to S. frugiperda in Puerto Rico are now documented to be due to resistance development (Matten et al. 2008). To our knowledge, this was


313

the first documented report of a field control failure of Bt crops that was caused by resistance development in any where of the world. The corn-insect eco-systems in Puerto Rico might create a very favourable scenario for resistance development. These factors may include a closed island insect population, continuous corn production throughout the year, high insect pest pressure, and drought conditions. However, because baseline susceptibility data are not available before the introduction of Cry1F corn, it is still unclear if the field resistance was caused by selection pressures from planting Bt plants or if these populations were naturally more tolerant to the Bt toxin or both factors. After the field control failures were observed, selling of Cry1F corn seeds was discontinued in Puerto Rico (Matten et al. 2008). 18.4.2

Resistance to Cry1Ab corn in B. fusca in South African

Cry1Ab corn was first introduced for control of a corn stalk boring complex of B. fusca and Chilo partellus in South Africa during 1998/1999 growing season (van Wyk et al. 2008). Since then adoption of Bt corn has increased rapidly. Approximately 50,000 ha of Cry1Ab corn (Event Mon810) were planted in the country during the first season. The Bt corn area increased to 943,000 ha (35% of the total corn hectarage) in 2006 and approached more than 1.3 million ha (52% of the total corn hectarage) in 2008 (Table 18.1, van Rensburg 2007). Cry1Ab corn hybrids were effective against the stem borers in vegetative plant stages during the first planting season in South Africa. However, at the harvest of the crop in 1999, diapause larvae and damaged plants were observed in various Bt corn hybrids (van Resburg 2007). During 2004/2005 growing season, severe damage of B. fusca to vegetative stages of various Bt corn hybrids, which were produced by different seed companies, was observed in several locations. All affected plantings were grown under irrigation and all fields had a history of continuous Bt corn production (van Rensburg 2007). Field and greenhouse tests showed that progeny survival and larval mass gain of borer populations derived from insects collected from Bt corn plants were considerably greater than those collected from non-Bt corn areas (van Rensburg 2007). The results confirmed that the significant damage of Bt corn plants in the field was due to resistance development in B. fusca. Reasons for resistance development in B. fusca so rapidly are still unclear, but there are some similarities between the two documented cases of field-evolved resistance. The Bt plants apparently did not express a high dose of Cry proteins against S. frugiperda and B. fusca. It is well known that S. frugiperda is relatively less susceptible to Cry toxins compared to most other corn lepidopteran pests such as O. nubilalis and D. grandiosella (US EPA 2001). Majority of the currently used Bt corn hybrids do not express a high dose of Bt proteins against these insect pests. Similarly, B. fusca is much less susceptible to Bt corn compared to C. partellus (van Rensburg 2007). The survival of B. fusca on Cry1Ab corn plants observed during 1998/1999 season, the first season of

314

Fangneng Huang

planting Bt corn in South Africa, is a good indication of non-high dose Cry protein expression of these Bt corn hybrids against B. fusca. In addition, field selection pressures were very high in both cases where resistance was developed. In South Africa, for practical reasons, corn producers do not like to plant non-Bt refuge corn inside the irrigated area, which could have contributed to increase selection pressure towards the development of the field resistance (van Rensburg 2007). 18.4.3

Resistance to Bt cotton in H. zea in the southern region of the United States

It has been documented that both the first generation of Bt corn (YieldGard® corn) and Bt cotton (Bollgard® cotton) do not express a high dose of Cry proteins against H. zea, a major lepidopteran pest of corn and cotton in the United States. In the southern region of this country, H. zea feeds on wild hosts and/or corn for two generations during spring and summer seasons. After corn is harvested, it moves to cotton for 2 to 3 additional generations (US EPA 2001). Because both Bt corn and Bt cotton do not produce a high dose of Bt proteins for this insect species, H. zea could feed on Bt corn plants first and the survivors could move to Bt cotton fields and continue exposure to Bt proteins. This local movement of H. zea could create a multiple-generation of exposure to Bt proteins within a same crop growing season (US EPA 2001). Therefore, relative to the North Central Corn Belt of the United States, a greater percent of non-Bt corn refuge size is required by the United States Environmental Protection Agency (U.S. EPA) for IRM in the southern region where corn and cotton are overlapped. Laboratory bioassays showed that field populations of H. zea collected from Bt corn and Bt cotton fields were less susceptible to Cry1Ac toxin compared to insects collected from non-Bt crops (Ali et al. 2006; Ali and Luttrell 2007), suggesting a possible shift in resistance frequency. A slight decrease in susceptibility of H. zea to Cry1Ac also was reported during the first three years of commercial use of Bt cotton in Mississippi Delta area where Bt cotton had been widely planted (Hardee et al. 2001). These data were recently interpreted as documents for field resistance development to Bt cotton in H. zea (Tabashnik et al. 2008). However, the conclusion of this field evolved resistance is questionable because field control failures or reduced efficacies of Bt cotton due to resistance development have not been reported in this area (Moar et al. 2008). 18.4.4

Resistance to Cry1Ac cotton in H. armigera in north China

H. armigera is the most economically important insect pest of cotton in China (Wu 2007). It has been difficult to control this pest with chemical insecticides because of its high level of resistance to various classes of insecticides (Wu and Guo 2005). Bt cotton was first commercialized during 1996/1997 in China (James 2006). Like the three cases of resistance development described above,


315

Cry1Ac cotton varieties planted in China also does not produce a high dose of Bt proteins against H. armigera. Nevertheless, adoption of Bt cotton in China has increased dramatically, up to 3.6–3.8 million ha annually (66 to 68% of its total cotton area) during 2004–2008 (James 2008). Transgenic Bt cotton technology has been the primary tool for managing H. armigera in China during the last decade. A nine-year (1999–2007) monitoring of Bt resistance indicated that resistance allele frequency to Cry1Ac cotton in a Qiuxian County population of H. armigera in north China increased from 0.0058 in 1999 to 0.091 in 2007 (Xu et al. 2009; Liu et al. 2009). Bioassays with purified Bt toxin also demonstrated that populations collected from Bt corn plants in 2007 were significantly less susceptible to Cry1Ac toxin than a laboratory susceptible colony. In addition, field survey showed that the number of H. armigera eggs laid on Bt cotton plants increased consistently from 2003 to 2007 (Liu et al. 2009). These results strongly suggest that field resistance to Bt cotton in H. armigera might already occur or is developing in Qiuxian area. However, field control failures due to resistance development have not yet been reported from this area. Bt microbial insecticide sprays was one of the major tools for controlling H. armigera in Qiuxian County prior to the use of transgenic Bt cotton. In 1998, Cry1Ac cotton was first introduced to this region for controlling this devastative cotton pest (Xu et al. 2009). Since then, Bt cotton growing area had increased rapidly during the first three years, approached 100% in 2001and remained the 100% adoption level during 2002–2007. The long-term and large-scale adoption of Bt corn likely has placed a heavy selection pressure on H. armigera populations in this area (Xu et al. 2009). Requiring farmers to plant refuge is difficult in China because of the challenges associated with educating and monitoring>10 million small-cotton growers (Wu 2007). IRM plans for Bt cotton growers are therefore not required in China. In addition, the partial survival of H. armigera populations due to non-high dose expression of Bt proteins in the cotton varieties might also enhance the selection for resistance development.

18.5 Bt resistance allele frequency in target pest species of Bt corn and Bt cotton Resistance monitoring is a key component for a successful IRM plan. Resistance monitoring techniques for Bt corp IPM should be able to detect resistance allele frequency at very low levels (e.g. 0.001). Therefore, proactive actions can be implemented before field-evolved resistance occurs (Andow and Alstad 1998; Huang 2006). However, monitoring of rare resistance alleles in field populations has been a challenge. Traditional methods for monitoring insecticide susceptibility do not have the sensitivity for identifying rare resistance alleles. During the past decade, several techniques have been evaluated for this purpose (Andow and Alstad 1998; Zhao et al. 2002, Bourguet et al. 2003; Huang 2006). Among these, the F2/F1 generation screening procedures (Gould et al. 1997; Andow and Alstad

316

Fangneng Huang

1998; Bourguet et al. 2003; Yue et al. 2008; Xu et al. 2009) and the DNA-marker method (Tabashnik et al. 2006; Gahan et al. 2007) have demonstrated the ability for identifying rare resistance alleles in field insect populations. By using these novel methods, Bt resistance allele frequencies have been determined for several insect species targeted by Bt corn or Bt cotton (Huang 2006). 18.5.1

Bt resistance allele frequency in target species of Bt corn

For target species of Bt corn, resistance allele frequency has been estimated for several stalk boring species including O. nubilalis in the United States and Europe (Andow and Alstad 1998; Andow et al. 1998, 2000; Bourguet et al. 2003; Farinós et al. 2004; Kaiser-Alexnat et al.2005; Stodola et al. 2006), D. grandiosella in Louisiana (USA) (Huang et al. 2007b), D. saccharalis in Louisiana and Texas (USA) (Huang et al. 2007a, 2008, 2009; Huang and Leonard 2008; Yue et al. 2008) and Sesamia nonagrioides in Greece and Spain (Andreadis et al. 2007). Resistance genes that allowed resistant insects to complete larval stages on Cry1Ab corn (e.g. YieldGard® corn) have been documented in three Louisiana populations of D. saccharalis. Major resistance genes to Bt corn have not been detected in any other target species, except B. fusca in south Africa as described above. The global monitoring results demonstrate that Bt resistance allele frequencies in target species of Bt corn are low, especially for the primary target, O. nubilalis. 18.5.2

Bt resistance allele frequency in target species of Bt cotton

For the major lepidopteran pests targeted by Bt cotton, Bt resistance frequency has been estimated for H. virescens in the southern United States (Gould et al. 1997; Gahan et al. 2007), P. gossypiella in Arizona, California, and Texas (USA) (Patin et al. 1999; Tabashnik et al. 2000, 2006), H. zea in North Carolina (USA) (Burd et al. 2003), and H. armigera in Australia (Akhurst et al. 2003; Gunning et al. 2005) and China (Li et al. 2004; Xu et al. 2009). Major resistance alleles to Cry1Ac cotton have been identified in H. virescens in southern United States, P. gossypiella in southwestern United States, H. zea in North Carolina, and H. armigera in Australia and China. Bt resistance alleles in H. virescens, the primary target of Bt cotton in the United States, was estimated to be very rare (Gould et al. 1997; Gahan et al. 2007). Resistance allele frequency in the Arizona populations of P. gossypiella was originally estimated to be as high as 0.16 during 1997 (Tabashnik et al. 2000), but a recent study by using the DNA screening method revised the Bt resistance allele frequency to be very low in Arizona, California, and Texas (Tabashnik et al. 2006). Bt resistance alleles in the North Carolina (USA) populations of H. zea was also estimated to be rare (Burd et al. 2003). However, Bt resistance allele frequency in H. armigera in Australia and China appears to be greater than that of the species described above. Resistance allele


317

frequency in H. armigera in Qiuxian County area (China) was reported to be 0.0058 in 1999 and 0.091 in 2007 (Xu et al. 2009; Liu et al. 2009). In Australia, Downes et al. (2007) screened >400 feral individuals of H. armigera during 2002–2006 and identified 7 individuals carrying resistance alleles to Cry2Ab, a Cry protein expressed in Bollgard II® cotton, but no insects possess resistance alleles to Cry1Ac.

18.6

Inheritance of resistance to Bt crops

Inheritance characters of Bt resistance have been evaluated for several target species of Bt corn or Bt cotton. Published studies showed that high levels of resistance to Bt plants was often controlled by a single recessive or incompletely recessive gene (Tabashnik and Carrière 2007). However, at low toxin concentrations, resistance can be more dominant when it is measured as effective dominance (Liu and Tabashnik 1997; Liu et al. 2001). For example, our current studies showed that Cry1Ab resistance of D. saccharalis was completely recessive at 128 μg/g, but it was incompletely recessive at 1 μg/g and nearly completely dominant at 0.016 μg/g (Wu et al. 2009b). Similar results were also reported in P. gossypiella (Liu et al. 2001; Tabashnik et al. 2002), H. virescens (Gould et al. 1992, 1995) and H. armigera (Kranthi et al. 2006; Liang et al. 2008). The toxin-concentration-depended dominance suggests that increasing expression levels of Bt proteins in plants could make a genetically non-recessive resistance become functionally recessive.

18.7

Fitness of Bt resistance

Knowledge on fitness of resistant insects is useful in developing IRM strategies (Gassmann et al. 2009). For resistance associated with fitness costs, especially for dominant fitness costs, resistance evolution could be delayed or even be reversed in absence of selection pressure (Tabashnik et al. 2005; Gassmann et al. 2009). Fitness of Bt resistance has been evaluated for several insect species targeted by Bt crops (Akhurst 2003; Snow et al. 2003; Bird and Akhurst 2004; Cerda and Wright 2004; Vacher et al. 2004; Carrière et al. 2005, 2006; Higginson et al. 2005; Raymond et al. 2005; Anilkumar et al. 2008; Gassmann et al. 2009). Published data showed that Bt resistance is often associated with fitness costs, but most of the costs are recessively inherited (Anilkumar et al. 2008). Sometimes, fitness costs of Bt resistance could be interacted with environmental factors (Carrière et al. 2004; Carrière et al. 2005; Higginson et al. 2005; Gassmann et al. 2006; Janmaat and Myers 2005, 2006; Raymond et al. 2005, 2006, 2007; Bird and Akhurst 2007; Raymond et al. 2007). However, our recent study indicated that a lack of fitness costs was associated with Cry1Ab resistance in D. saccharalis (Wu et al. 2009c).

318

18.8

Fangneng Huang

Incomplete resistance

Resistance to Bt plants has been found to be incomplete in most cases where it has been investigated. Compared to on non-Bt plants, resistant insects on Bt plants may develop slower, have a greater mortality, or produce less offspring (Liu et al. 1999; Bird and Akhurst 2004; Huang et al. 2007a; Van Rensburg 2007). For examples, relative to susceptible larvae on non-Bt cotton, resistant P. gossypiella on Bt cotton plants required an average of 5.7 d longer to develop to the pupal stage (Liu et al. 1999) and had a lower survival rate and smaller pupal weight (Carrière et al. 2006). Bird and Akhurst (2004) reported that life history parameters of resistant H. armigera larvae on Bt cotton plants also had a significantly developmental delay compared to the susceptible strain on non-Bt plants. They also found survival of resistant larvae on Bt cotton plants was>50% lower than on non-Bt cotton. Recently, Huang et al. (2007a) also reported that neonates of Bt resistant D. saccharalis on Cry1Ab corn plants took 20 to 30 more d to reach the pupal stage than on non-Bt plants. Similar results also were observed for the field-evolved resistant B. fusca (van Rensburg 2007). The considerable lower fitness of resistant populations on Bt plants than on non-Bt plants is often referred as a result of incomplete resistance (Tabashnik and Carrière 2007). This kind of incomplete resistance would be important for resistance management if it has a pleiotropic effect of the resistance alleles that should slow down resistance development in the field (Carrière et al. 2006; Huang et al. 2007a).

18.9

Cross-resistance

Cross-resistance has been investigated in several target insect species of Bt corn and Bt cotton. Cross-resistance is common among Bt toxins (Bauer 1995; Gould e t a l . 1 9 9 5 ; Tabashnik et al. 2000; Zho et al. 2001; Ferré and van Rie 2002; Li et al. 2005a; Ali et al. 2007; Wu et al. 2009a). However, several studies also demonstrated that no cross-resistance is existed among Bt toxins that are already presented in Bt corn or Bt cotton. For example, a Cry1Ab resistant strain of O. nubilalis was not resistant to Cry9C and had only a very low level of crossresistance to the Cry1F protein (Siqueira et al. 2004). Jackson et al. (2007) also reported that Cry1Ac resistant H. virescens did not resist to a vegetative Bt insecticidal protein, Vip3A, which has been presented in the recently released VipCot Bt cotton (Table 2). Similarly, a Cry2Ab resistant strain of H. armigers was not resistant to the purified Cry1Ac toxin or Cry1Ac cotton plants (Mahon et al. 2007). Our studies also showed that a Cry1Ab resistant D. saccharalis strain was susceptible to Cry2Ab2 toxin (Wu et al. 2009a). These results provide valuable information to support the use of a “gene pyramiding” approach for Bt resistance management (Moar and Anilkumar 2007). Unfortunately, it is impossible to establish any models for predicting crossresistance because cross-resistance patterns can vary for different insect species,


319

Bt toxins, and even among insect strains of a same species (Bauer 1995; Zhao et al. 2001; Ferré and Van Rie 2002; Ali et al. 2007). One of the most likely underlying mechanisms of the variable cross-resistance patterns may be associated with the specific binding sites for Bt toxins in the insect midgut. However, for target species of Bt corn or Bt cotton, specific Bt binding sites are still not well understood. In the diamondback moth, Plutella xylostella, a major vegetable insect pest, probably four Bt binding sites are existed in the midgut (Ferré and Van Rie 2002).

18.10

Mechanisms of Bt resistance

The proposed mode of action for Bt toxins involves sequential events, including solubilization and activation of protoxins, specific binding of active toxins, and formation of nonselective pores (Gill et al. 1992). Theoretically, any changes in insect gut physiology and/or biochemistry that affect one or more steps in this process could interfere with toxicity and lead to development of resistance. However, up to date, most of the studies on Bt resistance mechanisms have focused on two steps in the mode of action: proteolytic activation of Cry protoxins and binding of Bt toxins to receptors in midgut. Currently, the mechanism of Bt resistance has been evaluated in three major cotton pests, H. virescens, (Heckel et al.1997; Gahan et al. 2001; Jurat-Fuentes et al. 2002, 2003; Morin et al. 2003; Jurat-Fuentes and Adang 2004, 2006) and P. gossypiella (Morin et al. 2003; Gonzalez-Cabrera et al. 2003; Tabashnik et al. 2005), and H. armigera (Xu et al. 2005). The molecular basis of Bt resistance has not been well understood (Gahan et al. 2001). The most common mechanism in these three cotton insect species was due to a reduction in binding of toxins to cadherin receptors in midgut. In addition, resistance mechanisms also have been investigated in P. xylostella, O. nubilalis, and the Indianmeal moth, Plodia interpunctella. Resistance in some strains of these three species was due to changes in binding affinity (Ferré and van Rie 2002; Herrero et al. 2001; Baxter et al. 2005), while for some strains, resistance could be associated with changes in midgut proteinase activity (Oppert et al.1997; Huang et al. 1999b; Li et al. 2005b). Other mechanisms or multiple mechanisms could also play an important rule in Bt resistance evolution (Griffitts et al. 2001; Jurat-Fuentes et al. 2003; Gunning et al. 2005).

18.11

Current IRM strategies for Bt crops

Since Bt corps were first commercially used in 1996, mandatory IRM plans have been successfully implemented for planting Bt crops in the United States, Canada, and Australia. Basically, two different strategies are currently used for managing insect resistance to Bt crops in these countries: 1) a high dose/refuge strategy and 2) a gene pyramiding method.

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High dose/refuge strategy

To delay resistance development to Bt crops, a “high dose/refuge” IRM strategy (Ostlie et al. 1997) has been used in the United States, Canada, and Australia. This IRM strategy involves planting high dose Bt plants in a portion of an area that can kill heterozygous resistant insects. The remaining area is planted with non-Bt plants that can serve as a refuge for Bt-susceptible insects. The relatively large populations of Bt-susceptible insects from refuge areas should mate with the rare surviving resistant homozygotes. Thus, most of their offspring that carry any resistance alleles will be heterozygous for Bt resistance. Since heterozygotes should be killed by high dose Bt plants, resistance frequencies in the target insect populations should be maintained at low levels for a long period of time (Ostlie et al. 1997). To be effective for the “high dose/refuge” IRM strategy, there are four key assumptions that must be met. First, Bt crops must produce a sufficiently high dose of Bt proteins that is able to kill heterozygous resistant individuals. Second, resistance is controlled by one recessive or at least partial recessive gene. Third, initial resistance alleles frequency in field insect populations is very low (e.g. < 0.001). Finally, resistant individuals survived from Bt corn plants should random mate with susceptible insects from non-Bt refuge plants (US EPA 2001). The “high dose/refuge” strategy has been implemented successfully in the United States and Canada. A two-year survey conducted in the United States indicated that >85% growers complied with the refuge requirements (Agricultural Biotechnology Stewardship Technical Committee 2002). Canadian surveys during 2001–2007 also demonstrated that >80% growers followed the refuge requirements (Canadian Corn Pest Coalition 2005, 2007). The relatively high compliance rates in the United States and Canada suggest that the “high dose/ refuge” IRM plan is feasible for their growers. The successful implementation of the “high dose/refuge” strategy might have contributed a significant role for the long-term success of Bt crops in the north America. A similar IRM strategy has also been recommended in Australia and India for planting Bt cotton (APCoAb 2006), but the compliance levels for the two countries are unknown. 18.11.2

Gene pyramiding

Another IRM strategy currently used for Bt crop resistance management is called “gene pyramiding” method. This technique relies on transferring two or more Bt genes with different mode of actions into the same plants such that resistance to one Bt protein should not resist to other toxins (Moar and Anilkumar 2007). In order to delay resistance development, the gene pyramiding approach requires lack of cross-resistance among the toxins pyramided. Modeling has showed that resistance development to Bt crops with pyramided genes could be significantly delayed compared to the use of single gene expressed Bt crops (Onstad and Gould 1998; US EPA 2001). Because of the unpredictability of cross-resistance patterns, careful selection of different Bt genes without cross-resistance for pyramiding is


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essential to ensure the success of this strategy for managing Bt resistance. The gene pyramiding method could be considered as a primary IRM strategy for developing countries because implementation of this strategy is not necessary to relay on their tens of millions of small-sized crop farmers. The gene pyramiding strategy has been used for managing resistance to Bt cotton in the United States and Australia since 2003 when Bollgard II® became commercially available. The Bollgrad II ® contains two different Bt genes, Cry1Ac and Cry2Ab2. The two proteins act independently by binding to different receptors in the insect midgut (Moar and Anilkumar 2007). Most of the currently planted Bt cotton in the United States contains the Bollgrad II traits. In Australia, all Bt cotton varieties planted have been the Bollgrad II® since the 2004/2005 growing season (Farrell 2006). The gene pyramiding strategy is expected to be used for managing resistance to Bt corn in the United States and Canada during 2009. Recently, genes encoding for Cry1A.105 and Cry2Ab2 proteins have been transferred into corn hybrids using the VecTran technology (Monsanto 2007). One of the transgenic products produced with this technology is the event MON89034 corn. This event contains both Cry1A.105 and Cry2Ab2 genes and is expected to offer a valuable IRM tool (Monsanto 2007). MON89034 corn has been approved by U.S. EPA for commercial use beginning in 2009 under the trade name of YieldGard VT ProTM (Wu et al. 2009a). Our recent bioassays exhibited that a Cry1Ab resistant D. saccharalis strain demonstrated only a relatively low cross-resistance (4-fold) to Cry1A.105 protein and no cross-resistance to Cry2Ab2 (Wu et al. 2009a). Our on-going studies also showed that MON89034 corn plants can completely overcome our Cry1Ab corn resistant D. saccharalis in greenhouse tests (MG, FH unpublished data). These results provide a showcase that Bt crops with pyramided genes may be a very useful tool to delay resistance development. 18.11.3

Other IRM strategies suggested for Bt crops

Several other strategies have also been suggested for managing resistance for Bt crops. As discussed above, studies have showed that high levels of Bt resistance are often associated with fitness costs. If this is true, rotating Bt varieties with non-Bt varieties could be a useful strategy for resistance management, especially if the fitness cost is dominant. A dominant fitness cost would allow resistance gene frequency to decrease in the field when selection pressure is removed (Tabashnik and Carrière 2007). Another suggested strategy is to rotate among Bt corn hybrids that express different proteins with different mode of action (Roush 1996). To be effective, this strategy also requires a lack of cross-resistance among the toxins and may work better if fitness costs are involved with resistance. In addition, studies have showed that Bt resistance could be caused by reducing midgut proteinase activity, which convert Bt protoxins to active toxins (Oppert et al. 1997; Huang et al. 1999b; Li et al. 2007). This type of resistance should be managed by planting Bt crops that express activated Bt proteins (Huang et al. 1997).

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18.12 Integration of Bt crop technology within integrated pest management programs Bt crops should be considered as only one component of integrated pest management (IPM) programs and should be incorporated in IPM programs (Kennedy 2008). Combining IRM with IPM programs should be the best strategy for managing Bt resistance, because IPM can diversify mortality factors and thus should reduce selection pressure from any single mortality mechanism (McGaughey and Whalon 1992).

18.13

Prospect

Since 1996 when transgenic Bt crops became commercially available, adoption of this technology has increased rapidly in the world. Crop growers have recognized great benefits from planting Bt crops. Resistance development in target insect pests has been considered as a major threat for the long-term success of Bt crop technology. Major resistance alleles have been detected in several target species, but field resistance that led to control failures has been documented in only two isolated cases. In both cases, the Bt corn hybrids planted apparently did not express a high dose of Bt proteins and, in the meantime, a very high selection pressure was involved for the resistance development. After more than a decade of commercial use of Bt crops, resistance allele frequency in the field remains relatively low in most target pest species, except for H. armigera in Australia and China. The great success in use of Bt crops for more than a decade long will certainly encourage more countries and more crop growers to use this technology. Recently, a new Bt corn, SmartStaxTM corn, which contains eight transferredgenes, is expected to become commercially available during 2010 in the United States (Biosafety Clearing-House 2008). Two of the eight genes are for herbicide resistance and the other six are Bt Cry genes for resistance to insect. By planting this novel gene-stacked/pyramided Bt corn, non-Bt refuge size required for IRM in the United States is expected to be reduced significantly, from current level of 20% to 5% in areas where cotton is not planted and from 50% to 20% in the corn and cotton overlapping areas. In addition, an approach of mixing Bt and non-Bt seeds for providing refuge plants, known as refuge in the bag (RIB), is expected to be approved by U.S. EPA during 2009 for planting Bt corn for managing corn rootworms (Benbrook 2009). The use of RIB approach will eliminate the requirement to plant non-Bt corn refuge varieties separately from Bt corn areas and thus should greatly facilitate growers in use of Bt crops. Moreover, Bt rice is expected to become commercially available in China in the near future (James 2008; Huang et al. 2005). Commercial use of Bt rice in China will have a great influence on the global adoption of Bt crop technologies. Furthermore, the current global food crisis and the rapid increase in demand for fuel energy and food make the needs of


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such great pest management technologies even more urgently. It is, therefore, widely expected that the use of transgenic Bt crop technologies for managing insect pests will continue to increase rapidly in the future, especially in the developing countries. IRM for Bt crops will be more complicated and continue to be a big challenge. Nevertheless, we believe that the knowledge and experience gained during the past thirteen years of the successful commercialization of Bt crops should make us more confident for the sustainable use of transgenic Bt crop technologies in the future. Acknowledgements I thank Drs. Lixin Mao, Mukti Ghimire, and Zhaorigetu Chen for reviewing the manuscript. This article is published with the approval of the Director of the Louisiana Agricultural Experiment Station as manuscript number No. 2009-234-3252.

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Tabashnik B E, Gassmann A J, Crowder D W, Carrière Y. Insect resistance to Bt crops: evidence versus theory. Nature Biotech., 2008, 26: 199–202. US EPA (United States Environmental Protection Agency). 2001. Biopesticides registration action document- Bacillus thuringiensis plant-incorporated protectants. E: Benefit assessment. http:// www. epa.gov/oppbppd1/biopesticides/pips/Bt_brad2/ 5-benefits.pdf. [Last accessed 24 April 2007]. Vacher C, Bourguet D, Rousset F, et al. High dose refuge strategies and genetically modified cropsreply to Tabashnik et al. J. Evol. Biol., 2004, 17: 913–918. van Rensburg J B J. First report of field resistance by the stem borer, Busseola fusca (Fuller) to Btresistance maize. S. Afr. J. Plant Soil., 2007, 24: 147–151. vanWyk A, van den Berg J, Van Rensburg J B J. Comparative efficacy of Bt maize events Mon810 and Bt11 against Sesamia calamistis (Lepidoptera: Noctuidae) in South Africa. Crop Protect., 2008, 28: 113–113. Whalon M E, McGaughey W H. Insect resistance to Bacillus thuringiensis. // Kim L. Advanced Engineered Pesticides. New York: Marcel Dekker, 1993: 215–231. Whalon M E, McGaughey W H. Bacillus thuringiensis: use and resistance management. // Ishaaya I andDegheele D. Insecticides with Novel Modes of Action. New York: Springer, 1998:106–137. Wu K M, Guo Y Y. The evolution of cotton pest management practices in China. Ann. Rev. Entomol., 2005, 50: 31–52. Wu K M. Environmental impacts and risk management strategies of Bt cotton commercialization in China. J. Agri. Biotech., 2007, 15: 1–4. Wu X, Leonard B R, Zhu Y C, et al. Susceptibility of Cry1Ab-resistant and-susceptible sugarcane borer (Lepidoptera: Crambidae) to four Bacillus thuringiensis toxins. J. Invert. Pathol., 2009a, 100: 29–34. Wu X, Huang F, Leonard B R, et al. Inheritance of resistance to Bacillus thuringiensis Cry1Ab protein in the sugarcane borer (Lepidoptera: Crambidae. J. Invert. Pathol., 2009b, 102: 44–49. Wu X, Huang F, Leonard B R, et al. Growth and development of Bacillus thuringiensis Cry1Absusceptible and - resistant sugarcane borer on diet and conventional corn plants. Entomol. Exp. App., 2009c, 133: 199–207. Xu X, Yu L, Wu Y. Disruption of a cadherin gene associated with resistance to Cry1Ac δ-endotoxin of Bacillus thuringiensis in Helicoverpa armigera. App. Environ. Microbiol., 2005, 71: 948– 954. Xu Z, Liu F, Chen J, et al. Using F2 screen to monitor resistance allele frequency to Bt cotton in field populations of Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae). Pest Manag. Sci., 2009, 65: 391–397. Yue B, Huang F, Leonard B R, et al. Verifying an F1 screen for identification and quantification of rare Bacillus thuringiensis resistance alleles in field populations of sugarcane borer (Lepidoptera: Crambidae) Entomol. Exp. App., 2008, 129: 172–180. Zhao J Z, Li Y X, Collins H L, et al. Different cross-resistance patterns in the diamondback moth (Lepidoptera: Plutellidae) resistance to bacillus thuringiensis toxin Cry1Ac. J. Econ. Entomol., 2001, 94: 1547–1552. Zhao J Z, Li Y X, Collins H L, et al. Examination of the F2 screen for rare resistance alleles to Bacillus thuringiensis toxins in the diamondback moth (Lepidoptera: Plutellidae). J. Econ. Entomol., 2002, 95: 14–21.

CHAPTER 19

Potential Use of Proteinase Inhibitors, Avidin, and other Bio-reagents for Synergizing Bt Performance and Delaying Resistance Development to Bt Yucheng Zhu, Mingshun Chen and Craig A. Abel

Abstract After being ingested by target insects, the insecticidal proteins from Bacillus thuringiensis (Bt) need to go through a proteolytic process by insect midgut proteinases to become activated. At the same time, Bt can be hydrolyzed and degraded by midgut proteinases to become non-toxic to target insects. Once activated, the Bt proteins need to bind to midgut brush border membrane vesicle (BBMV) to cause gut lining lesions and eventually death in the target insect. A few bio-reagents may interact with the Bt binding to the receptors. By applying proteinase inhibitors to Bt-containing (sublethal dose) diet, the growth and development of Helicoverpa zea were significantly decreased when compared with the Bt only control. Midgut samples tested against the substrates for major midgut enzymes showed significant decreases in the protease activity of larvae fed Bt plus inhibitor versus control. Avidin, causing sequestration of biotin and vitamin deficiency, potentially interacts with Bt by binding to biotin-containing proteins. Besides possessing insecticidal toxicity itself, avidin at a sublethal dose could significantly synergize Bt toxicity against H. zea larvae. Because of different modes of action from that of Bt, proteinase inhibitors, avidin, and other bio-reagents could be used to enhance Bt performance, delay resistance development to Bt, and expand control range beyond lepidopterans. Keywords Proteinase Inhibit, Bio-reagents, Synergizing Bt, resistance, cotton bollworm Transgenic cotton, which produces toxins from the soil bacterium, Bacillus thuringiensis Berliner, effectively controls many economically important insects. Yucheng Zhu, Craig A. Abel Southern Insect Management Research Unit, USDA-ARS, Stoneville, MS, 38776, USA E-mail: [email protected] Mingshun Chen Department of Entomology, Kansas State University, USDA-ARS, Manhattan, KS, 66506, USA


Potential Use of Proteinase Inhibitors, Avidin, and other Bio-reagents

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Since the first introduction into the U.S. in 1995 (Benedict and Altman 2001), Bt cotton has been widely used to reduce feeding damage from many lepidopteran pests. Right now, Bt cotton has become the dominant cotton variety in the Midsouth of the US (Williams 2005). In current cotton systems, large scale deployment of Bt transgenes and the high efficacy of transgenic Bt plants against target pest insects impose high selection pressure on naturally existing Bt-resistant genes to increase their frequencies in the populations. However, the potential evolution of Bt resistance in lepidopteran cotton pests (Gould et al. 1997; Gahan et al. 2001) is a potential threat which could rapidly decrease the value of this Bt biotechnology. Bt resistance is attainable in lepidopteran insects and a number of insect species have developed resistance through laboratory selection (Tabashnik 1994), including the tobacco budworm with over 10,000-fold resistance (Gould et al. 1995) and the pink bollworm Pectinophora gossypiella (Saunders) with over 3,100-fold resistance (Tabashnik et al. 2002). Resistance to transgenic Bt cotton has also been observed in Helicoverpa armigera (Hübner) (Meng et al. 2004). If resistance management strategies are not developed and properly carried out, the effectiveness of this environmentally sound method of pest control could eventually be negated. To prolong the benefit of this biotechnology, alternative control measures should be developed to relieve selection pressure and slow down resistance development among many lepidopteran insects. Avidin is a bioactive glycoprotein found naturally in the egg white of bird, reptile, and amphibian eggs. Chicken egg white contains up to 500 ppm of avidin, which tends to be less stable and vulnerable to degradation by proteinases under acidic conditions in the human stomach (Kramer 2004). Transgenic corn seeds contain as much as 3000 ppm avidin (Kramer 2004). Avidin corn was not found to be toxic to mice when administered as the sole component of their diet for 21 days (Kramer 2000). Avidin has a very strong affinity for the vitamin biotin, which is a coenzyme required for enzymes that catalyze carboxylation, decarboxylation, and transcarboxylation reactions in all forms of life (Kramer 2004). Avidin has a long history of use in a variety of biochemical and medical diagnostic procedures. Insecticidal activity of chicken avidin has been known since 1959 (Levinson and Bergmann 1959). Sequestration of biotin causes vitamin deficiency and in turn leads to stunted growth and mortality of many insect species. Transgenic avidin corn (Kramer et al. 2000), tobacco (Markwick et al. 2003), and apple (Markwick et al. 2003) plants showed strong resistance and insecticidal activities against many field crop and stored product insects. However, using avidin has not been attempted previously to control cotton insects. By providing a different mode of action, the incorporation of an avidinproducing gene into the Bt cotton genome could certainly expand control ranges and delay resistance development to Bt. The major functions of the gut proteinases include hydrolyzation of ingested protein into peptides during the initial stages of protein catabolism. Because

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proteins are critical nutrients for insect growth and development, proteinases, such as trypsin, play an essential function for protein digestion and absorption in insects. Midgut proteinases are also involved in Bt protoxin activation, which is a necessary step for Bt proteins to achieve insecticidal function. Activated Bt toxins may be subjected to further proteolysis and degradation by gut proteinases to form non-toxic segments. One of the proposed Bt resistance mechanisms indicated that reduced sensitivity to Bt is associated with the absence of proteolytic activity or an activation process that is present in the wild type strain (Oppert et al. 1996; Oppert et al. 1997; Li et al. 2004; Li et al. 2005), or excessive degradation by gut proteinases in resistant strains or less sensitive stages (Forcada et al. 1996; Keller et al. 1996; Shao et al. 1998). Currently, Bt toxins used for cotton insect control have a narrow range against lepidopteran pests only. Due to large scale adoption of Bt cotton and reduced chemical applications, many originally secondary pests, such as the tarnished plant bug, Lygus lineolaris, and stink bugs, have emerged causing serious economic losses to cotton production. Because the gut proteinases play an important biochemical role in insect growth and development, these enzymes have been targets for proteinaceous inhibitors whose genes have been incorporated into transformed plants (Hilder and Boulter 1999). Jongsma (2004) recently proposed use of novel proteinase inhibitor genes to control sucking mouthpart pests. The introduction of proteinase inhibitors into host plants may substantially suppress protein digestion, and subsequently achieve insect control in a broad range through nutrient deficiency. In addition, introduction of proteinase inhibitors into the gut will certainly modify biochemical balance within target insects feeding on Bt cotton. Bt toxins may become stable and more effective against target insects. This report includes a review of previous studies (Zhu et al. 2005, 2007) to show the potential use of avidin for controlling a wide range of lepidopteran pests, and how avidin and proteinase inhibitors interact with Bt Cry1Ac toxin and on larval growth, development, and midgut proteinase activities in H. zea. New results and discussion are also included to expand assessments of the bioreagents to 5 lepidopteran species for their insecticidal activities and potential synergism to Bt.

19.1

Experimental procedures

Insects. Five lepidopteran species were selected for bioassays with avidin or proteinase inhibitors. These included: bollworms, H. zea; the tobacco budworm, Heliothis virescens (F.); the fall armyworm, Spodoptera frugiperda J.E. Smith; beet armyworms, Spodoptera exigua (Hübner); and the velvetbean caterpillar, Anticarsia gemmatalis (Hübner). Insect artificial diet and bio-reagents. Tobacco budworm (Heliothis


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virescens) diet (#F9915B), USP agar (#7060) and USDA vitamin premix (#6265) were obtained from Bio-Serve (Frenchtown, NJ). Low melting point agarose (Invitrogen, Ultrapure L.M.P Agarose) was used to replace regular agar in diets testing the effects of selected avidin and proteinase inhibitors. Avidin, benzamidine, phenylmethylsulfonyl fluoride (PMSF), and N-α-tosyl-L-lysine chloromethyl ketone (TLCK), azocasein, α-benzoyl-DL-arginine-p-nitroanilide (BApNA), and N-succinyl-alanine-alanine-proline-phenylalanine-p-nitroanilide (SAAPFpNA) were obtained from Sigma Chemical Company (St. Louis, MO). Examination of insecticidal potency of selected bio-reagents and their synergism to Bt. Avidin and proteinase inhibitors were individually incorporated into artificial diet to test toxicity against selected insect species. These bioreagents were also combined with Bt to examine their synergistic effect. Experiment procedures (Zhu et al. 2005, 2007) were summarized below. Avidin toxicity assay. To determine insecticidal activity of avidin against selected insect species, larvae were reared on artificial diet. Based on preliminary experiments, the artificial diet was supplemented with 7 concentrations of avidin (10, 15, 20, 40 60, 80, and 100 ppm) for assaying avidin toxicity to the bollworm, and was supplemented with two concentrations (10 and 100 ppm) for other insects tested. Diet was cooled in a water bath to 33oC before the appropriate avidin quantities were added. Larval mortality was initially determined at 5 days after treatment (DAT) and observed at 7 and 10 DAT. Three replications of ten larvae were examined for each treatment. For each replicate, larval mortality was corrected from the appropriate natural mortality using Abbott’s Formula (Abbott 1925). Means for each treatment were compared using analysis of variance and separated using the Fisher’s Protected Least Significant Difference procedure at the α = 0.05 level (PROC MIXED, Littell et al. 1996, SAS Institute 1997). Activity of Avidin and Synergy with Bt against the Bollworm. Because the bollworm is the major lepidopteran pest on Bt cotton (Williams 2005), the activity of avidin and its relationship with Bt was examined for this species. Bioassays (Zhu et al. 2005) were conducted for the bollworm as described above. In brief, avidin at 10 ppm was supplemented to the artificial diet containing 10 and 20 ppb of Bt protoxin (Cry1Ac protoxin from MVP II™) (Monsanto Corp. St. Louis, MO). The Bt concentrations were determined for the bollworm through preliminary experiments that demonstrated low mortality but substantial effects on larval weight (data not shown). Neonates were placed on diet supplemented with avidin only, Bt only, or avidin + Bt and monitored every 2-3d for changes in larval mortality and weight until either pupation or death. Bioassays with proteinase inhibitors. Newly molted 3rd instar larvae of H. zea were fed various artificial diets to monitor the effects of added proteinase inhibitors, Bt, and any synergistic effects of proteinase inhibitors with Bt. Inhibitor stock solution was prepared in ethanol for PMSF, and in d-H2O for the remaining inhibitors. Artificial diet, designed to monitor proteinase inhibitor effects, was made by using a low melting point agarose and cooled to 33oC in a water bath before inhibitors were added to the diet. The concentration of Bt

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remained constant at 15 ng/ml. The concentrations (w/v) for proteinase inhibitors in diet were as follows: benzamidine (0.5%), PMSF (0.02%), and TLCK (0.04%, 0.08%, 0.16%, and 0.32%). Larvae were regularly measured for changes in weight and length. Three repetitions of 6 larvae each were monitored until death or pupation occurred. Preparation of Midgut fluid. After two, four and six days of feeding on diets, midguts of larvae were dissected in cold 0.1M Tris-HCl, pH 8.0 over an ice block. Midguts were homogenized then centrifuged at 5,000 rpm for 5 minutes to remove debris. Supernatant was collected and protein concentration was determined by the Bradford method (Bradford 1976) using Coomassie Plus Protein Assay kit (Pierce, Rockford, IL), with BSA as the protein standard. Midgut enzyme solutions were diluted to 1 mg/ml. Proteinase Activity Assays. Total proteinase activity was measured with azocasein. Enzyme solution (5 μl) was mixed with 20 μl 200 mM tris-chloride, pH 8, 5 μl H2O, and 10 μl azocasein solution made in 0.05% SDS (sodium dodecyl sulfate). The reaction was allowed to run for 2.5 hr at room temperature. After incubation, 30 μl of 10% TCA (trichloro acetic acid) was added. Reactions were placed on ice for 30 minutes and centrifuged at 14,000g for 5 minutes to remove precipitated protein. After centrifugation, 60 μl of supernatant was added to 40 μl of 1M NaOH. Absorbance was measured at 405 nm. To study trypsin-like and chymotrypsin-like proteinase activities, the substrates BApNA and SAAPFpNA were used, respectively. For assays using these substrates, 5 μl enzyme solution was mixed with 45 μl universal pH buffer (Frugoni 1957), pH 8. Activities were determined by the addition of 50 μl BApNA (1 mg/ml) or SAAPFpNA (1 mg/ml) in Frugoni Buffer, pH 8.5 (final substrate concentration was 0.5 mg/ml). Absorbance at 405 nm was monitored for 15 minutes at 37oC with measurements taken every 15 seconds. Bt protoxin and three proteinase inhibitors, benzamidine, PMSF, and TLCK, were used to treat diet for feeding bollworm larvae. Gut enzymes were prepared after larvae were fed for 2, 4, and 6 days. Activities were analyzed using azocasein for general proteinase activity, BApNA for tryptic proteinase activity, and SAAPFpNA for chymotrypsin-like proteinase activity. Data analysis. Means for each treatment were compared using ANOVA and separated using the Fisher's protected least significant difference procedure at the α = 0.05 level (PROC GLM, SAS version 9.1, SAS Institute Inc. Cary, NC). Synergism is defined as the combined effect which is greater than the sum of their individual effects (Zhu et al. 2007).

19.2

Insecticidal potency and synergism to Bt

Effects of Avidin to Lepidopteran Species. Bioassay results (Zhu et al. 2005) showed that avidin is a very potent insecticide against the five selected lepidopteran pests. Adverse effects ranged from retarded insect growth with reduced larval body length and body weight at a sublethal dose or low


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concentration (10 ppm) to complete death (100% mortality) at a higher concentration. For all species tested, mortality was significantly higher (P < 0.05) when larvae were reared on diet containing 100 ppm when compared to 10 ppm at 7 and 10 DAT. By 10 DAT, larval mortality for all species approached 100% on diet containing 100 ppm avidin. Therefore, avidin toxicity and its quick action against target species seemed to be highly concentrationrelated. When the bollworm was selected for testing dose responses to avidin, the LC50 was 66.2 ppm at day 7. At day 10, the LC50 reduced to 22.6 ppm. The toxicity of avidin to all selected Lepidoptera species was similar. No significant differences (P>0.05) in larval mortality and no significant interaction between species and treatments (P>0.05) were observed across the species and among the two treatments (10 and 100 ppm). Synergism of avidin and Bt toxin. Sublethal doses of both avidin and Bt toxin were used for an examination of synergism of the bio-reagents. Zhu et al. (2005) found that avidin toxicity against the bollworm was synergized by the addition of Bt. A significant interaction (P < 0.05) was observed between time of evaluation and the treatments. At 5 and 7 DAT, no significant differences (P>0.05) in larval mortality were observed among the treatments. However, at 10-17 DAT, significant differences in larval mortality (P < 0.05) among the treatments was observed At 12, 14 and 17 DAT, larval mortality was significantly greater (P < 0.05) for avidin at 10 ppm containing Bt at both 10 or 20 ppb compared to all other treatments. At day 17, avidin (10 ppm) alone and Bt (10/20 ppb) alone resulted in 10% and 13/23% mortality, respectively. The synergism of Bt and avidin increased larval mortality to 63% for 10 ppm avidin + 10 ppb Bt and 67% for 10 ppm avidin + 20 ppb Bt treatment. In addition, larval weights were significantly different among the treatments for all time periods (P < 0.001). The treatments containing both avidin and Bt had numerically lower weights compared to treatments containing avidin or Bt alone, or the untreated control. Influences of Bt and benzamidine on insect growth and gut proteinase activities. Benzamidine acts as a competitive inhibitor by binding in the specificity pocket of the substrate. A previous study (Zhu et al. 2007) showed that benzamidine itself has certain insecticidal function against cotton bollworm. After feeding for 7 days, treatment with a combination of Bt and benzamidine significantly reduced body weight by almost 2-fold (P < 0.001) than the additive effect of Bt alone and benzamidine alone. The interaction of Bt and benzamidine was significant (P < 0.05). Larval body length changes were similar to the body weight changes. After feeding for 7 days, treatment with a combination of Bt and benzamidine also significantly reduced body length by more than 2-fold than the additive effects of Bt alone and benzamidine alone (P < 0.05). Midgut proteinase enzyme activity assays using 3 different substrates showed significant synergism of Bt and benzamidine. After 6 days, the Bt + benzamidine treatment reduced general proteinase activity by 1.5-fold of the additive effects of Bt alone and benzamidine alone. Similarly, Bt + benzamidine treatment suppressed approximately 1.54-fold more tryptic activity than the additive

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effects of Bt alone and benzamidine alone. Although significantly higher chymotrypsin activity was suppressed more by Bt + benzamidine treatment compared to Bt alone or benzamidine alone, the suppression was not greater than the additive effects of Bt alone plus benzamidine alone. Influences of Bt and PMSF on insect growth. PMSF (phenylmethylsulphonyl fluoride) is a serine protease inhibitor. PMSF binds specifically and selectively to the active serine residue in serine proteases. Results showed that treatments with Bt alone or Bt plus PMSF consistently reduced larval body weight and length after 2, 4 or 6 days of post treatment (Zhu et al. 2007). The body weight in the treatments of Bt alone and Bt + PMSF reached only half or less than half of the body weight of the control. PMSF alone had only a minor effect on larval body weight decrease. Interaction of Bt and PMSF on body weight at day 6 was significant (P < 0.05). Bt treatment alone and PMSF alone didn’t reduce body length substantially (less than 8%). However, the combination of Bt and PMSF significantly reduced body length by up to 34% (P < 0.01). Synergism of Bt and PMSF on midgut proteinase activities was also observed by Zhu et al. (2007). More than twice of the general proteinase activities were suppressed by Bt + PMSF than the additive effects of Bt alone and PMSF alone. Bt + PMSF treatment suppressed trypsin-like activity by 84%, which was close to the additive effects (88%) of Bt alone and PMSF alone. Approximately 1.3fold chymotrypsin-like activity was inhibited by Bt + PMSF compared with the additive suppression by Bt alone and PMSF alone. Influences of Bt and TLCK on insect growth. TLCK (Nα-Tosyl-Lyschloromethylketone$HCl) irreversibly inhibits trypsin-like serine proteases. It covalently binds with histidine residues at the active site of the enzymes and may bind to cysteine residues. Larval growth of H. zea was significantly retarded after feeding on diet treated with a combination of Bt and TLCK (Zhu et al. 2007). High concentrations of TLCK resulted in substantially adverse effects on larval growth and development. The picture (Fig. 19.1) shows feeding activities of H. virescens on diet treated with Bt and TLCK. The larvae, fed with diet containing Bt only (Panel B) and TLCK only (Panel E and F), showed active feeding and fast growth which were basically similar to control insects (Panel A). The larvae on diet treated with Bt + TLCK (Panel C and D) showed less feeding and slow growth. Zhu et al. (2007) also showed that Bt alone and 0.04% TLCK alone treatments had similar body weights with the control at day 8 after treatment. A combination of 0.04% TLCK with 15 ng/ml Bt significantly reduced body weight by 49%. At high concentrations (0.08%, 0.16%, and 0.32%), TLCK alone also induced significant reduction of larval body weight up to 76%. With the addition of 15 ng/ml Bt, the three highest concentrations of TLCK reduced body weight further. Interaction of TLCK and Bt on larval body weight was significant (P < 0.001). Similarly, the interaction of TLCK and Bt on larval body length was also significant (P < 0.05). With addition of 15 ng/ml Bt, TLCK reduced larval body length by up to 55%. Midgut proteinase activities were also substantially suppressed by TLCK.


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Fig. 19.1 Rearing plates to show visual feeding activities in different treatments. A: Control; B: Bt only at 15 ppb; C: 0.1% TLCK + 15 ppb Bt; D: 0.2% TLCK + 15 ppb Bt; E: 0.1% TLCK; F: 0.2% TLCK. (In E and F: larvae and diet in the bottom 2 rows were remove before the picture was taken).

Treatments with Bt alone and 0.04% TLCK alone did not alter general proteinase activity significantly. A TLCK only treatment at a concentration of 0.32% suppressed general proteinase activity by half when compared with the activity observed in control samples. Up to 67% of the activity was suppressed by the Bt + TLCK treatment. Interaction of Bt and TLCK was significant (P < 0.001). Tryptic activities were significantly suppressed. Up to 78% of the BApNAse activity was inhibited by the 0.32% TLCK alone treatment. With the addition of 15 ng/ml Bt, TLCK further suppressed tryptic activity, showing an increasing trend (up to 89% suppression), as its concentration increased from 0.8% to 0.32%. Chymotrypsin-like activity was also sensitive to Bt and TLCK treatments. TLCK alone inhibited 24%–61% with increased inhibition as TLCK concentration increased. The combination of Bt and TLCK suppressed more chymotrypsin activity (up to 69% suppression) than either of the individual treatments. Interaction of Bt and TLCK was significant (P < 0.01). Interaction of Bt and TLCK on insect development and mortality was also investigated (Zhu et al. 2007). TLCK at 0.04% and 0.08% levels did not delay insect development, and 100% of the larvae pupated on day 12 which was the same as for the control. For those larvae feeding on 0.16% TLCK diet, they spent 15 days longer (total 27 days) to reach 100% pupation. Most (approximately 85%) larvae pupated after feeding on 0.32% TLCK diet for 18 days. The Bt only treatment showed a certain delay (3 d) of larval development. Combination of Bt and TLCK delayed larval development even further. At day 12, less than 30% of the larvae developed to pupal stage (P < 0.0001). Interaction of Bt and TLCK was significant. Bt + 0.32% TLCK treatment maintained lower pupation (c.a. 30%) for 30 days, and only 80% of the larvae pupated after 42 days. Bt alone and certain concentrations (0.04%–0.16%) of TLCK alone treatments did not kill any larvae. As much as 10% of the larvae were killed by the 0.32% TLCK treatment.

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Bt + 0.04%–0.16% TLCK killed 3%–6% larvae. Bt combined with 0.32% TLCK resulted in 20% mortality.

19.3

Summary and discussion

Transgenic plants have had a profound impact on agriculture and the environment. The development of transgenic cotton varieties with insecticidal Bt genes has been one of the most successful applications of biotechnology research. Unlike conventional chemical applications, Bt cotton continuously maintains a toxic dose of Bt proteins, and delivers the toxins directly to the most sensitive stage of the target pest, i.e. once the insect hatches and starts to feed. After ingestion, Bt proteins are subjected to proteolytic processes by insect gut proteinases. Bt protoxins are activated mainly by trypsins and/or chymotrypsins. The activated toxins may bind directly to target sites on gut membrane and subsequently cause lysis of the gut and death of the insect. Intensive and longterm implementation of Bt cotton may cause target insects to adapt or to develop resistance to Bt. Limited research has been conducted to increase Bt toxicity against H. armigera and the diamondback moth, Plutella xylostella (L.) (Wang et al. 1999; Liu et al. 2000; Qiu et al. 2002). Up to a 3-fold increase in toxicity was obtained by the addition of inorganic additives. Currently, there is an urgency to seek out natural bio-reagents or bio-pesticides so that their genes can be inserted into the cotton genome to either synergize or enhance Bt performance and to increase the control range of transgenic cotton. Current studies attempt to explore the potential use of avidin and proteinase inhibitors to alter midgut biochemistry for stabilization of Bt toxin and facilitating Bt binding to midgut membrane. Through laboratory bioassays, we attempted to reveal potential toxicity of selected bio-reagents and to examine interaction and synergism of these bioreagents with Bt. Results showed that the growth of all three lepidopteran insects was greatly retarded by diet which contained avidin (Zhu et al. 2005). In addition, it was also observed significant activity of avidin against the fall armyworm and the tobacco budworm. Avidin in diet at a concentration as low as 40 ppm could kill up to 100% of the larvae. More importantly, avidin could synergize Cry1Ac Bt toxicity against the bollworm. The results (Zhu et al. 2005) were consistent with those of Burgess et al. (2002). Avidin is a bioactive protein, and the gene coding for its synthesis could be inserted into cotton (or other field crops) genomes alone or stacked with Bt genes. By targeting different sites, avidin and Bt together could be more effective in suppressing lepidopteran insects. Unlike Bt toxins, avidin targets different binding sites. In the current study, the synergistic effect of avidin with the Cry1Ac Bt toxin against the bollworm (also seen with Cry1Ba against H. armigera, Burgess et al. 2002) suggests strong potential for stacking these genes into one transgenic plant. Incorporation of an avidin gene into the cotton genome alone or stacked with Bt toxin genes could be advantageous for Bt resistance management. Furthermore, bollworms could


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perhaps be completely controlled by avidin alone if cotton could express the toxin at levels comparable to corn (3000 ppm) as shown by Kramer (2004). In addition, avidin could also help delay development of resistance to Bt. Du & Nickerson (1996) demonstrated that an avidin analog, streptavidin, could bind to 85 and 120 kDa proteins of BBMV in Manduca sexta. The binding was influenced by biotin and Bt toxin, suggesting that biotin containing proteins might be common receptors for both avidin and Bt. It has not been confirmed whether synergism or interference occur in binding between Bt toxin and avidin. Based on Zhu et al. (2005), it was likely that Bt synergizes avidin binding, or vice versa, because the combination of Bt and avidin consistently showed synergism in larval body weight and length suppression and mortality increase in the target insects. Considering different modes of action and synergism, avidin is a very potent bio-pesticide. Stacking the avidin gene over transgenic Bt cotton might be very useful for increasing control range and delaying resistance evolution. Because of the potential threat of resistance development, it is also very urgent to seek alternative insecticidal proteins to be inserted into the cotton genome for controlling not only lepidopteran pests, but also sucking insects. Due to the widespread use of transgenic Bt cotton, the number of chemical insecticide applications has been substantially reduced, which has resulted in the resurgence of sucking insects as a serious problem in cotton (Snodgrass 1996; Snodgrass and Scott 2000). Therefore, transgenic cotton with genes that target a wider range of pests is urgently needed. In addition to its efficacy against lepidopteran insects (Burgess et al. 2002; Marwick et al. 2001, 2003), avidin is highly insecticidal against many coleopteran pests (Morgan et al. 1993; Allsopp and McGhie 1996; Kramer et al. 2000). If avidin is shown to also be effective against sucking insects, its introduction into transgenic cotton, along with its synergistic action with Bt toxins, could be a major advance in cotton insect control and Bt resistance management. Another alternative for increasing control range and for delaying Bt resistance development is proteinase inhibitors. Serine proteinases are common luminal enzymes in midguts of many Lepidoptera species (Applebaum 1985; Terra and Ferriera 1994). Serine proteinases in the midgut also activate protoxins, thereby mediating Bt toxicity (Milne, R., H. Kaplan 1993; Martínez-Ramírez, A.C., M.D. Real 1996), and they also may play a concurrent role in the hydrolytic degradation and subsequent inactivation of the toxic protein. It is also likely that development of Bt resistance is associated with gut proteinase profile and activity changes. Forcada et al. (1996) provided evidence that midgut extracts from a Btresistant strain of H. virescens degraded the Bt toxin in vitro at a rate faster than extracts from a susceptible strain. Shao et al. (1998) further proved that the excessive degradation of protoxin in H. armigera midgut juice, which contained trypsin-, chymotrypsin-, and elastase-like enzymes, was responsible for the low sensitivity of this species to Bt, with chymotrypsin playing a major role in the degradation process. Cotton varieties transformed with genes for Bt toxins are designed to disrupt

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gut epithelial cell physiology of the target insects. However, the potential for gut proteinases to modulate the effectiveness of Bt protoxins in insects is high and was the impetus for more detailed studies on the interaction of Bt toxins and gut proteinase. Zhu et al. (2007) demonstrated the interaction of Bt and proteinase inhibitors. Insect growth was consistently retarded more by Bt + inhibitor treatments than by treatments of Bt alone or inhibitor alone. A synergistic effect was observed in most of the bioassay experiments. Stunted growth was further reflected by suppressed gut proteinase activities in insects treated with Bt + inhibitor. The mechanisms of the Bt and proteinase inhibitor interaction have not been studied previously. By analyzing growth and gut proteinase activities, we suggest that Bt alone at a sublethal dose may damage gut tissues and reduce production of the proteinases (Fig. 19.2). The damage could extend to a change in gut permeability and absorption of nutrients. These limited proteinases still maintain proteolytic activity for providing basic nutrients, and for processing Bt protoxins to toxins and degradation of toxins as well. Limited reduction of growth and proteinase activity are attributed to the damage of gut cells incurred by Bt binding (Fig. 19.2). Most proteinase inhibitors function competitively with the substrate to share the active site of the enzyme to cause nutritional deficiency. Retarded growth is a common symptom of proteinase inhibition in insects. By feeding on a diet containing a relatively low dose of the proteinase inhibitor, insects were able to

Fig. 19.2 Schematic model of Bt protoxin activation and toxin degradation by midgut proteinases without addition of proteinase inhibitors. The model suggests that Bt toxin degradation is fast in resistant insects which results in less Bt toxin binding to midgut brush board membrane vesicle.


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live without substantial reduction of body weight and length. The insects still had enough free proteinases to hydrolyze dietary proteins. Proteinase activity data proved that there was no substantial suppression of gut enzyme activity after inhibitor only treatment. By applying a combination of Bt and individual proteinase inhibitor, significant reduction of larval body weight and length was observed. The interaction of Bt and proteinase inhibitor for most measurements was greater than the additive effect of individual Bt and proteinase inhibitor treatment. Interaction was further reflected by proteinase activity data. Treatment of Bt + Inhibitor consistently and significantly suppressed the enzyme activity in the gut. The interaction was mostly synergistic, which is consistent with the results of the bioassay. Bt-inhibitor interaction is most likely achieved by suppression of Bt degradation when a proteinase inhibitor is present (Fig. 19.3). Although low dose inhibitor treatments did not critically suppress the hydrolyzation of dietary proteins for nutritional requirements, it might significantly suppress gut proteinases, and subsequently suppress Bt degradation by gut enzymes. More Bt toxin stays actively bound to targets and creates more damage to the gut membrane. Extended gut damage results in further suppression of gut proteinase production and a subsequent decrease in proteinase activity (Fig. 19.3).

Fig. 19.3 Schematic model of Bt protoxin activation and toxin degradation by midgut proteinases with addition of proteinase inhibitors. The model suggests that Bt toxin degradation is substantially suppressed which results in more gut damage and less proteinase production due to more Bt toxin binding to midgut brush board membrane vesicle. Acknowledgements We thank Dr. Susan Li (USDA-ARS, Stoneville, MS USA) and Dr. Lingxiao Zhang (Mississippi State University DREC, Stoneville, MS USA) for reviewing and improving an earlier draft of this manuscript. Authors appreciate Sandy West for the laboratory assistance and efforts for improving this manuscript, and Dr. Carlos Blanco for providing part of reagents. Mention of a trademark, warranty,

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proprietary product or vendor does not constitute a recommendation or endorsement by the USDA and does not imply approval or recommendation of the product to the exclusion of others that may be suitable.

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Section 4: Emerging Pest Management Strategies and Technologies

sdfsdf

CHAPTER 20

Advances and Prospects of RNAi Technologies in Insect Pest Management Xin Zhang, Jianzhen Zhang and Kunyan Zhu

Abstract RNA interference (RNAi) is a common mechanism of posttranscriptional gene silencing that destroys mRNA of a particular gene to prevent translation to form an active gene product (most commonly a protein). The discovery of RNAi has not only provided a breakthrough in methodology for functional analysis of genes, but also opened a novel avenue for treating human diseases and protecting crops against insect pest damages. Space injection of double-stranded RNA (dsRNA) is the most commonly used method for RNAi in entomological research. Recent studies, however, have demonstrated that specific suppression of gene expression in insects can also be accomplished by feeding and topical application of dsRNA in certain insect species. The specific gene silencing using RNAi with feeding and topical application methods holds great promises of application of RNAi for controlling both agriculturally and medically important insects. Indeed, transgenic plants expressing dsRNA of specific genes have already been demonstrated for plant resistance against insect pests. This paper reviews and discusses the current advances and prospects of RNAi technologies in insect pest management. Keywords environmental RNAi, host-delivered RNAi, insect pest management, RNA interference, systemic RNAi

20.1 Introduction: Mechanism of RNAi and its applications in insect science RNA interference (RNAi) is a phenomenon of downregulation of gene expression by double-stranded RNA (dsRNA) or small interfering RNA Xin Zhang, Jianzhen Zhang, Kunyan Zhu Department of Entomology, Kansas State University, Manhattan, KS, 66506, USA E-mail: [email protected] Jianzhen Zhang College of Life Science and Technology, Shanxi University, Taiyuan, Shanxi, 030006, China


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Xin Zhang, Jianzhen Zhang and Kunyan Zhu

(siRNA). RNAi is a post-transcriptional control mechanism involving degradation of a target mRNA mediated through the production of siRNAs from the dsRNA, which is cleaved by dsRNA-specific endonucleases known as dicers. The siRNAs are 21 bp dsRNA fragments carrying two base extensions at the 3′ end of each strand; one strand of the siRNA is assembled into an RNA-induced silencing complex (RISC) in conjunction with the argonaute multi-domain protein, which contains an RNaseH-like domain responsible for target degradation (Price and Gatehouse 2008). RNAi was first elegantly described in the nematode, Caenorhabditiselegans (Fire et al. 1998), and now it has been found to be a conserved mechanism in eukaryotes including fungi, plants, insects and mammals (Mello and Conte 2004). As a breakthrough technique, RNAi has broadened our understanding of gene regulation and has revolutionized methods for genetic analysis, which has been widely used in model organisms such as C. elegans and Drosophila melanogaster. RNAi has been widely used in genetic research in insects of different orders at least including Diptera, Lepidoptera, Coleoptera, Hymenoptera, Orthoptera, Hemioptera, Boattodea, and Neoptera. Delivery of dsRNA to initiate RNAi in insects has predominately been via microinjection of nanogram amounts of long dsRNA into insect body cavity. In C. elegans, RNAi can be produced by feeding bacteria expressing dsRNA (Timmons et al. 2001), or even by soaking nematodes in dsRNA solution (Tabara et al. 1998). This triggers RNAi in response to dsRNA molecules encountered in their environment and is referred to as environmental RNAi (Whangbo and Hunter 2008). Recently, feeding RNAi has also been reported in the light brown apple moth, Epiphyas postvittana (Turner et al. 2006), bug Rhodnius prolixus (Rajagopal et al. 2002), and termite, Reticulitermes flavipes (Zhou et al. 2008). Most recently, plantmediated RNAi has been demonstrated as a viable approach for control of insect pests in agricultural settings (Baum et al. 2007; Mao et al. 2007). These results show the great potential of applying RNAi in insect pest management in agriculture. Here, we review the current understanding of RNAi and the advances of RNAi-mediated insect pest management and discuss its future directions in this emerging field.

20.2 Genetic basis of systemic RNAi and environmental RNAi in C. elegans The phenomenon in which local administration of dsRNA (e.g., the gut through feeding) leads to an RNAi response in the whole body through the amplification and spread of silencing to other cells and progeny is known as systemic RNAi. Systemic RNAi has been well studied in plants and nematodes. The basis of systemic RNAi lies on the presence of an RNA-dependent RNA polymerase (RdRP) that is able to interact with the RISC complex and generate new dsRNA based on the partially degraded target template by using the hybridized siRNA strands as primers (Sijen et al. 2001). In plants, the targeted mRNA is converted

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to long double-stranded RNA by the RdRP and new siRNAs (secondary siRNAs) are generated after cleavage by dicer (Baulcombe 2007). In C. elegans, however, an argonaute protein associated with a primary siRNA in the RISC targets a longstranded RNA and recruits an RdRP that synthesizes 22–23 nucleotide secondary siRNAs directly. Thus, once dsRNAs are introduced into a cell, the RNAi effect can be amplified through the generation of secondary siRNAs. The secondary siRNAs can also be exported to neighboring cells to spread the gene silencing, leading to a persistent RNAi effect through the organism and over its developmental time. Another property of RNAi in C. elegans is that dsRNA in environment can trigger strong responses in the whole organism. Studies on feeding RNAi in C. elegans have provided information on how dsRNA molecules enter an organism from environment to trigger RNAi. dsRNA is uptaken through the intestinal lumen when fed by C. elegans. The ingested dsRNA and resulting silencing signals spread systemically to distant cell within the animal. For spreading of silencing signals, the target gene is not required to be expressed in the intestine (Winston et al. 2007). In order to study the uptake of dsRNA, genetic analysis in C. elegans has been carried out and two systemic RNA interference deficient genes, sid-1 and sid-2, have been identified (Winston et al. 2002). sid-1 encodes a conserved multiple transmembrane channel protein that is expressed on the cell surface with homologs in most animals (Winston et al. 2002). sid-1 mutant worms remain competent in cell autonomous RNAi, but cannot perform systemic RNAi in response to feeding, soaking, injection of dsRNA or in vivo expression of dsRNA from transgenes. When expressed in Drosophila S2 cells, sid-1 enhanced the ability of S2 cells to uptake dsRNA at sub-optimal dsRNA concentrations by an apparently passive mechanism (Feinberg and Hunter 2003). sid-2 encodes a gut-specific transmembrane protein (SID-2) with a single transmembrane region with homologs identified only in C. briggsae and C. remanei. SID-2 is necessary for the initial import of dsRNA into the animal from the gut lumen but is not required for the systemic spread of silencing signals between cells and tissues (Winston et al. 2007). Localization in the apical membranes of the intestinal cell also suggests that SID-2 facilitates the import of ingested dsRNA from the intestinal lumen. However, SID-2 alone is not sufficient for dsRNA import from the intestinal lumen because sid-1 mutants are also unable to uptake environmental dsRNA. Interestingly, the nematode C. briggsae has its own sid-2, but is defective in uptake of dsRNA from the gut lumen. Once transformed with C. elegans sid-2, C. briggsae gains a systemic RNAi response (Winston et al. 2007). Investigation of the RNAi in the other eight available Caenorhabditis species showed that only one of these tested species was proficient in environmental RNAi, indicating that the ability to perform environmental RNAi might be uncommon, or the process might be regulated by factors that are not present in standard laboratory growth conditions (Whangbo and Hunter 2008). Besides sid-1 and sid-2, other genes that are involved in environmental RNAi

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have been discovered in C. elegans, which include the feeding defective (fed) mutants (fed-1 and fed-2) (Timmons et al. 2003) and the RNA spreading defective (rsd) mutants (rsd-2,-3, -4, -6 and -8). Mutants of rsd-4 and rsd-8 are resistant to RNAi by feeding both in germline and somatic cells. While mutants of rsd-2, rsd3, and rsd-6 are resistant to feeding RNAi in germline, but are still sensitive to feeding RNAi in somatic cells (Tijsterman et al. 2004). Recent studies performed in C. elegans and D. melanogaster using a reverse genetic screen suggest that receptor-mediated endocytosis is involved in dsRNA uptake from environment (Saleh et al. 2006; Ulvila et al. 2006). Twenty-three genes involved in endosytic pathway are identified are required for uptake of dsRNA by cultured Drosophila S2 cells from medium. Knockdown of 10 of the 23 homolog genes in C. elegans disrupted the ability to trigger an RNAi response by feeding the worms with dsRNA (see Table 20.1; and review by Whangbo and Hunter 2008). These results suggest that endocytosis might be a common mechanism for dsRNA uptake and might occur in different insect orders. If so, it might be very promising that RNAi can be done in insect pests from different orders which is effectively targeted by oral delivery of dsRNA (Price and Gatehouse 2008).

20.3

Systemic and environmental RNAi in insects

Very limited information on the genetic basis of systemic and environmental RNAi is available in insects compared with that from C. elegans. To date, it Table 20.1

Genes involved in environmental RNAi in C. elegans

Gene name

Gene product

RdRP

RNA-dependent Enzyme for siRNA amplification leading RNA polymerase. to persistent and systemic RNAi

Sijen et al. 2001

sid-1

Multiple transmembrane channel protein (likely dsRNA channel)

Mutant being defective in systemic and environmental RNAi

Winston et al. 2002

Gut-specific transmembrane protein (likely dsRNA receptor)

Mutant being defective in feeding and soaking RNAi, but sensitive to injection RNAi

Winston et al. 2007

Feeding RNAi defective protein

Mutant being defective in feeding RNAi, Timmons et al. 2003 but sensitive to injection RNAi

rsd-4, rsd-8, rsd-2, RNA spreading rsd-3, rsd-6 defective protein

Mutant being defective in feeding RNAi, Tijsterman et al. 2004 but sensitive to injection RNAi

arl-1, F22G12.5, cgoc-2, ZK1098.5, Feeding RNAi vps-41, vps-34, defective protein ger-1, bre-3, sedl-1, sym-3

Knockdown by RNAi leading to defective Saleh et al. 2006 feeding RNAi

sid-2

fed-1, fed-2

Function in environmental RNAi

Reference


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appears that insects lack RdRP needed to drive this RNAi amplification in C. elegans. Homologues of the C. elegans sid-1 gene have been identified in several insect species such as Tribolium castaneum, Bombyx mori, Apis mellifera and aphids, but not in dipterans such as Drosophila and mosquitoes (Gordon and Waterhouse 2007; Tomoyasu et al. 2008). Systemic RNAi has been reported in several insect species although its genetic mechanism is still elusive as described above. Insect systemic RNAi was first documented in the coleopteran insect, the red flour beetle, T. castaneum. Tomoyasu and Denell (2004) reported that injection of Tc-ASH (a homologue of the Drosophila sensory bristle-forming gene) dsRNA into larvae at a single discrete site resulted in a ‘loss-of-bristle’ phenotype over the entire epidermis of adult insects. In another study, injection of adult females with dsRNA specific to three genes-Distalless (leg development gene), maxillopedia (homeotic gene), and proboscipedia (encoding a homeotic protein required for the formation of labial and maxillary palps)- led to RNAi effect in both mother insects and developing progeny embryos after egg hatch (Bucher et al. 2002). With the robust systemic RNAi response and the completed genome sequence, Tribolium is becoming a model organism for genetic analysis by using RNAi. Intriguingly, a recent genome comparison of C. elegans and Tribolium revealed a lack of conservation of a systemic RNAi mechanism (Tomoyasu et al. 2008). For example, Tribolium lacks a C. elegans-like RdRP, so the signal amplification observed in Tribolium must be based on a different gene with a similar activity, or possibly even a different mechanism. Three Tribolium homologs of C. eleganssid-1 were identified. However, these Tribolium sid-like genes do not seem to be required for systemic RNAi. Only ortholog of C. elegans rsd-3 has been identified in Tribolium. Considering Drosophila also has the rsd-3 ortholog, the possibility is low for this gene to confer systemic RNAi in Tribolium. In addition to Tribolium, grasshopper also demonstrated systemic RNAi response in the eyes by injection of the dsRNA of a gene in the abdomen of the nymphs (Dong and Friedrich 2005). Effective feeding for RNAi is a prerequisite to use of RNAi for crop protection against insect pests. Feeding RNAi was documented in larvae of the light brown apple moth (Epiphyas postvittana) by incorporation of dsRNA of gut-specific carboxyesterase genes through droplet feeding. Systemic RNAi response with persistence RNAi signal was also shown as the that knockdown of a gene expressed in the adult antenna could be achieved through feeding dsRNA to larvae (Turner et al. 2006). Oral delivery of dsRNA of two cytochrome P450 genes, CYP6BG1 and CYP6BF1v4, by droplet feeding has been demonstrated to reduce the gene expression efficiently in the larvae of Plutella Xylostella. Repression of CYP6BG1 transcript was evident not only in the midgut but also in the larval tissues enclosed in the carcass (Bautista et al. 2009). However, feeding RNAi against a midgut aminopeptidase-N gene was not successful in another lepidopteran Spodoptera litura, though downregulation of this gene by microinjection of dsRNA into the insect haemoceol was very efficient (Rajagopal

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et al. 2002). Feeding dsRNA of a salivary gland gene encoding nitroporin 2 (NP2) led to downregulation of the gene expression in the bug Rhodnius prolixus (Hemioptera) (Araujo et al. 2006). A recent study in the termite Retculitermes flavipes demonstrated that silencing of two genes led to significant reduction of group fitness and mortality after feeding of the dsRNA. One of the genes is a proximally expressed in salivary glands encoding an endogenous digestive cellulose enzyme (Cell-1) and the other is distally expressed in fatty body encoding a caste-regulatory hexamerin storage protein (Hex-2) (Zhou et al. 2008). These results indicate that the target genes for feeding RNAi are not necessarily restricted to the gut-specific genes. The gene knockdown by feeding was also reported in a blood-sucking Diptera insect, Glossina morsitans morsitans, by incorporating dsRNA in the bloodmeal. At transcript and protein level, delivering dsRNA in the bloodmeal is as effective as injection in knockdown of an immunoresponsive midgut-expressed gene Tsetse EP. However, feeding dsRNA fails to knockdown the fatty body-expressed transferrin gene, 2A192, previously shown to be silenced by dsRNA injection (Walshe et al. 2009). These studies demonstrated that dsRNAs can be successfully delivered and trigger silencing of target genes in different insect orders. Insect control through RNAi is dependent on the feeding delivery of the dsRNA. The studies on feeding RNAi are increasing very fast these days and the list of the successful examples of feeding RNAi is getting larger (Table 20.2). Most recently and for the first time, Pridgeon et al. (2008) reported that topical application dsRNA of an inhibitor of apoptosis protein I gene (AaeIAP1) could lead to a down-regulation of the gene and was able to kill female adults of Aedes aegypti. The highest mortality caused by dsRNA-IAP1-D could reach to 48% after 24 h of topical application. These results show great potential in the development of molecular pesticides for the control of medically important disease vectors by targeting the critical pathways or genes.

20.4

Plant-delivered RNAi and insect pest management

Silencing of endogenous genes in insects by ingesting dsRNA suggests the possibility of applying RNAi technology for insect pest control. This strategy, known as host-delivered RNAi or in planta RNAi, was first reported for the control of root knot nematode, Meloidogyne incognita, and several studies show that host-delivered-RNAi could trigger sequence-specific gene silencing and could confer effective resistance to the plant-parasitic nematodes (reviewed by Fuller et al. 2008). Similarly, two recent studies have shown that transgenic plants expressing dsRNA of specific insect genes can enhance resistance to these herbivorous insects. In one study, a total of 290 essential genes were screened by feeding the larvae with artificial diet supplemented with specific dsRNA in western corn rootworm (Diabrotica virgifera virgifera). A total of 14 genes were identified to lead to specific downregulation of their expression at low dsRNA concentrations

Advances and Prospects of RNAi Technologies in Insect Pest Management Table 20.2

Delivery methods that have been tested for potential insect control

Delivery method

Insect species

Epiphyas postvittana

Feeding of dsRNA with food

Topical application of dsRNA

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Gene name

Gene product

EposCXE1; EposPBP1;

A larval gut carboxylesterase and a pheromone binding protein, respectively.

Reference

Turner et al. 2006

Plutella Xylostella CYP6BG1, and CYP6BF1v4

Cytochrome P450 monooxygenases expressed in the midgut and other tissues.

Rhodnius prolixus NP2

A salivary gland nitroporin 2.

Retculitermes flavipes

Endogenous digestive cellulose in salivary glands, and a Zhou et al. 2008 caste-regulatory hexamerin storage protein in fatty body, respectively.

Cell-1; and Hex-2

Bautista et al. 2009

Araujo et al. 2006

Glossina morsitans TsetseEP morsitans

Immunity-related protein highly expressed in midgut.

Apis mellifera

AmVg

Vitellogenin

Nunes and Simões 2009

Aedes aegypti

AaeIAP1

An inhibitor of apoptosis protein I.

Pridgeon et al. 2008

Diabrotica V-ATPase A virgifera virgifera Transgenic plants expressing small Helicoverpa armigera hairpin RNA

CYP6AE14 and GST1

A midgut vacuolar ATPase, subunit A. A cytochrome P450 monooxygenase and a glutathione S-transferase expressed in midgut.

Walshe et al. 2009

Baum et al. 2007

Mao et al. 2007

and result in insect stunting and mortality. Transgenic corn expressing one of the most effective dsRNAs, directed against a gene encoding V-type ATPase A, showed a significant level of protection against rootworm infestation. When dsRNA designed to target rootworm genes were tested in larvae from two other insects pests, mortality declined with decreasing sequence identity between the rootworm genes and their orthologs in the other species, indicating that gene silencing was poetically very selective (Baum et al. 2007). In the second study, the product of a cytochrome P450 gene, CYP6AE14, which is highly expressed in the midgut of cotton bollworm (Helicoverpa armigera) has been identified as the antidote against one of the cotton’s natural

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secondary metabolite (the toxic compound gossypol). The larvae of cotton bollworm fed on transgenic tobacco (Nicotiana tobacum) and Arabidopsis thaliana plants expressing dsRNA of this gene showed downregulation of the endogenous CYP6AE14 and significant larval growth retardation when transferred to artificial diets (Mao et al. 2007). An expected followup study is to examine whether transgenic cotton plant expressing CYP6AE14 dsRNA can confer similar resistance to H. armigera. The above two studies share some common factors for success: 1) identification of suitable insect target genes (V-ATPase and CYP6AE14), 2) target genes mainly expressed in the midgut, 3) in planta expression of dsRNA, and 4) a continuous supply of dsRNA to the gut cells of the feeding insects (Price and Gatehouse 2008). An important issue from both studies is whether the systemic RNAi responses occur within the insect. Insect genomics indicates that cotton bollworm and western corn rootworm lack RdRP, which is same as that in Tribolium. Perhaps unamplified RNAi is effective against the P450 and V-ATPase genes because they are mainly expressed in the midgut and the gut cells of the feeding insect receive a continuous supply of hairpin RNA. The hairpin RNA would be diced into siRNAs in these cells and may also spread into surrounding cells and tissues via the intercellular dsRNA transport SID proteins, whose genes are present in almost all animals, including arthropods. It would be interesting to discover whether phenotypic silencing can be observed for genes not expressed in the midgut (Gordon and Waterhouse 2007).

20.5

Conclusion and future directions

The specific gene silencing using RNAi with feeding and topical application methods have shown great potentials to use RNAi for controlling both agriculturally and medically important insects. Most importantly, the transgenic plants expressing dsRNA of specific genes have shown significant plant resistance against insect pests. Theses results have demonstrated the feasibility of using RNAi in crop protection against herbivorous insects. This approach is promising for the future pest management because it allows a wide range of potential targets for suppression of gene expression in insects to be exploited. The minimum requirement of using RNAi in insect pest control involves following steps: 1) enough supply of dsRNAs and its effective delivery, 2) uptake of dsRNA from environment either through intestinal cells or cuticle, and 3) target gene silencing via cell autonomous RNAi machinery. Definitely, systemic RNAi responses including amplification of signals and spreading to distant cells and tissues will increase the efficiency significantly (Fig. 20.1), which has been demonstrated in insects such as Tribolium (Tomoyasu et al. 2008), light brown apple moth (Turner et al. 2006), and grasshopper (Dong and Friedrich 2005), though RdRP has not been identified in any insects. Expression of dsRNA in transgenic plants provides a continuous supply of dsRNA for the feeding insects. Thus, it is enough to lead to RNAi responses in the midgut of insects and finally


Fig. 20.1 Diagram of systemic RNA interference triggered by double-stranded RNA (dsRNA) in lower animals. Ingested exogenous dsRNA in intestinal lumen is imported into intestinal cells through transport proteins SID-1 and SID-2, which have been identified in Caenorhabditis elegans, or possibly through endocytosis. Relatively long dsRNA is first processed by a dicer (a member of the RNase III nuclease family) in an ATP-dependent reaction into primary small interfering RNA (siRNA) which is 21~23nucleotide dsRNA with a 2-nucleotide 3’ overhang. The siRNA is then incorporated into the RNA-induced silencing complex (RISC) which consists of an argonaute protein. Argonaute cleaves and discards the passenger (sense) strand of the siRNA duplex leading to activation of RISC. The remaining guide (antisense) strand of the siRNA then guides RISC to its homologous target mRNA through base-pairing and induces cleavage of the target mRNA. The systemic RNAi in C. elegans and some of other organisms (e. g., ticks) also relies on the presence of RNA-dependent RNA polymerase (RdRP). RdRP uses the antisense siRNA as a primer and the target mRNA as a template to synthesize a “secondary” population of siRNA. Such a new population of the siRNA can either be used to initiate a RNAi effect through the RISC-dependent pathway as described above or be exported from the cell (may be through SID-1) to spread the RNAi effect to other cells.

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results in plant resistance against the feeding insects even without systemic RNAi responses in the targeting insects. RNAi response in adult mosquitoes has been documented with a topical application method. Thus, it is also promising to apply dsRNAs as molecular pesticides directly by large amount of dsRNA production using the bacterial expression system for which the technique has already been successfully established. In C. elegans, it is obvious that the midgut cells use a different mechanism to uptake dsRNA from the gut lumen compared with that from the hemolymph. SID-2 is exclusively involved in uptake of dsRNA from gut lumen instead of the hemolymph. The mechanism is still not clear for insects to uptake dsRNA. Strong systemic RNAi response has been well documented in Tribolium. However, SID1-like proteins are not involved in systemic RNAi in Tribolium. It is still not known if SID-1-like proteins are the mechanism for dsRNA uptake from the body cavity. Similarly, nothing is known on the mechanism of uptake of dsRNA from the gut lumen. It is still possible that different mechanisms are involved in uptake of dsRNA from hemolymph and from gut lumen. To date, RNAi by injection of dsRNA has been well documented in insects, whereas feeding RNAi is only reported in very limited insect species. It is still not clear whether it is common to perform RNAi by feeding methods in insects. Thus, it is critical to reveal the exact mechanism of dsRNA uptake from gut lumen for future application of RNAi in insect pest management. Target gene selection is crucial for the success of RNAi-mediated insect pest control. A series of potential target genes were identified by Baum et al. (2007), and most of which involved in essential physiological process in insects. Thus, these genes such as V-type ATPase A might easily be extended to other insect species. The approach of Mao et al. (2007), in which insect detoxification mechanisms towards plant allelochemicals are targeted, has the advantage of being predictable and specific to pests that feed on a crop producing a defined defensive chemical. Insects have multiple enzymes (P450s, glutathione Stransferases and carboxylesterases) for detoxification of plant chemicals. Genes encoding these enzymes are attractive targets for applying dsRNA to enhance the susceptibility of insects against plant allelochemicals. Mao et al. (2007) have demonstrated that silencing of a glutathione S-transferase gene in bollworm. These findings strongly suggest that plant-delivered silencing of insect detoxification genes could be a powerful strategy for controlling insect pests (Gordon and Waterhouse 2007). The challenges are still there. Fox example, whether all detoxification genes are equally amenable to this approach? Will this technology extend to other insect groups? Companying the exciting potential of a widespread approach for the management of insect pests, many important issues should be addressed before this technology can be applied on a large scale. Firstly, the specificity or the offtarget effects must be well studied in either the crops, crop pests or other consumers of the transgenic crops (i.e. livestock and people). Effects on nontarget insects should be minimized. Secondly, the risk of resistance development


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by the pest populations should be considered. Could pest populations circumvent RNAi through acquisition of point mutations or by sequence polymorphisms in the target gene? Despite of these questions, there is no doubt that researchers and farmers have grounds to look forward to a new era in insect pest control. Acknowledgements The authors thank Ming-Shun Chen and Xuming Liu for their review of an earlier draft of this manuscript. The relevant research was supported by Arthropod Genomics Center and Ecological Genomics Institute, both funded by K-State Targeted Excellence Program, and Kansas Agricultural Experiment Station. This manuscript is contribution No. 09-333-B from the Kansas Agricultural Experiment Station, Kansas State University, Manhattan, Kansas, USA.

References Araujo R N, Santos A, Pinto F S, et al. RNA interference of the salivary gland nitrophorin 2 in the triatomine bug Rhodnius prolixus (Hemiptera Reduviidae) by dsRNA ingestion or injection. Insect Biochem. Mol. Biol., 2006, 36: 683–693. Baulcombe D C. Amplified silencing. Science, 2007, 315: 199–200. Baum J A, Bogaert T, Clinton W, et al. Control of coleopteran insect pests through RNA interference. Nat. Biotechnol., 2007, 25: 1322–1326. Bautista M A M, Miyata T, Miura K, Tanaka T. RNA interference-mediated knockdown of a cytochrome P450, CYP6BG1, from the diamondback moth, Plutella xylostella, reduces larval resistance to permethrin. Insect Biochem. Mol. Biol., 2009, 39: 38–46. Bucher G, Scholten J, Klingler M. Parental RNAi in Tribolium (Coleoptera). Curr. Biol., 2002, 12: R85–R86. Dong Y, Friedrich M. Nymphal RNAi: Systemic RNAi mediated gene knockdown in juvenile grasshopper. BMC Biotechnol., 2005, 5: 25. Feinberg E H, Hunter C P. Transport of dsRNA into cells by the transmembrane protein SID-1. Science, 2003, 301: 1545–1547. Fire A, Xu S Q, Montgomery M K, et al. Potent and specific genetic interference by doublestranded RNA in Caenorhabtidis elegans. Nature, 1998, 39: 806–811. Fuller V L, Lilley C J, Urwin P E. Nematode resistance. New Phytol., 2008, 180: 27–44. Gordon K H, Waterhouse P M. RNAi for insect-proof plants. Nat. Biotechnol., 2007, 25: 1231– 1232. Mao Y B, Cai W J, Wang J W, et al. Silencing a cotton bollworm P450 monooxygenase gene by plant-mediated RNAi impairs larval tolerance of gossypol. Nat. Biotechnol., 2007, 25: 1307– 1313. Mello C C, Conte D. Revealing the world of RNA interference. Nature, 2004, 431: 338–342. Nunes F M F, Simõs L P Z. A non-invasive method for silencing gene transcription in honeybees maintained under natural conditions. Insect Biochem. Mol. Biol., 2009, 39: 157–160. Price D R, Gatehouse J A. RNAi-mediated crop protection against insects. Trends Biotechnol., 2008, 26: 393–400. Pridgeon J W, Zhao L, Becnel J J, et al. Topically applied AaeIAP1 double-stranded RNA kills female adults of Aedes aegypti. J. Med. Entomol., 2008, 45: 414– 420. Rajagopal R, Sivakumar S, Agrawal N, et al. Silencing of midgut aminopeptidase-N of Spodoptera

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litura by double-stranded RNA establishes its role as Bacillus thuringiensis toxin receptor. J. Biol. Chem., 2002, 277: 46849–46851. Saleh M C, van Rij R P, Hekele A, et al. The endocytic pathway mediates cell entry of dsRNA to induce RNAi silencing. Nat. Cell Biol., 2006, 8: 793–802. Sijen T, Fleenor J, Simmer F, et al. On the role of RNA amplification in dsRNA triggered gene silencing. Cell, 2001, 107: 465–476. Tabara H, Grishok A, Mello C C. RNAi in C. elegans: Soaking in the genome sequence. Science, 1998, 282: 430–431. Tijsterman M, May R C, Simmer F, et al. Genes required for systemic RNA interference in Caenorhabditis elegans. Curr. Biol., 2004, 14: 111–116. Timmons L, Court D L, Fire A. Ingestion of bacterially expressed dsRNAs can produce specific and potent genetic interference in Caenorhabditis elegans. Gene, 2001, 263: 103–112. Timmons L, Tabara H, Mello C C, Fire A. Inducible systemic RNA silencing in Caenorhabditis elegans. Mol. Biol. Cell, 2003, 14: 2972–2983. Tomoyasu Y, Denell R E. Larval RNAi in Tribolium (Coleoptera) for analyzing adult development. Dev. Genes. Evol., 2004, 214: 575–578. Tomoyasu Y, Miller S C, Schoppmeier M, et al. Exploring systemic RNA interference in insects: A genome-wide survey for RNAi genes in Tribolium. Genome Biol., 2008, 9: R10. Turner C T, Davy M W, MacDiamid R M, et al. RNA interference in the light brown apple moth, Epiphyas postvittana (walker) induced by doublestranded RNA feeding. Insect Mol. Biol., 2006, 15: 383–391. Ulvila J, Parikka M, Kleino A, et al. Double-stranded RNA is internalized by scavenger receptormediated endocytosis in Drosophila S2 cells. J. Biol. Chem., 2006, 281: 14370–14375. Walshe D P, Lehane S M, Lehane M J, Haines L R. Prolonged gene knockdown in the tsetse fly Glossina by feeding double stranded RNA. Insect Mol. Biol., 2009, 18: 11–19. Whangbo J S, Hunter C P. Environmental RNA interference. Trends Genet., 2008, 24: 297–305. Winston W M, Molodowitch C, Hunter C P. Systemic RNAi in C. elegans requires the putative transmembrane protein SID-1. Science, 2002, 295: 2456–2459. Winston W M, Sutherlin M, Wright A J, et al. Caenorhabditis elegans SID-2 is required for environmental RNA interference. Proc. Natl. Acad. Sci. USA, 2007, 104: 10565–10570. Zhou X, Wheeler M M, Oi F M, et al. RNA interference in the termite Reticulitermes flavipes through ingestion of double-stranded RNA. Insect Biochem. and Mol. Biol., 2008, 38: 805–815.

CHAPTER 21

Anti-tick Vaccine Development: Status and Perspectives Quentin Q. Fang and Oscar J. Pung

Abstract Ticks (Acarina: Arachnida: Arthropoda) are obligate, nonpermanent ectoparasites of terrestrial vertebrates. In arthropods, ticks are second only to mosquitoes in the number of pathogens they transmit to people, domestic pets, livestock, and wild animals. Ticks are vectors to a variety of human diseases, including Rocky Mountain spotted fever, Lyme disease, Colorado tick fever, tickborne encephalitis, babesiosis, tularemia, and tick-borne relapsing fever. In addition, other maladies due to tick bites, such as tick paralysis, tick toxicosis, and anaphylaxis, are common. Each year, worldwide public health and agricultural industry costs due to tick-transmitted diseases are estimated to be in the range of several billion U.S. dollars. To date, the control of ticks has relied heavily on chemical pesticides (acaricides). The excessive use of pesticides in tick control has resulted in environmental contamination, food safety concerns, and pesticide resistance. Consequently, use molecular technology to develope anti-tick vaccines is an essential alternative strategy in tick and tick-borne disease control. Recent studies and the status of anti-tick vaccine development are reviewed in this chapter. Keywords external parasites, anti-tick vaccine, vectors, tick-borne disease

Ticks are small arachnids in the superfamily Ixodoidea that, along with other mites, constitute the subclass Acarina in the phylum Arthropoda (Oliver 1989; Sonenshine 1991). Ticks are classified into three families: The family Argasidae (soft ticks, ~183 species), the family Ixodidae (hard ticks, ~683 species), and the family Nuttalliellidae (containing the monospecific species Nuttalliella namaqua) (Horak et al. 2002). All ticks are obligate, nonpermanent ectoparasites of terrestrial vertebrates. Hard ticks have three distinct life stages: larva (six legs), Quentin Q. Fang, Oscar J. Pung Department of Biology, Georgia Southern University, Statesboro, GA, 30460, USA E-mail: [email protected]


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Quentin Q. Fang and Oscar J. Pung

nymph (eight legs), and adult (eight legs). All three stages must feed on blood for their development, but only one blood meal is taken during each of the three life stages. Adult female hard ticks lay one batch of thousands of eggs and then die. The entire life cycle of a hard tick can take less than a year in tropical regions and over three years in cold climates, where certain stages may enter diapause until hosts are again available. The hard ticks are slow feeders, usually requiring at least 24 hours to two weeks for full engorgement (Oliver 1989; Matuschka et al. 1990; Sonenshine 1991; Falco et al. 1996), and many can go for several months without feeding if not unduly stressed by environmental conditions (Sonenshine 1991, 1993). The life cycle of soft ticks also consists of larval, nymphal and adult stages but these are not readily distinguishable from each other. For example, many soft ticks go through multiple nymphal stages, gradually increasing in size until their final molt to the adult stage. Soft ticks feed several times during each life stage, and females lay multiple small batches of eggs between blood meals throughout their lives. Many soft ticks have an uncanny resistance to starvation, and can survive for years without a blood meal (Furman and Loomis 1984). The entire life cycle of the soft ticks is generally much longer than that of hard ticks, lasting several years. Although there are only around 860 species of ticks worldwide, these arthropods can be found on every continent of the world (Horak et al. 2002). Ticks are second only to mosquitoes in their transmission of pathogens to people, domestic pets, livestock, and wild animals (Dennis and Piesman 2005). They are vectors of a variety of pathogens responsible for human disease including Rocky Mountain spotted fever, Lyme disease, Colorado tick fever, tick-borne encephalitis, babesiosis, tularemia, tick-borne relapsing fever, and others. In addition, pathological sequelae associated with tick bites such as paralysis, toxicosis and anaphylaxis are common. Worldwide public health costs due to tick-transmitted diseases and bites are estimated to be in the range of several billion U.S. dollars per year. Ticks also cause tremendous losses in the livestock industry. For example, tick-transmitted East Coast fever (caused by the protozoan parasite Theileria parva) is responsible for annual losses of 1.1 million cattle and 168 million dollars (Norval et al. 1992). From 1981 to 1993, 72% of all livestock deaths in Tanzania were due to tick-borne diseases (Lynen et al. 2008). The control of tick populations is an effective strategy in the prevention of tickborne infectious diseases. To date, tick control measures have relied heavily on chemical pesticides (acaricides) (Sonenshine 1991, 1993; George 2000). The excessive use of pesticides is problematical and is responsible for environmental contamination, food safety concerns, and the appearance of pesticide resistance in ticks (Matthysse and Lisk 1968; Solomon 1983; Nolan 1990; Barre et al. 1993; Berge et al. 1996; Villarino et al. 2003; Miller et al. 2007; Thullner et al. 2007; Baffi et al. 2008; Ntondini et al. 2008; Rosado-Aguilar et al. 2008; Rosario-Cruz et al. 2009). For example, Miller et al. (2001) reported that brown dog ticks (Rhipicephalus sanguineus) collected from the Corozal Army Veterinary Quarantine Center in Panama were highly resistant to pyrethroid and organophosphate

Anti-tick Vaccine Development: Status and Perspectives

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acaricides, possibly due to the continuous application of these compounds over the course of several years. Thus, it is extremely important to investigate new alternative strategies of tick control. The use of molecular technology to develop anti-tick vaccines is a potentially promising alternative strategy (Willadsen 1987, 2004; de la Fuente and Kocan 2006). Vaccines have been used over the past several decades to control infectious diseases. However, because ticks are the vectors of multiple disease agents, controlling tick-borne illnesses with pathogen-specific vaccines would require developing several different vaccines and is probably implausible. An additional complication to anti-tick borne disease vaccine development is the complexity of the tick life cycle. Generally, both adult ticks and their instars require a blood meal to develop or survive. They often feed on more than one host, potentially transmitting pathogens over a wide range of animals that act as zoonotic disease reservoirs. Thus, a more plausible alternative control strategy would be to develop anti-tick vaccines rather than anti-pathogen vaccines (Willadsen 1987, 1990, 2004; Wikel 1988; Nuttall et al. 2006; Sonenshine et al. 2006). While vaccines against bacteria and viruses protect the host from infection, an anti-tick vaccine would reduce the numbers of engorged ticks. This could have the effect of reducing tick populations as a result of increased mortality and reduced egg production. However, it is important to note that it may not be possible to completely eliminate tick infestations using such a vaccine. The successful development of anti-tick vaccines is complicated, timeconsuming, and challenging. The candidate antigens have to be identified by gene cloning and expression. The expressed, purified proteins must then be characterized and assayed for function. Host immunity triggered by a potential antigen would require evaluation, and if exposure induces a protective host response, the impact of the antigen on tick feeding tested. Finally, demonstration that the vaccine is safe, that it can be produced commercially, and that it can pass vaccine trials and the approval process would be required.

21.1

Interaction between tick and host

Generally, anti-tick vaccine development focuses on two types of antigens: exposed and concealed. Exposed antigens are proteins or polypeptides found in tick saliva and secreted into the body of the host during feeding (Willadsen and Williams 1976; Willadsen 1987; Willadsen and McKenna 1991; Wikel et al. 1994; Willadsen and Jongejan 1999; Nuttall et al. 2000; Nuttall and Labuda 2004; Steen et al. 2006; Francischetti et al. 2009). In contrast, concealed antigens are those proteins or polypeptides that are not secreted into the host such as tick intestinal proteins. Because exposed antigens often play a role in tick feeding, the identification and characterization of the structure and function of tick salivary gland proteins, particularly these injected into hosts, are of central importance in understanding tick feeding and tick-host interactions and crucial to the development of a successful anti-tick vaccine (Sauer et al. 1995; Bowman

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et al. 1997; Jaworski 2003; Bowman and Sauer 2004; Valenzuela 2004; Steen et al. 2006). When acquiring a blood meal, ticks need to penetrate the host’s skin and damage enough blood vessels for the release of blood. The lesion develops gradually with the formation of a hematoma and is caused by neutrophils attracted to the feeding site where they degranulate and induce inflammation (Ribeiro 1987; Kotsyfakis et al. 2006). While feeding, the ticks inject saliva into the host skin. The saliva contains a large number of molecules including a wide array of bioactive proteins that can inhibit host hemostasis, alter vasoconstriction and down regulate inflammation and immune responses such as activation of the alternative complement system (Wikel et al. 1994; Wikel 1996; Bowman et al. 1997; Brossard and Wikel 1997; Barriga 1999; Steen et al. 2006; Mans and Ribeiro 2008). The effects of bioactive salivary proteins benefit the tick in its quest to locate blood and to keep blood flowing without incurring a host response. It is estimated that tick saliva contains over 400 proteins. Only a small fraction of the salivary proteins from hard ticks have been cloned, sequenced, and characterized in recent years (Narasimhan et al. 2002; Valenzuela 2002, 2004; Francischetti et al. 2005; Fukumoto et al. 2006; Ribeiro et al. 2006; AlarconChaidez et al. 2007; Daix et al. 2007; Harnnoi et al. 2007; Paveglio et al. 2007; Schroeder et al. 2007; Kotsyfakis et al. 2008; Schuijt et al. 2008). Over 8,000 expressed sequence tags (EST) sequences from a cDNA library of Ixodes scapularis salivary glands have been obtained (Ribeiro et al. 2006).

21.2

Tick feeding and pathogen transmission

The molecular interaction between microbes and arthropods is important for the transmission and persistence of pathogens in a complex enzootic life cycle. Many arthropod-borne pathogens are transmitted to a host via the saliva of the vector when a blood meal is taken (Munderloh and Kurtti 1995; Bowman and Sauer 2004; Ueti et al. 2009). Tick saliva contains a cocktail of molecules that not only help ticks obtain a blood meal from their vertebrate hosts but promote pathogen transmission as well. For example, the life cycle of the spirochete Borrelia burgdorferi, the agent of Lyme disease in the US, mainly involves I. scapularis ticks and Peromyscus leucopus, the white footed mouse. Ixodes ticks have the capacity to feed on a wide variety of vertebrate hosts and deposit spirochetes into these hosts while feeding. The tick vector I. scapularis plays a key role in maintaining the enzootic life cycle of the bacterium (Wikel 1999; Oliver et al. 2003; von Lackum et al. 2007; de Silva et al. 2009). Larval ticks acquire spirochetes by feeding on infected mice and remain infected in the subsequent nymphal and adult stages of development. A comprehensive understanding of the tick-pathogen interface will also be fundamental to the development of new and novel measures for the control of both tick infestations and tick-borne pathogens (de la Fuente et al. 2008). It is evident that B. burgdorferi evolved several strategies to survive in a complex


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enzootic cycle involving tick vectors and a diverse range of hosts despite strong host antibody responses (Schwan 1996; Schwan and Piesman 2000; Pal and Fikrig 2003; Bubeck-Martinez 2005; Fikrig and Narasimhan 2006; Hovius et al. 2007). Numerous studies have demonstrated that tick salivary gland molecules play a role in Lyme disease transmission. Lyme Borrelia interacts with (adhere, recognize, and respond to) vector ticks and mammalian hosts via its three outer membrane surface proteins, the OspA, OspB, and OspC (Howe et al. 1986; Burkot et al. 1994; Woodman et al. 2008). In the vector-host transmission cycle, the spirochetes alter their outer membrane surface protein structure inside the engorging tick (Schwan and Piesman 2000; Ohnishi et al. 2001). Borrelia burgdorferi upregulates OspA during entry into ticks and this protein contributes to the colonization of spirochetes within the tick gut (Pal et al. 2000, 2001). When a tick feeds on a mammal, B. burgdorferi spirochetes in the tick gut swiftly upregulate OspC, invade the salivary gland of the tick, and are then transmitted to the host dermis along with tick saliva (Schwan et al. 1995). This protein, therefore, has been proposed to play a role in spirochete transmission from the vector (Gilmore and Piesman 2000) and in early mammalian infection (Schwan and Piesman 2000). Grimm et al. (2004) reported that OspC is strictly required for B. burgdorferi to infect mice but not to localize or migrate in the tick. Osps A and B are upregulated during entry into ticks, and evidence indicates that both contribute to the colonization of the bacterium within the gut of the vector (Fingerle et al. 2007). Borrelia burgdorferi sigma54 is required for mammalian infection and vector transmission but not for tick colonization (Fisher et al. 2005). A thorough understanding of the changes in gene expression and the function of the differentially expressed antigens during the life cycle of the spirochete will allow a better control of Lyme disease and the design of effective vaccines to prevent infection (Anguita et al. 2003; Hovius et al. 2007; Tyson et al. 2007).

21.3

Immunity induced by tick saliva (exposed antigens)

It is likely that some of the components of tick saliva are capable of evoking a host immune response, protecting the host from infection, and decreasing the viability of the ticks. However, tick-host and host-pathogen interactions that occur during tick feeding are immunologically complex so a thorough understanding of acquired immunity to ticks is essential for rational development of anti-tick vaccines. It has been observed that tick saliva and salivary gland extracts modulate the immune defense mechanisms of the host (Willadsen 1980; Wikel 1996; Bowman et al. 1997; Valenzuela 2002; Brossard and Wikel 2004; Bowman and Sauer 2004; Kovar 2004). Studies indicate that tick feeding induces host immune regulatory and effector pathways involving antigen-presenting cells, T lymphocytes, antibodies, complement, and other bioactive molecules (Wikel et al. 1994; Wikel 1996; Brossard and Wikel 1997, 2004; Szabó et al. 2004; Kovar 2004). Vancova et al. (2007) reported that chemokine-binding factors from tick saliva regulate the activity of host immunocompetent cells

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during blood-feeding. Narasimhan et al. (2007) demonstrated that the dynamic balance between acquired resistance and tick modulation of host immunity affects engorgement and pathogen transmission. Acquired resistance may impair tick engorgement, ova production, and viability (Wikel 1996). The identification and characterization of proteins expressed in tick salivary glands during feeding and pathogen transmission are the primary rate-limiting step in the detection of suitable antigenic targets for vaccine development (Wikel and Whelen 1986; Barriga 1994; Nuttall et al. 2006). Several investigators have focused their efforts on identifying and expressing tick salivary gland proteins that play a role in host immune responses (Bowman et al. 1997; Valenzuela 2002; Jaworski 2003; Bowman and Sauer 2004; Kovar 2004; Nuttall and Labuda 2004; Santos et al. 2004; Valenzuela 2004; Francischetti et al. 2005; Ribeiro et al. 2006; Steen et al. 2006; Yu et al. 2006; Alarcon-Chaidez et al. 2007; Harnnoi et al. 2007; Prevot et al. 2007; Schroeder et al. 2007; Rachinsky et al. 2008). Narasimhan et al. (2007) discovered that immunity directed against salivary proteins expressed in the first 24 hours of tick attachment is sufficient to evoke all the hallmarks of acquired tick-immunity, to thwart tick feeding, and impair Borrelia transmission. Therefore, defining this subset of proteins will promote a mechanistic understanding of novel I. scapularis proteins critical for the initiation of tick feeding and Borrelia transmission. In addition, Kotsyfakis et al. (2008) observed that guinea pig vaccination against a secreted tick salivary immunomodulator, sialostatin L2, decreased the feeding ability of I. scapularis nymphs. Increased rejection rate, prolonged feeding time, and apparent signs of inflammation were observed for nymphs attached to vaccinated animals, indicating a protective host immune response. These results are encouraging in that they demonstrate the potential value of exposed antigens for anti-tick vaccine development. Although progress has been made in development of exposed anti-tick vaccine, there are no commercial exposed antigen vaccines currently available. Characterization of tick salivary proteins is still one of the central tasks in exposed antigen vaccine development. On the other hand, the development of an exposed antigen vaccine has been impeded by the observation that tick salivary proteins do not necessarily induce significant anti-tick immunity and may, in fact, down-regulate the host immune response (Ribeiro 1989). A co-evolutionary arms race has been taking place for millions of years between ticks, the pathogens they transmit, and their hosts (Hoogstraal and Kim 1985; Steen et al. 2006; Maritz-Olivier et al. 2007). To enhance survival, the host develops immune responses against tick feeding and, in turn, ticks counterattack by evolving mechanisms to minimize host immunity (Decrem et al. 2008; Mans et al. 2008). For example, it has been demonstrated that the soft tick Ornithodoros moubata produces a salivary gland protein capable of inhibiting activation of the complement (C) system (Paesen et al. 1999; Nunn et al. 2005; Schroeder et al. 2007; Roversi et al. 2007; Chmelar et al. 2008; Mans et al. 2008). In addition, Fang and his colleagues in the Center for Ecology and Hydrology at the University of Oxford, UK have characterized, expressed and


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determined the structure of a 16.7 KDa lipocalin (OmCI) derived from O. moubata. This protein binds directly to complement component C5 and is a potent inhibitor of complement activation. Laboratory assays indicate that OmCI also binds to leukotriene fatty acids; most notably leukotriene at sites of inflammation (unpublished data, see summary in Nunn et al. 2008).

21.4

Search for concealed tick antigens of significance

The identification and test of concealed antigens for use as anti-tick vaccine candidates is another strategy. A concealed antigen originating from the absorptive surface of the tick gut and not introduced into the host during tick feeding may be an alternative to exposed antigens and a potential component for use in an anti-tick vaccine. A tick blood meal containing antibodies induced in the host by a concealed antigen vaccine could damage the tick intestine, prevent tick feeding, impair ova production, and even cause tick death. A commercial, concealed antigen anti-tick vaccine, registered as TickGardTm and Gavac, for control of the cattle tick Rhipicephlus (Boophilus) microplus was successfully introduced in Australia, Cuba, Mexico and other Latin American countries (Kemp et al. 1989; Willadsen and Kemp 1989; Willadsen and McKenna 1991; Willadsen et al. 1995). The vaccine contains a tick gut glycoprotein, designated Bm86 that is never injected into the host during tick feeding (Kemp et al. 1989; Rand et al. 1989; Willadsen et al. 1989; Willadsen and McKenna 1991). The glycoprotein is located on the luminal surface of the plasma membrane of tick gut epithelial cells. When used to vaccinate cattle, the vaccine stimulates an immune response that protects cattle against subsequent tick infestation. The Bm86 nucleotide sequence of the cDNA contains a 1982-basepair open reading frame and predicts that Bm86 contains 650 amino acids including a 19-amino acid signal sequence and a 23-amino acid hydrophobic region adjacent to the carboxyl terminus. The main feature of the deduced protein sequence is the repeated pattern of 6 cysteine residues, suggesting the presence of several epidermal growth factor-like domains (Rand et al. 1989; Richardson et al. 1993). Animal model experiments demonstrated that ticks engorging on cattle vaccinated with these inclusion antibodies were significantly damaged as a result of the immune response against the cloned antigen (Richardson et al. 1993). Under field conditions the Bm86-based vaccine was able to control R. (B.) microplus populations (Pipano et al. 2003). Friesian cattle were immunized with two inoculations of anti-tick Bm86 ( TickGardTm) vaccine and were challenged 30 or 90 d later with Boophilus annulatus larvae derived from 1.2 g of eggs. No nymphs or adult ticks were found on the immunized cattle during four weeks after challenge. Repeated infestations (2 to 4) with larvae on three other calves during a period of 160 and 390 d after the immunization did not result in development of nymphal and adult stages. In control, non-immunized cattle infested with corresponding batches of larvae 1380 to 4653 replete adult female ticks were collected. Larvae issued from Babesia bovis-infected female ticks transmitted the

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infection to Bm86-immunized cattle, but the progeny of B. bigemina-infected females did not. The isolation and characterization of Bm86 homologues from other tick species were also explored. Garcia-Garcia et al. (2000) isolated the gene Bm95 from Argentinean strain A cattle ticks, which show low susceptibility to Bm86 vaccine. Similar to Bm86, Bm95 contains seven EGF-like domains and a lipid-binding GPI-anchor site at the C-terminal region. Cattle immunized with the Bm95 antigen from strain A were protected against infestations with Bm86sensitive and Bm86-resistant tick strains, thus suggesting that Bm95 could be a more universal antigen to protect cattle against infestations by R. (B.) microplus strains from different geographic areas. Odongo et al. (2007) characterized two Bm86 homologues (designated Bd86-1 and Bd86-2) from Rhipicephalus (Boophilus) decoloratus, a species closely related to R. (B.) microplus. Two distinct sequences Bd86-1 and Bd86-2 were 95% similar at the DNA level and exhibited 93% identity at the amino acid level. Amino acid sequence identity between Bd86 homologues (Bd86-1 and Bd86-2) and Bm86 was 86% and 85%, respectively. Another Bm86 homologue, designated as the Hl86, was cloned from semi-engorged female Haemaphysalis longicornis ticks in Japan (Liao et al. 2007). The predicted amino acid sequence of the Hl86 gene shows a 37% identity to the Bm86 gene. Hl86 is predicted to be a GPI-anchored membrane-bound glycoprotein with a 19-amino acid signal sequence and a 22-amino acid hydrophobic region adjacent to the carboxyl terminus. The most important feature that Hl86 has in common with Bm86 is the repeated pattern of 6 cysteine residues forming epidermal growth factor-like domains. RT-PCR analysis showed that Hl86 mRNA transcripts are expressed in all the life cycles of H. longicornis, and the expression was found in the midgut of the adult tick. Fang and Xu isolated and cloned a Bm86 homologue (Rs86) from the brown dog tick R. sanguineus (GenBank accession #EF222203). The complete cDNA sequence of Rs86 consists of 2304 nucleotides, including a 92-bp region of untranslated sequence at the 5’-end and a 205 bp fragment of untranslated region at the 3’-end. The amino acid coding region is 1962 bp in length and encodes for a polypeptide of 654 amino acids putatively. Rs86 has 79.8% identity with Bm86 at the amino acid level. The amino acid identities of Rs86 with the other five known Bm86 homologues are from 71.1% to 79.7%. The expression and purification of the Rs86 protein were also attempted using different expression systems. Peconick et al. (2008) explored a synthetic vaccine (sbm7462) against the cattle tick Rhipicephalus (Boophilus) microplus in different strains from South America. Azhahianambi et al. (2009) isolated a Bm86 homologue (named rHaa86) from the hard tick Hyalomma anatolicum. Seven epidermal growth factor-like domains predicted in Haa86 were structurally similar with that of its Bm86 counterpart. The identity between the corresponding epidermal growth factor-like domains of Bm86 and Haa86 ranged from 51.3% to 78.3%. The molecular weight of the rHaa86 was 120–140 kDa, with possible 50–70 kDa glycosylation. The purified rHaa86 was characterized immunologically and evaluated for its immunoprotective potential against homologous challenge infestation in three groups of


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cross-bred calves. Promising immune responses were observed in these animal hosts. The data demonstrated that rHaa86 antigen based vaccine could serve as an effective components in the vaccine-based control of H. a. anatolicum. In addition, the non-Bm86 type of concealed antigens, serine protease inhibitors (RAS-1 and-2) were investigated (Imamura et al. 2006). RASs show promise as anti-tick vaccine candidates. The molecular weight of RAS is around 60–64 kDa. Vaccination of cattle with a combination of rRAS-1 and-2 conferred significant protective immunity against ticks, resulting in 61.4% reduction in nymph engorgement rate, and in 28 and 43% increased mortality rate in adult female and male ticks, respectively. This is the first report on an anti-tick vaccine trial using a combination of two different serpins derived from R. appendiculatus, and using cattle as a natural host. A similar study has been extended to RAS-3 and-4 in the tick Rhipicephalus appendiculatus (Imamura et al. 2008). Seixas et al. (2008) investigated tick Vitelin-Degrading Cysteine Endopeptidase (VTDCE), a peptidase with an active role in Rhipicephalus (Boophilus) microplus embryogenesis for anti-tick vaccine development. VTDCE is found in the tick's eggs and was shown to be the most active protein in vitellin (VT) hydrolysis of the three peptidases. The study indicated that bovines vaccinated with VTDCE induce a partial protective immune response against R.B. microplus infestation. Immunized bovines challenged with R.B. microplus larvae presented an overall protection of 21%, and a reduction in the weight of fertile eggs of 17.6%. The data obtained indicate that VTDCE seems to be important for tick physiology, however, the immunity may not be strong enough and may not be effective for the prevention of tick infestations and population control.

21.5

Summary and perspectives

Ticks are vectors of many diseases of humans, domestic pets, livestock, and wild animals. Control of ticks and tick-borne diseases has relied heavily on the use of acaricides. This chemical-based control method has resulted in acaricide resistance, environmental problems, and food safety concerns. The development of an anti-tick vaccine may serve as an effective alternative strategy in tick and tick-borne disease control. The expectation is that an effective anti-tick vaccine would not necessarily kill ticks but would reduce tick feeding on hosts, impair tick development and egg production, and prevent pathogen transmission. However, compared to anti-bacterial and anti-viral vaccines, the development of an anti-tick vaccine is much more challenging because ticks are multi-cellular organisms with complicated life cycles. Anti-tick vaccine development is a long and complicated process. The search for candidate anti-tick antigens has taken two routes: exposed tick antigens and concealed antigens. There are advantages and disadvantages associated with each type of tick antigens. The exposed antigens, which the tick injects into the host during blood feeding, have mostly been identified by their ability to elicit an immune response during tick infestations, but the immunity induced may not

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strong enough to effectively control tick populations. In some instances these molecules also downregulate host immunity. Concealed antigen vaccines are not secreted into the host during tick feeding. Vaccination with a concealed tick antigen induces specific immunoglobulins that are taken up with the blood meal as the tick feeds. Antibodies interact with the concealed antigen on the surface of the tick gut, and may cause rupture of the gut and reduction of feeding. However, this type of immunity necessitates a time delay between tick ingestion of immunoglobulin and the onset of pathology in the tick gut thus perhaps reducing the effectiveness of the vaccine in preventing tick feeding. Despite the fact that progress has been achieved in the development of anti-tick vaccines in recent years, difficulties remain. The search for new antigen candidates has been inadequate and the isolation and characterization of these antigens remains a central task of monumental proportions. Furthermore, the mechanisms and processes involved in immunity to these antigens are poorly understood. For example, a host vaccinated with a single, isolated antigen may not evoke an immune response sufficient to decrease tick feeding or disease transmission. Results of various investigations suggest that multi-subunit vaccines that target tick salivary gland components and tick gut antigens, as well as the pathogen, may be the most effective because they would target the several facets of both the tick vectors and the pathogens. Alternately, the identification of more effective adjuvants may be another means of inducing a more potent host immune response. The successful use of a commercial concealed tick antigen vaccine for the control of cattle ticks in Australia and Central America over the past decade has demonstrated that anti-tick vaccines have the potential to replace, or at least supplement the use of acaricides for tick control. The development of improved vaccines in the future will be greatly enhanced by the use of new and efficient molecular technologies and by the completion of tick genome projects.

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CHAPTER 22

Hymenopteran Parasitoids and Their Role in Biological Control of Vegetable Liriomyza Leafminers Tongxian Liu, Le Kang, Zhongren Lei and Ricardo Hernandez

Abstract Many Liriomyza leafminers are economically important pests of vegetables, ornamentals and field crops worldwide. At present, chemical insecticides are still the most common method for leafminer control worldwide. The effectiveness of these insecticides has been dogged by their indiscriminate use, impact on natural enemies and the development of resistance to several groups of insecticides and their natural enemies are eliminated by overuse and misuse of insecticides. Extensive investigations of natural enemies of Liriomyza have been conducted worldwide, and more than 150 species of hymenopteran parasitoids have been reported. Several species of parasitoids have been commercially mass-reared and used for biological control of some important Liriomyza species on ornamental and vegetable crops under protected environmental conditions. Conservation biological control promoting natural control and using biorational management strategies should be harmoniously integrated in leafminer management programs. Keywords leafminers, parasitoids, biological control, integrated pest management, vegetable

Tongxian Liu Key Laboratory of Applied Entomology, Northwest A&F University, Yangling, Shaanxi, 712100 China E-mail: [email protected] Tongxian Liu, Ricardo Hernandez Texas AgriLife Research,Texas A&M University System,Weslaco,Texas 78596,USA Le Kang State Key Laboratory of Intergrated Management of Pest Insects and Rodents, Institute of Zoology, Chinese Academy of Sciences, Beijing, 100101, China Zhongren Lei Institute of Plant Protection, Chinese Academy of Agricultural Sciences, Beijing, 100081, China


Hymenopteran Parasitoids and Their Role in Biological Control of Vegetable Liriomyza Leafminers 377

22.1

Introduction

Liriomyza leafminers are economically important pests of field crops, ornamentals and vegetables throughout the world (Parrella 1987; Kang 1996; Kang et al. 2009). More than 20 species of Liriomyza have been reported as being economically important, including L. sativae (Branchard), L. trifolii (Burgess), L. huidobrensis (Branchard), L. bryoniae (Kaltenbach), L. strigata (Meigen), and L. longei Frick (van der Linden 2005; Morgan et al. 2000). At present, synthetic chemical and natural insecticides for leafminer control have been extensively used worldwide by smallholder farmers and large-scale producers alike. Unfortunately, the overuse and misuse of insecticides have cause great concerns in their negative impacts on natural enemies, resistance to common insecticides, and contamination of food crops and the environment and human safety (Lange et al. 1980; Parrella et al. 1984; Sanderson et al. 1989; Ferguson 2004). At present, integrated pest management (IPM) does not seem to have been researched extensively for the management of Liriomyza under field conditions. However, there is now much interest in around the world where the IPM paradigm has been successfully developed for other pests. Extensive investigations of natural enemies of Liriomyza have been conducted worldwide and more than 150 species of parasitoids (Table 22.1). Of those species, several species have been mass-reared and used for biological control of Liriomyza species under confined environmental conditions. In this chapter, we will cover the major species of parasitoids of Liriomyza, especially the species that have been successfully used or have potential to be major biological agents. We will also discuss the progress of conservation biological control that promoting natural control in integrated leafminer management programs.

22.2

Parasitoid species

There are more than 150 species of parasitoids that are known to attack Liriomyza species (Table 22.1). Minkenberg and van Lenteren (1986) reviewed the European species of parasitoids. LaSalle and Parrella (1991) listed 23 Nearctic species of parasitoids of Liriomyza. At least 14 parasitoid species are known from Florida alone (Stegmaier 1966, 1972; Schuster et al. 1991; Schuster and Wharton 1993). Vega (2003) listed 72 species of parasitoids from various countries, and most are from South America. There are several regional reviews for the leafminer parasitoids in Asia and the Pacific Islands (Waterhouse and Norris 1987; Xu et al. 1999; Fisher et al. 2005), including 28 species in Japan (Konishi 1998), 14 species in China (Murphy and LaSalle 1999; Zhu et al. 2000, 2002; Chen et al. 2003; Zeng et al. 2006), 11 species in Indonesia (Rauf et al. 2000), 8 species in Malaysia (Murphy and LaSalle 1999), and many species in Vietnam (Tran et al. 2005b, 2006). Çikman and Uygun (2003) and Çikman et al. (2006) reviewed the parasitoid species in Turkey. Al-Ghabeish and Allawi (2001) found

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that L. huidobrensis was attacked by 16 parasitoid species, while the other leafminers were attacked only by 1–3 species. As shown in Table 22.1, of the hymenopterous parasitoid species attacking Liriomyza, at least 41 species are in the family of Braconidae. At least 6 species have been reported in Cynipidae. At least 3 species in Eucoilidae, 81 species in Eulophidae, and at least 13 species in Pteromalidae. Some parasitoid species are also hyperparasitoids such as Neochrysocharis formosa, which is an endohyperparasitoid of several parasitoids of Liriomyza, and hyperparasitism by N. formosa could be as high as 100% in two months after the first release of D. isaea (Ozawa et al. 2002). Table 22.1

Parasitoids of Liriomyza spp. (from various sources)

Family (No. species)

Genus and species

Braconidae (44 spp.)

Aphidiu ervi Haliday; A. sp. Chorebus amplicator (Nees Esen.); C. artemisiellus Griffiths; C. daimenes (Nixon); C. misellas (Marshall) Dacnusa areolaris (Nees); D. discolor (Föster); D. hospita (Föster); D. maculipes Thomson; D. nipponica Takada; D. sasakawai Takada; D. sibirica Telenga; D. veronicae Griffiths; Mesopora sp. Oenonogastra microrhopalae (Ashmead) Opius agromyzicolae Fischer; O. aridis Gahan; O. basalis Fischer; O. biroi Fischer; O. browsvillensis, O. bruneipes Gahan; O. caricivorae Fisher; O. chromatomyiae Belokobylski & Wharton; O. cinerariae Fisher; O. concolor Szèpligeti; O. dimidiatus (Ashmead); O. dissitus Muesebeck; O. exilis Halliday; O. exiguus Wesmael; O. insulicola Belokoylskij; O. levis Wesmael; O. liromyzae Fischer; O. mirabilis Fischer; O. monilicornis Fischer; O. pallipes Wesmael; O. parvulum (Wesmael); O. phaseoli Fischer; O. propodealis Fischer; O. pulchiventris Fischer; O. scabriventris Nixon; O. suturalis Gahan; O. thoracoseiia; O. turcicus Fischer

Cynipidae (6 spp.)

Charip sp. Cothonapis pacifica Yoshimoto; C. purpureus (Howard) Ganaspidium hunteri (Crawford); G. pusillae Weld; G. utilis Baerdsley (= G. nigrimanus Weld)

Eucoilidae (3 spp.)

Gronotom guamensis Abe & Konishi; G. micromorpha Perkins Nordlanderi plowa Quinlan

Eulophidae (87 spp.)

Achrysocharoides zwoelferi Delucchi Apleurotropis kumatai Kamijo Asecodes delucchi (Boucek) ; A.erxia (walker) Chrysocharisainsliei Crawford; C. caribea Boucek; C. crassiscapus (Thomson) (= C. mallochi Delucchi); C.mallochi Gahan; C. entedonoides Walker (= C. albicans Delucchi); C. flacilla (Walker) (= C. phytomyzae (Brèthes); Euparacrias phytomyzae (Brèthes)); C. gemma (Walker); C. giraulti Yoshimoto; C. ignota Hansson; C. liriomyzae Delucchi; C. oscinidis Ashmead (= C. parksi Crawford); C. orbicularis (Nees); C. pentheus (Walker); C. phryne (Walker); C. pubicornis (Zetterstedt); C. viridis (Nees); C. vonones (Walker, 1839) (= C. brethesi Schauff & Salvo) Chryomatomyia horticola Goureau; C. formosus Westwood; C.okazakii Kamijo; C. smaragdulus (Graham)


(Continued) Family (No. species)

Genus and species

Eulophidae (87 spp.)

Chrysonotomyia recticulata Cirrospilus ambiguous Hansson & La Salle; C. flavoviridis Walker ; C. vittatus Walker (= C. hytomyzae (Ishii)); Closterocerus agromyzae (Crawford, 1913) (= Achrysocharellaagromyzae (Crawford); Derostenus agromyzae Crawford)); C. arizonensis (Crawford) (= Neochrysocharisarizonensis (Crawford)) ; C. beasleyi (Fisher & La Salle) (= Neochrysocharisbeasleyi Fisher & La Salle); C. cinctipennis Ashmead; C. diastatae (Howard) (= C. diastatae (Howard); C. punctiventris (Crawford); Neochrysocharispunctiventris (Crawford)); C. losterocerus erxias Walker (= Asecodeserxias Walker); C. formosus Westwood (= Derostenus fullawayi (Crawford); Achrysocharellavariipes (Crawford); Neochrysocharis formosa Westwood); C. lyonetiae (Ferrière); C. mirabilis Edwards & La Salle; C. okazakii (Kamijo) (= Neochrysocharis okazakii (Kamijo)); C. pictipes (Crawford)(= Neochrysocharis albipes Kurdjumov) C. purpureus (Howard); C. trifasciatus Westwood; C. utahensis Crawford Derostenus sp. Diaulinopsis arenaria Erdös; D. callichroma Crawford Diglyphus albiscapus Erdös; D. begini (Ashmead) (= Solenotusbegini (Ashmead)); D. chabrias (Walker); D. crassinervis Erdös; D. horticola Khan; D. intermedius (Girault) (= Solenotus intermedius (Girault)); D. isaea (Walker); D. minoeus (Walker); D. poppoea Walker; D. pulchripes (Crawford); D. pachyneurus Graham; D. pusztensis (Erdös et Novicky); D. websteri (Crawford) Hemiptarsenus indicus Khan; H. ornatus (Nees); H. unguicellis (Zetterstedt); H. varicornis (Girault) (= H. semialbiclavus (Girault)); H. zanglerii (Walker); H. zilahisebessi Erdös Meruana liriomyzae Boucek Neochrysocharis ambitiosus (Hansson); N. sericae (Walker) Oomyzus liriomyzae Narendran Pediobius indicus Khan; P. metallicus (Nees) (= P. acantha (Walker)) Phytomyza lappae Goureau; Pnigalio katanosis (Pteromalidae); P. katonis (Ishii); P. minio (Waller) (= P. flavipes (Ashmead)); P. phragmitis (Erdös); P. soemius (Waller) Quadrastichus liriomyzae Hansson & La Salle Ratzeburgiola incompleta Bouce Stenomesius japonicas (Ashmead) Sympiesis gordius (Walker) Tetrastichus sp. Zagrammosom americanum Girault; Z. latilineatum Ubaidillah; Z. lineaticeps (Girault) (= Mirzagrammosoma lineaticeps (Girault)); Z. mirum Girault; Z. multilineatum (Ashmead)

Figitidae (2 spp.)

Agrostocynips robusta Ashmead Disprygma pacifica Yoshimoto

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Family (No. species)

Genus and species

Pteromalidae (11 spp.)

Cyrtogaste vulgaris Walker Eupteromalus sp. Halticoptera aenea (Walker); H. circullus (Walker); H. crius (Walker); H. helioponi De Santis; H. patellana (Dalman) Lamprotatus tubero Walker Sphegigaster hamugurivora Ishii Thinodytes cyzicus (Walker) Trichomalopsis oryzae Kamijo and Grissel

22.3

Natural control

Many species of parasitoids play key roles in regulating Liriomyza populations in the nature. Generally, parasitoid assemblages in natural, unmanaged habitats tended to be more species rich than assemblages of parasitoids on leafminers in agricultural habitats, although significant heterogeneity occurs among studies. Johnson et al. (1980a) have shown the importance of parasitoids in the regulation of Liriomyza spp. In the former study, parasitism of L. sativae on tomato by C. oscinidis was found usually low early in crop development and gradually increases as the crop matures. Neuenschwander et al. (1987) found that five indigenous eulophid species plus five other rare parasitoids frequently caused 90% parasitism on L. trifolii; rates were highest in fields free from insecticides. In Victoria, Australia, Bjorksten et al. (2005) found that natural control exerted by local parasitoids was high, with 100% control of L. chenopodii reached in beets within 1–3 weeks of mines appearing and 100% control of L. brassicae within 6 weeks. Although parasitoids can be important in natural control, and in the absence of insecticides usually keep this insect at low levels of abundance (Harding 1965; Trumble 1981), natural control often does not provide adequate suppression, which results in the application of other pest management solutions including augmentative biological control. Interestingly, invading leafminer populations have sometimes been observed to decline naturally after a few years and it has been hypothesized that this is due to the action of local natural enemies (Murphy and LaSalle 1999). Also, the rate at which local leafminer parasitoids can expand their host range can be quite fast. For examples, after the invasion of L. trifolii in Senegal in the early 1980s, many species of parasitoids had been found (Neuenschwander et al. 1987). L. huidobrensis and L. sativae were first found in Indonesia in 1994, and eight species of parasitoids were found in 1998 (Rauf and Shepard 1999) although only one has been found to be common at the present time. Similar diversities have been found on these leafminers in Iran (Talebi et al. 2005), Malaysia (Sivapragasam and Syed 1999), Vietnam (Tran et al. 2005, 2006), Japan (Yano 2004), and China (Wen et al. 2002; Xu et al. 1999). Many species of parasitoids have become common on the leafminers in only a few years (Wen et al. 2002; Tran et al. 2005, 2006).


Conserved biodiversity contains a pool of potential biological control agents. Thus regional biodiversity can contain indigenous parasitoids that will contribute to the control of invading pests. With respect to invading Liriomyza, initial surveys indicate that this certainly is occurring in many countries, although high levels of pesticide usage make this difficult to quantify. Despite the difficulties in quantifying the effect of indigenous parasitoids, they should be considered as a resource and protected as much as possible. Also, one of the largest threats to indigenous species is introduced species (Reid & Miller 1989), and there is concern about the possible effects of introduced biological control agents on nontarget organisms (Howarth 1991). Of particular concern when considering introductions are generalists species which show little discrimination in switching hosts, and most leafminer parasitoids seem to fall into this category. In light of these points we now consider the relative roles that conservation/enhancement and classical biological control might play in an IPM program for Liriomyza spp. in field crops. Interestingly, numerous parasitoid species have been recorded from the leafminers in particular areas. Species of parasitoids found varied greatly among crop systems and geographic locations. In Texas, Chandler (1982) found 8 native hymenopterous species, and a species in Chrysonotomyia accounted for 68.3% of all the specimens collected. In Florida, Schuster and Wharton (1993) found that D. intermedius, D. begini, and N. punctiventris were the most abundant larval parasitoids of L. sativae and L. trifolii (28.8, 26.3, and 15.6%, respectively), and O. dissitus was the most abundant larval-pupal parasitoid (51.8 and 12.6%, respectively). Johnson et al. (1980b) found that Chrysonotomyia punctiventris (J. C. Crawford) and Chrysocharis parksi J.C. Crawford were the predominant larval and larval-pupal parasites, respectively, in Irvine, California. Over a 3-year period of survey on the Helianthus annus L. and Xanthium strumarium L. in northern California, Gratton and Welter (2001) found 16 species of parasitoids, and the most common species were Diglyphus and N. arizonensis. In Senegal, Neuenschwander et al. (1987) found that the most important parasitoids were H. semialbiclavus, which dominated in the 2nd half of the dry season, and two species of Chrysonotomyia, which were abundant in the rainy season. Niranjana et al. (2005) surveyed the parasitoids of L. sativae in Sri Lanka, and found four species of parasitoids, and D. isaea accounted for a parasitism of 58.7%. In Victoria, Australia, Bjorksten et al. (2005) found that the most common parasitoids of Liriomyza were H. varicornis (42.5%), D. isaea (14.6%), C. mirabilis (10.5%), and O. cinerariae (8.5%). Weekly collections from Chinese cabbage infested with L. brassicae and beetroot infested with L. chenopodii, they found H. varicornis and D. isaea were the most abundant in both crops. Habitat diversification and management can be also important for enhancement of Liriomyza parasitoids. For example, weed patches near crops can be important reservoirs for parasitoids (Schuster et al. 1982). Salvo et al. (2005) analyzed the parasitic assemblages of L. huidobrensis were analyzed in relation to natural, urban and cultivated habitats through experimental and comparative methodol-

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ogies. Their results showed that overall parasitism and parasitoid species richness were not lower in simple and disturbed than in complex habitats. Pooled data indicated that parasitism of L. huidobrensis increased in the sequence natural < urban < cultivated on both experimentally exposed and naturally occurring weeds. Small leafminer populations attracted the highest total number of parasitoid species in cultivated habitats. Some degree of habitat specialization was detected in eulophid species which were particularly scarce in cultivated habitats, the reverse being found for braconids. Although many parasitoid species are polyphagous, attacking several dipterous leafminer species, some are strongly influenced by host plants (Murphy and LaSalle, 1999). More generally there is the observation that some parasitoid species seem to be better adapted to particular crop plants. Johnson and Hara (1987) emphasize that effective biological control may depend on matching the most effective natural enemies with a given Liriomyza host and crop. Crop monoculture has been considered one of the factors contributing to the disruption of parasitoids; some of the major parasitoids have particular crop ‘preferences’ and thus their impact may be reduced on ‘non-preferred’ crops. The abundant parasitoids reared from Liriomyza species differed largely among crops. Parasitoids and parasitism are associated with crops and crop phenology, and parasitism can be ranged from as high as 90% on French bean, eggplant, soybean and beet to as low as less than 10% on potato, celery and Welsh onion (Rauf et al. 2000). Parasitoid species and the proportion of each species reared from the leafminers varied with host plants, crop systems and geographic locations (Tran et al. 2006). Gratton and Welter (2001) studied the population dynamics and parasitoid assemblages of L. helianthi Spencer over a 3-year period on the Helianthus annus L. and Xanthium strumarium L. in northern California. They found that the most common species, Diglyphus spp. and N. arizonensis show no bias in association with a particular leafminer or plant species, and however, one parasitoid, C. ainsliei Crawford, was strongly associated with H. annuus.

22.4

Classical biological control using parasitoids

Classical biological control of Liriomyza under field conditions achieved mixed results, and most works have been targeted against L. trifolii and L. sativae (Murphy and La Selle 1999), and works on biological control in greenhouses have mostly been developed in horticultural industries under protected environments around the world (Chow and Heinz 2005c; van der Linden 2005). In general, biological control efforts under greenhouse conditions have focused on augmentation. Most successes being achieved in greenhouses and these have been successful because of the closed protected conditions and regulated climate (Minkenberg and van Lenteren 1986). In many cases, an introduced parasitoid has failed to establish or the result of the introduction is not known. However, a few notable successes on vegetables have been achieved. Of more relevance here are the biological control programs that have been conducted


against Liriomyza under field condition or in under partly covered crops. Biological Control under Field Conditions. Biological control strategies appropriate for Liriomyza spp. in field vegetables could include conservation or enhancement of local natural enemies and/or the introduction of appropriate natural enemies from the area of origin of the pests or from related leafminers from other areas. However, these strategies are not mutually exclusive, as it is clear that any introductions should take into account the existing local natural enemy community. Some notable successes in the control of Liriomyza spp. have been achieved on some of the Pacific Ocean islands – Hawaii, Tonga and Guam. In Hawaii, a particularly successful program of parasitoid introductions against L. trifolii and L. sativae has been achieved in the late 1970s and 1980s. G. utilis was introduced in 1977 has a major impact on L. trifolii and L. sativae on watermelons and may be important on L. trifolii on celery, and N. diastatae also had a significant impact on both leafminers on several vegetable crops (Johnson 1993). In Tonga, G. utilis and C. oscinidis were released in 1988 for the control of L. trifolii on watermelon, pumpkin, tomato, bean and Irish potato with great success (Johnson 1993). In Senegal, there was a high mortality in the shipments of parasitoids and thus it was difficult to establish healthy cultures (Neuenschwander et al. 1987). Johnson and Hara (1987) reported that the predominant parasites reared from the 4 major species of Liriomyza infesting 12 different host crops in North America and Hawaii. They found that no single parasitoid species was the predominant biological control agent in most crops. D. begini, H. circulus and C. punctiventris were either the 1st or 2nd most reared species in 60.9, 26.1 and 21.7% of the studies, respectively. Because of uneven distribution of parasitoids among crops, it is suggested that effective biological control may depend on matching the 'most effective' parasitoid species complex with a given Liriomyza species and crop. Augmentative Biological Control. Yano (2004) summarized current status of implementation and research about augmentative biological pest control in protected culture in Japan. Augmentation involves efforts to increase populations of natural enemies. Inoculative augmentation refers to the application of an indigenous agent to enhance subsequent buildup in the biocontrol agent population, whereas inundative augmentation refers to the mass application of an agent with the primary objective of high initial kill. In both economic and ecological terms, a classical biological control agent that becomes established and exerts a controlling influence on a pest over an indefinite time period is the ideal control agent. Because parasitoids often provide effective suppression of leafminers in the field when disruptive insecticides are not used, there has been interest in release of parasitoids into crops. This occurs principally in greenhousegrown crops, but is also applicable to field conditions. The ultimate success of augmentative biological control may depend on releases of biological control agents that maximize establishment, are released in synchrony with the host, and can be integrated into integrated pest management programs in conjunction with

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insecticides. Thus, determining optimal release rates that maximize the effectiveness of natural enemies can increase the effectiveness of augmentation biological control and increase its potential economic benefits. The attitudes regarding the augmentation of natural enemies for biological control have been changed. The practical application of this method has been hindered by the high cost of natural enemies, problems associated with the availability and quality of natural enemies, lack of rigorous research on success versus release rates and economic analyses, and the belief that such a strategy utilizes natural enemies as a pesticide, so taking the ecology out of biological control. Mass Rearing. Mass rearing sufficient host, Liriomyza, is essential for massrearing parasitoids. Rearing quality Liriomyza needs to have quality host plants under ideal environmental and nutritional conditions. The ideal host plants should be the ones that can be easily propagated and maintained, be attractive to females for oviposition, bear more leafminer larvae, and produce more and better leafminers. Various host plants have been used to rear Liriomyza, including lima bean plants (Webb and Smith 1970a; Petitt and Wietlisback 1994), tomato plants (Ushchekov et al. 2000), and cowpea plants by Jeyakumar and Uthamasamy (1997) under different environmental conditions. The ideal environmental conditions for growing host plants relatively higher temperature (252°C) and higher humidity (80%–95% RH), and longer lighting (14–16 h). Other conditions, including fertilization, irrigation, soil, or potting media are also important to grow quality host plants. Several species of parasitoids of Liriomyza have been mass-reared or attempted mass-rearing for augmentation as biological agents of Liriomyza (Table 22.2). The mass-rearing procedures involve in maintaining host plants, rearing Liriomyza, and the parasitoids. Effective mass rearing systems of parasitoids are essential to successful biological control because cost of parasitoid production is often too high and lowering the cost is crucial to economically reasonable practices of biological control. The commercial natural enemies is likely to be high-cost until mass production can be scaled up to meet demand, and we would expect use to be focused on environmentally sensitive areas. Because of this limitation, and because its use against field Liriomyza is unlikely, the demand for parasitoids and other natural enemies may well be much more uniform than the demand for chemical insecticides. Overproduction of males in mass rearing of parasitoids contributes to higher costs for biological control because only females directly kill pests. Ode and Heinz (2002) found that host size positively affects both male and female wasps, and females laid more daughters in larger hosts and more sons in smaller hosts. They developed a technique by presenting female D. isaea groups of sequentially larger leafminer hosts (L. huidobrensis) to attack; they are able to generate progressively more female-biased sex ratios. After three days of providing increasingly larger hosts, we were able to reduce the sex ratios produced by individual females from 57% male to 36% male; sex ratios produced by groups of


Table 22.2 Species of parasitoids that have been mass-reared and mass-released for biological control of Liriomyza spp. Parasitoids

Rearing hosts

Chrysocharis flacilla

L. huidobrensis

Dacnusa sibirica

L. bryoniae

Dacnusa sibirica

Host Plants Broad bean (Vicia faba)

Target L. huidobrensis

Tomato (Solanum lycopersicum) L. bryoniae Chrysanthemum

L. trifolii

Dacnusa sibirica

Phytomyza caulinaris

Persian buttercup (Ranunculus asiaticus)

L. huidobrensis

Diglyphus begini

L. trifolii

Lima bean (Phaseolus lunatus)

L. trifolii

Diglyphus begini

L. trifolii

Chrysanthemum

L. trifolii

Dacnusa sibirica

L. bryoniae

Tomato

L. bryoniae

Diglyphus isaea


Persian buttercup

L. huidobrensis

Diglyphus isaea

Chrysanthemum

L. trifolii

L. langei

Chrysanthemum

L. langei

Ganaspidium utilis

L. trifolii

Lima bean (Phaseolus lunatus)

L. trifolii

Halticoptera helioponi

L. huidobrensis

Broad bean (Vicia faba)

L. huidobrensis

Neochrysocharis formosa

L. trifolii

Kidney bean (Phaseolus vulgaris)

L. trifolii

Opius pallipes

L. bryoniae

Tomato (Solanum lycopersicum) L. bryoniae

Opius pallipes


Persian buttercup (Ranunculus asiaticus)

L. huidobrensis

Phaedrotoma scabriventris

L. huidobrensis

Broad bean (Vicia faba)

L. huidobrensis

Diglyphus isaea

females dropped from 64% male to 45% male. Chow and Heinz (2005b, 2006) developed a similar technique as Ode and Heinz (2002) to generate less malebiased sex ratios for L. langei. They used chrysanthemum as the host of L. langei and D. isaea. Presenting individual females with only large host increased mean sex ratio from 32 to 67% male over 2 days, and with progressively larger hosts over 1 or 2 days reduced mean sex ratio from 90% to 100% males to less than 30% males. Using both small and large hosts produced similar numbers of parasitoids as using only large hosts, but male cohorts reduced from 66% to 56%. They found that using both small and large hosts produced slightly lower percentage of males than using only large hosts. Therefore, the use of both small and large hosts for mass rearing of D. isaea could reduce actual costs of females by 23%. This technique can reduce overproduction of males in D. isaea with no compromise in biological control efficacy. Storage. Duration of storage of parasitoids can be affected by temperature, duration and host plants. Chien et al. (2005) investigated the suitable life stages and conditions for storage of H. varicornis, and offspring production, and found that maintaining a high fertility of H. varicornis for mass production, the female wasps could be stored at 15°C for 10 days or at 15°C for 20–30 days. For inoculative field release for host-killing or-feeding and parasitization, H. varicornis female wasps could be stored at 15°C for 10–30 days or 25°C for

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10–15 days and at 15°C for 40 days or 1-day-old pupae at 10°C for 1–4 weeks. Augmentation. It may, under some circumstances, be possible to augment local Liriomyza parasitoid populations. For example, in Western Sumatra, Indonesia, extension workers have successfully worked with farmers in areas of low pesticide usage to redistribute parasitized leafminers on crops such as cauliflowers to fields where parasitoids are absent or have low activity (Zamzami 1999). Inundative augmentation of parasitoids in greenhouses. Most effort in the biological control of invasive leafminers under field conditions has focused on the classical approach, i.e. the introduction of natural enemies from the area of origin of the pest. Several parasitoid species have been mass-released for controlling Liriomyza under different crop systems, including D. sibirica (Hendrikse 1980), D. begini (Parrella et al. 1989; Rathman et al. 1991), and D. isaea (Chow and Heinz 2006). Diglyphus isaea. D. isaea is commercially available. Inundative release of D. isaea against L. trifolii has been part of IPM program for management pests under greenhouses (Ozawa et al. 1993, 1999; Rodriguez et al. 1997; Ozawa et al. 1999). It has proved that D. isaea is effective against L. trifolii on tomato in greenhouses (Ozawa et al. 1999). Cabitza et al. (1993) released D. isaea to control L. trifolii, and found it was effective in spite of high levels of infestation of the pest (which reached 74 mines/plant), and the level of parasitism was up to 100%. Ushchekov (1994) reported that L. bryoniae in the summer-autumn rotation could be effectively reduced by a single release of 1 female/15 larvae of Liriomyza. Ushchekov et al. 2000 described a technique is outlined for the mass-rearing of the parasitoid D. isaea against L. bryoniae, using several varieties of tomato and bean. Ulubilir and Sekeroglu (1997) found that the release rate of D. isaea at 100 adults/100 m2 (at 10 different spots, each 10 adults/10 m2), larval populations of L. trifolii decreased to < 1 larva/leaf, which was similar to the larval densities at the cyromazine-treated plots. Sampson and Walker (1998) set up on commercial tomato nurseries to test whether the use of a simple pest density threshold for D. isaea release could be used to improve early parasitoid establishment in commercial crops. They used an action threshold of one new mine per plant per week to trigger releases of D. isaea for three times at 7-d intervals and found that D. isaea established faster, and number of parasitoids used and control costs were reduced. L. bryoniae populations were controlled without economic damage and no chemical treatments were required. Diglyphus begini. At present, D. begini is another species is not commercially available. It has been found that twice-weekly releases of the parasitoid D. begini over a period of three weeks to control L. trifolii on marigolds, and the leafminers were effectively controlled for six weeks after the first release and was maintained at a low level for a further eight weeks (Heinz et al. 1988). Later, they were inundatively released D. begini for control of L. trifolii on marigolds, and the leafminer population reduced to approximately zero within 8 weeks of the first


release, and remained at low level for the duration of the crop (Heinz and Parrella 1990). D. begini has also been successfully released for biological control of L. trifolii on greenhouse-grown chrysanthemums (Parrella et al. 1992). D. isaea and D. sibrica. Liriomyza trifolii has been successfully used to contol leafminers on tomato in greenhouses with > 90% parasitism (Ozawa et al. 1993; Matsumura et al. 2001). Landi (1993) reported that the release of D. isaea and D. sibirica resulted in 30% of plants had mined leaves by L. trifolii on chrysanthemum at the time of harvesting, which was slightly higher compared with plants treated with insecticides. Abd-Rabou (2006) reared and released D. isaea and D. sibirica imported from The Netherlands. A total of 90 000 of these parasitoids were on cucumber and tomato in greenhouses to control L. trifolii. The parasitism rates of D. sibirica reached 11.6 and 7.2% at the eleventh week from the release date, on cucumber and tomato, respectively. Also, the parasitism rates of the European strain of D. isaea increased until it reached 2.1 and 1.4%, at the tenth week from the release date, on cucumber and tomato, respectively. D. sibirica and the European strain of D. isaea can be established in Egypt. Hemiptarsenus varicornis. This species has been used for biological control against L. on cherry tomatoes in greenhouses, and was practically effective on tomatoes in greenhouses (Ozawa et al. 2004). Other species. Neochrysocharis formosa and O. pallipes have also been massreleased for control Liriomyza. Hondo et al. (2006) determine that the most suitable release density for the host was approximately 50 larvae or 5 pairs of L. trifolii adults/plant, and for N. formosa, 10 adults/50 host larvae. The use of O. pallipes resulted in better control of L. bryoniae (Schelt and van Altena 1997).

22.5

Inoculative releases

Neuenschwander et al. (1987) released nine species of parasitoids to control L. trifolii in Senegal in 1982 and 1983. Although many were recovered shortly after release, only O. dissitus was recovered in later samples and became relatively abundant. Boot et al. (1992) developed a deterministic model to simulate population growth of L. bryoniae and the parasitoid D. isaea. The model has two driving variables, ambient temperature and leaf nitrogen content of the tomato plant. Results of a glasshouse experiment were used to validate the model. The timing of successive generations of leafminers was simulated accurately over four generations. Population growth of leafminers was correctly simulated during the first two generations, but overestimated in the third generation. Mortality of leafminers due to parasitism was overestimated in the first generation after introduction of parasitoids: 73% instead of the observed 30%. A nearly 100% mortality of leafminers was correctly simulated in the second generation after introduction of parasitoids. Sensitivity analysis was performed for three types of variables: driving variables, temperature and leaf nitrogen content; parasitoid traits, searching efficiency and allocation of attacks to host feeding and

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oviposition, and introduction strategies for biological control, timing, number of releases and number of parasitoids per release. Population growth was sensitive to temperature, leaf nitrogen content, searching efficiency of parasitoids and numbers of parasitoids released. Ozawa et al. (2001) determined the effectiveness of the inoculative releases of D. isaea and D. sibirica at 47–125 wasps per ha, in controlling L. trifolii in greenhouse experiments. Selective insecticides were applied in biologically controlled greenhouses while various insecticides, including non-selective insecticides, were applied in chemically controlled greenhouses. The density of L. trifolii larvae in biological control greenhouses was maintained at the same level as that in chemical control greenhouses, and the mortality of the leaf miner increased up to 100%. The damage to tomato plants was not severe, and fewer insecticides were applied.

22.6

Release strategy

Parrella et al. (1992) and Heinz et al. (1993) developed a predictive mathematical model to utilizing augmentative releases of D. begini for biological control of L. trifolii on greenhouse-grown chrysanthemums. Their model determined the appropriate release rate necessary to reduce pest densities to < 1 larva/1000 chrysanthemum leaves within 40 days of planting, after which time aesthetically important foliage forms on chrysanthemums. Validation studies of the predictions provided mixed results. Following the release rates generated by the model, L. trifolii larval densities were not significantly greater than 1/1000 leaves 40 days after planting. However, the model did not always trace the succession and magnitude of pest population fluctuations accurately. Two factors probably contributed to errors in prediction: the assumptions inherent in the model were not met during the validation trials; and the leafminer subroutine of the model could not accurately predict L. trifolii densities in the absence of D. begini. In spite of these errors, leafminer damage to the harvested foliage was significantly lower in the treatments receiving D. begini releases than in the control treatments. In addition, when the model was tested in a commercial cut chrysanthemum greenhouse in California, L. trifolii was successfully controlled. Recommendations have been developed for Diglyphus and Dacnusa, specifying release rates per unit area for preventative or curative releases. However, these wasps are expensive, and other pests are not controlled, so only a few growers are utilizing these parasitoids regularly. Combining the parasitoids with nematodes or fungi for control of other major floricultural pests is now being investigated.

22.7

Host feeding

In biological control, host killing by host-feeding is profitable in the release step, but becomes unprofitable in the mass-production step because it does not result


directly in production of progeny. Hondo et al. (2006) found that seven species of eulophid parasitoids (P. katonis, H. varicornis, D. isaea, D. minoeus, D. pusztensis, C. pentheus and N. formosa) are solitary and idiobiont parasitoids, and their adults kill hosts directly by feeding on the hosts. Each female of an introduced and a native D. isaea could feed and kill up to 466 and 239 L. trifolii larvae at 20°C, respectively, in their lifespan. Parrella et al. (1982) found that D. intermedius, D. begini and C. parksi killed more L. trifolii larvae than parasitized. Chien and Ku (2001) found the native parasitoids of L. trifolii from Taiwan, H. varicornis, C. pentheus, C. okazakii, N. formosa, exhibit host-feeding except Opius sp. With parasitization or host-feeding, all five of these parasitoids have host instar preference, especially preferring to oviposit on third instars. But host selection behavior affects the ratio of idiobiont female progeny only.

22.8

Factors influencing biological control of Liriomyza

Although some notable successes in the control of Liriomyza spp. have been achieved on some of the Pacific Ocean islands, most parasitoid introductions have failed on these islands and elsewhere. Part of the failure relates to shipping techniques but other factors are important. Using natural enemies for management of Liriomyza spp. on vegetables and ornamental plants under greenhouses have different approaches. Fruiting vegetables could tolerate some foliar leafminer damage with no significant yield lose, whereas most ornamental plants have almost zero tolerance for leafmines. Therefore, biological control on greenhouses fruiting vegetables is likely. Low tolerance of pest damage is the main reason for difficulty in adopting biological control in flower production. The costs for mass-rearing parasitoids varied greatly, Rathman et al. (1991) reported that the recurring cost of producing 1,000 parasitoids/day was estimated as $7.2 for Ganaspidium utilis compared with $19.2 for another Liriomyza parasitoid, D. begini. Similarly, Parrella et al. (1989) reported that the daily cost (recurring costs only) to produce 1000 parasitoids was $19.4. The lack of a system for supplying enough natural enemies of good quality at reasonable costs was the limiting factor in biological control because private companies may not interested in biological control. In the augmentative biological control of pests, living natural enemies are released artificially. Most natural enemies are effective against only a small group of pest species. In addition, the most important technical factor in commercializing natural enemies is integrating the use of natural enemies with other control measures that are intended to control other pests on a crop. If the control measures for other pests are incompatible with the use of natural enemies, the practical use of natural enemies is almost impossible. Since it is impossible to control all pest species with natural enemies, the integrated use of natural enemies and other control measures, which include cultural control and the use of selective pesticides, is essential for commercial application.

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There are overwhelming evidences that chemical information play the most important role in host location process in tritrophic system of plant-leafminerparasitoid interaction. For instance, Dicke and Minkenberg (1991) provided evidence that volatile blend from leafminer-infested tomato leaves affected the behavior of D. sibirica in the absence of visual cues. Petitt et al. (1992) also reported that O. dissitus preferentially landed on leafminer-infested, rather than non-infested lima bean plants and this preference was confirmed by choice experiments in a four-armed olfactometer without any visual cue. Behavioral studies have demonstrated that odors and headspace volatiles from plant-leafminer complexes play an important role in host location of leafminer parasitoids (Zhao and Kang 2002; Wei and Kang 2006). The chemical cues for attraction are often composed of complex blends of leafminer-induced volatiles (Wei et al. 2007), thus making it difficult to understand the role of specific compounds in host location by parasitoids (Wei et al. 2006). However, it has been found that the olfactory sensilla of parasitoids only responds to a limited number of the compounds released by insect-damaged plants (Wei and Kang 2006). Recently, electrophysiological techniques have been successfully used to understand the underlying mechanism of odor perception of parasitic wasps of Liriomyza. For example, in a study involving the system of the vegetable leafminer, L. sativae, and D. isaea, Zhao and Kang (2002b) found that neither the healthy host nor non-host plants elicited distinctive electroantennogram (EAG) responses in the parasitoid. Wei and Kang (2006) proved that six-carbon alcohol and ester from mechanically damaged plants elicited strong EAG responses of parasitoids. Meanwhile, they identified nine EAD-active volatiles from the headspace extracts from both Liriomyza huidobrensis and L. sativae secondinstar larvae-damaged beans by GC-EAD techniques. Host and nonhost plant extracts, botanical insecticides, and elicitors induced compounds as repellent/ deterrent have been utilized to push leafminers away from agricultural crops. At the same time, some universally induced compounds as attractive/stimulant stimuli to pull parasitoids to leafminer-damaged plants have also been proposed. In push strategies, host and nonhost plant extracts (i.e. neem, Chinaberry tree, and essential oils), botanical insecticides (i.e., neem-based insecticide, rotenone), and elicitors (i.e. Jasmonic Acid) induced compounds as repellent/ deterrent have been used to push adult leafminers away from agricultural crops, to take effect of toxicity to the larvae of leafminers, and to be compatible with biological control agents (Weintraub and Horowitz 1997; Hossain and Poehling 2006). In pull strategies, the above examples address how leafminer-induced volatile blends or some key volatile compounds attract parasitic wasps of leafminer in laboratory (Wei and Kang. 2006; Wei et al. 2007). It is expected that application of these cues could manipulate the behavior of parasitoids and thus improve its effect in biological control of leafminer pests in the field. Others. Biocontrol agents used in such greenhouses must be highly adaptable to extreme temperatures for exotic natural enemies (van Lenteren 1986).


Therefore, it is imperative to evaluate their thermal tolerance, particularly in relation to development and reproduction throughout their lifetimes (Hondo et al. 2006). Yano (2004) discussed some social limiting factors in biological control, including low tolerance of pest damage, supply of natural enemies, poor advisory service for farmers, and need for registration. However, many species of arthropod natural enemies have been registered, and a sufficient number of imported or indigenous natural enemies are now supplied by private companies. An advisory service for farmers is very important if a good performance of natural enemies for promoting biological control is to be obtained.

22.9

Interactions among natural enemies

Generally, when releasing mass-reared natural enemies into a crop system of existing natural enemy populations, competitive interactions are likely to occur. Parasitoids-Parasitoids. Liriomyza parasitoids may interfere with each other when larval ectoparasitoids (e.g., Diglyphus spp.) and larval-pupal endoparasitoids (e.g., C. oscinidis and G. utilis) are present in a cropping system. Larval ectoparasitoids are able to parasitize leafminers that already contain a living endoparasitoid larva, resulting in the death of the endoparasitoid. Endoparasitoids do not parasitize leafminers with ectoparasitoids because the parasitized hosts will not pupate so the endoparasitoids can complete their life cycles. Bader et al. (2006) assessed the influence of two commercially available parasitoids D. isaea and D. sibirica attacking L. langei on chrysanthemum. They concluded that levels of interspecific competition among parasitoid species were undetectable at leafminer densities typical of field-grown ornamental crops (low densities), and thus, the efficacy of one species released into a backdrop of potentially competing parasitoids did not negatively or positively affect the outcome of the augmentative biological control, nor was there a positive outcome; however, crop quality at harvest was influenced. Parasitoids-Nematodes. A combination of entomopathogenic nematodes and parasitoids has been found more effective than either natural enemies used alone. Sher and Parrella (1999) found adult females of D. begini are able to detect and avoid ovipositing on nematode-infected hosts, and paralysed L. trifolii larvae. However, Head et al. (2003) found that 98% of eggs laid by the female wasps were deposited alongside healthy larvae although adult D. isaea did not discriminate between the healthy and S. feltiae nematode-infected leafminer larvae of L. huidobrensis for host feeding, indicating a synergetic effect when the entomopathogenic nematodes and the parasitoids are used together. Negative interferences are also seen, however, with nematodes decreasing the likelihood of D. begini developing to adults and nematodes directly infecting and killing D. begini larvae (Sher and Parrella 1999). Head et al. (2003) also found that in intact leaf mines of L. huidobrensis, larvae already parasitized by D. isaea which had developed to the larval or pupal stage, and D. sibirica in the larval stage were subsequently also infected by S. feltiae following foliar application. This reduced

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the potential of the wasps to survive until the adult stage. In the same study, they also found that a soil drench of imidacloprid did not cause a significant reduction in the number of D. sibirica, which survived the treatment and developed to adult emergence.

22.10

Host searching behavior and response to plant odors.

Although host location cues of parasitic wasps include visual, chemical, acoustic, and contact and taste cues, overwhelming evidences show that chemical information play the most important role in the tritrophic system of plantleafminer-parasitoid interaction. For instance, Dick and Minkenberg (1991) found that volatile blend from leafminer-infested tomato leaves affected the behavior of D. sibirica in the absence of visual cues. Petitt et al. (1992) also reported that O. dissitus preferentially landed on leafminer-infested rather than non-infested potted lima bean plants. The chemical cues for attraction are often composed of complex blends of leafminer-induced volatiles (Wei et al. 2007), thus making it difficult to understand the role of specific compounds in host location by parasitoids (Wei et al. 2006). However, a lot of researches has revealed that the olfactory sensilla of parasitoids only responds to a limited number of the compounds released by insect-damaged plants thereby substantially reducing the number of compounds that require testing (Wei and Kang 2006). Plant volatile compounds that have elicited antennal responses were also attractive to parasitoids in behavioral experiments (Wei and Kang 2006). Zhao and Kang (2002) found that neither the healthy host nor non-host plants of the L. sativae elicited distinctive electroantennogram (EAG) responses in D. isaea; odors of physically damaged leaves, whether host or non-host plants, elicited strong EAG responses of the leafminer and its parasitoid. This study implied that physical treatment can induce EAG-active compound from plant, which is distinct from healthy one. Wei and Kang (2006) proved that six-carbon alcohol and ester from mechanically damaged plants elicited strong EAG responses of parasitoids.

22.11

Parasitoid conservation

The modern pesticides have provided potent means in suppressing Liriomyza and numerous other pests. However, a biologically-based IPM strategy would seek to conserve or enhance natural enemies, particularly parasitoids because the massive overuse and misuse of insecticides have resulted in numerous problems, including increasing costs, pesticide resistance, contamination to environments, toxic to humans and non-target organisms. The differential destruction of natural enemies of Liriomyza through insecticide use was first reported in the early 1950s during the first leafminer outbreaks in North America. Hills and Taylor (1951)


found that repeated applications of DDT against L.sativae were reduced the parasitoid population, which resulted in a pest outbreak. Subsequent studies with many chlorinated hydrocarbon and organophosphorous insecticides have confirmed this conclusion (e.g. Oatman & Kennedy 1976). There is an extensive recorded literature concerning the compatibility of using biological control agents and selective insecticides for management of Liriomyza under different crop systems. The most commonly used insecticides for management of Liriomyza species include abamectin, cyromazine, spinosad, imidacloprid. Their effects on the major parasitoids are summarized. Avermectins. Abamectin is a mixture of natural fermentation products, avermectins, derived from the soil bacterium Streptomyces avermitilis. Many studies found that abamectin is not harmful to C. humilis,H. varicornis and O. chromatomyiae (Hidrayani et al. 2005), D. isaea (Chen et al. 2003), D. intermedius, D. begini, C. punctiventris, C. parksi and C. ainsliei and H. circulus under field conditions (Trumble 1985), and is slightly harmful to H. varicornis, D. isaea, Gronotoma micromorpha Perkins and Opius sp. (Prijono et al. 2004). In contrast, abamectin has negative effects on many species of parasitoids, including causing 100% mortality of N. okazakii on 12 h old dry residue (Huang et al. 1999), reducing adult D. isaea female longevity up to 5 days after application on chrysanthemum leaves and lethal for D. isaea larvae when applied directly to larvae or when contaminated leafminer larvae were consumed by parasitoid larvae (Kaspi and Parrella 2005), affecting the development of O. chromatomyiae and N. formosa, and strongly affected N. formosa adult emergence when topically applied at different immature stages of N. formosa (Hossain and Poehling 2006), and causing significant mortality to larvae and pupae of H. varicornis and D. isaea (Bjorksten and Robinson 2005). Weintraub and Horowitz (1998) and Weintraub (1999, 2001) reported that topical spray of abamectin significantly reduced D. isaea populations, but recovered soon. Kaspi and Parrella (2005) found that emergence of D. isaea was not affected by abamectin treatments when applied to chrysanthemum plants that contained parasitoid larvae, and the longevity of these emerged adults was not affected by abamectin applications. Azadirachtin. The active ingredient, azadirachtin (derived from the neem tree), acts as an insect growth regulator and is relatively non-toxic to natural enemies of Liromyza. Chen et al. (2003) found that azadirachtin had little effect on the population of the parasitoids, and its parasitism was significantly higher than that in the water control and other insecticide treatments. Hossain and Poehling (2006) tested side effects of two azadirachtin formulations (NeemAzalU, 17% azadirachtin and NeemAzal-T/S, 1% azadirachtin) on O.chromatomyiae and N. formosa, and found that adult emergence of O. chromatomyiae from parasitized L. sativae in NeemAzal-U (0.75, 1.5, 2.25 and 3 g/l) drenched soil was only slightly lower than that from untreated control hosts. Development of O. chromatomyiae was very much affected from topical applications of NeemAzalT/S. However, spraying of tomato leaves with NeemAzal-T/S revealed no detrimental effect on the adult emergence of N. formosa.

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Benzoylurea pesticides. Lufenuron, a benzoylurea pesticide, inhibits the production of chitin in larval insects. Tran et al. (2004) found that the numbers of host findings by female N. formosa, ovipositor insertions and host feedings were significantly lower on the leaves treated with lufenuron than the control. Tran et al. (2005) also found that the glass-film of lufenuron was harmful to N. formosa with reduced survival and longevity. Bionic pesticides. Dimehypo is a bionic pesticide. The dry residue of dimehypo caused 100% mortality of N. okazakii in 12 h (Huang et al. 1999). Prijono et al. (2004) recommended that it should be used cautiously when the parasitoids, H. varicornis, D. isaea, Gronotoma micromorpha Perkins and Opius sp., present. Carbamates. Carbosulfan and methomyl are commonly used insecticides for controlling Liriomyza species. Applications of carbosulfan decreased rates of parasitism by H. varicornis and O. chromatomyiae, and reduced numbers of C. humilis (Hidrayani et al. 2005). Trumble (1983) found that methomyl was not effective against the leafminer and reduced the seasonal percentage parasitism by reducing the survival of adults of D. intermedius, D. begini, C. punctiventris, C. parksi and C. ainsliei and H. circulus. Oatman and Kennedy (1976) reported that methomyl induced outbreak of L. sativae on tomato, and similarly, Ozawa et al. (1993) found that one application of methomyl eradicated D. isaea and the density of the leafminer increased. Neonicotinoids. Imidacloprid is one of widely used neonicotinoids with relatively low human toxicity. Ozawa et al. (1998) found that imidacloprid has a harmful effect on the attacking behavior of D. isaea against L. trifolii and topically application of imidacloprid decreased parasitism by D. isaea at a comparatively low level (Chen et al. 2003). Tran et al. (2004) found that N. formosa adults spent more time resting near or away from hosts and less time foraging for hosts on the leaves treated with imidacloprid than on the other leaves, and as a result oviposition and host feeding were reduced. Tran et al. (2005) found that the glass-film of imidacloprid was harmful to N. formosa and the parasitoid survival rapidly decreased with time, but imidacloprid is not harmful to parasitoid when imidacloprid is used through irrigation system. Organohydrochlorides. Cartap hydrochloride is a derivative of nereistoxin which is a naturally occurring insecticidal substance isolated from the marine segmented worms Lumbrinereis heteropoda and L. brevicirra. Application of Cartap hydrochloride significantly decreased the number of and parasitism by D. isaea and maintained these at a comparatively low level (Chen et al. 2003). Organophosphates. Chlorpyrifos, dichlorvos, dethamidophos, profenofos are wide-spectrum and highly toxic insecticides. Chlorpyrifo has been found harmful to H. varicornis, D. isaea, Gronotoma micromorpha Perkins and Opius sp. (Prijono et al. 2004). Its dry residue caused 100% mortality of N. okazakii in 1 h (Huang et al. 1999). Methamidophos significantly decreased the number of and parasitism by D. isaea and maintained these at a comparatively low level (Chen et al. 2003). Applications of profenofos decreased parasitism by H. varicornis and


O. chromatomyiae, and reduced numbers of C. humilis (Hidrayani et al. 2005). Pyrethroids. Bifenthrin, fenpropathrin, fenvalerate, and permethrin are wide spectrum insecticides. Chen et al. (2003) found that bifenthrin significantly decreased the number of and parasitism by D. isaea. Huang et al. (1999) reported that fenpropathrin’s dry residue caused 100% mortality of N. okazakii in 12 h. Mason and Johnson (1988) reported that D. begini was significantly more tolerant to fenvalerate and permethrin than C. punctiventris or G. utilis, and H. circulus and G. utilis had the lowest tolerances to permethrin. Spinosad. Spinosad is a widely used insecticide for controlling numerous pest insects, including Liriomyza. Hossain and Poehling (2006) found that spinosad is harmful to O. chromatomyiae and N. formosa, and that topical application of spinosad affected the development of O. chromatomyiae, and strongly affected N. formosa adult emergence when applied at different immature stages of N. formosa. Triazines. Two triazin insecticides, cyromazine and pymetrozine, are insect growth regulators, and have been widely used for management of Liriomyza. Studies show that cyromazine is compatible with H. varicornis and D. isaea (Chen et al. 2003; Prijono et al. 2004; Bjorksten and Robinson 2005). Application of cyromazine though drip irrigation systems had minor effects on D. isaea compared with spray which reduced the parasitoid populations (Weintraub 1999). Bjorksten and Robinson (2005) found that cyromazine did not cause significant mortality to larvae and pupae of H. varicornis and D.isaea, and progeny production and longevity of H. varicornis were not affected by adult exposure to cyromazine, nor did direct pupal exposure. However, Trumble (1985) found that cyromazine reduced parasitism on tomato. Tran et al. (2005) found that the glassfilm of pymetrozine was harmful to N. formosa with reduced survival and longevity. Fungicides. Mancozeb is one of the most commonly used fungicides. It has been found no negative effects on larvae and pupae of H. varicornis and D.isaea, progeny production and longevity of H. varicornis by adult and pupal exposures and leaf residence time on parasitism by H. varicornis (Bjorksten and Robinson 2005; Prijono et al. 2004). Insecticide application methods. The insecticide application can affect the effects on target pests as well as the parasitoids. Weintraub (1999) found that application of cyromazine by drip irrigation systems had minor effects on D. isaea compared with spray which significantly reduced the parasitoid populations. Parasitoids from heavily sprayed crops exhibit higher tolerance to insecticides. Rathman et al. (1990, 1995) found that D. begini from an untreated population that had been treated frequently with insecticides were 10- and 29-fold more resistant to several pyrethroids than a population with an unknown spray history.

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22.12


Conclusion

At present, a wide range of insecticides is used to manage these species but insecticide resistant populations are fast developing and this has added a new problem for farmers. Many newer chemical insecticides or biopesticides have no measurable impact on non-target organisms and should enhance the impact of natural enemies, leading to the development of truly integrated pest management. In spite of the fact that biological control of Liriomyza has been restricted for use in greenhouses or other protected environments, with the current availability of natural enemies, the possibility of building up a biologically based IPM scheme exists. More effort should be made to understand, conserve and enhance local natural enemies before the introduction of exotic parasitoids is considered. In particular, gaps should be identified in local parasitoid guilds such that ecologically compatible exotic agents can be identified. The final point should be that continued excessive pesticide usage may be a larger threat to local biodiversity than importing non-specialist leafminer parasitoids. If so, we can expect to see a reduction in leafminer insecticide treatments that goes beyond the area treated with beneficial arthropods.

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CHAPTER 23

Development of an Integrated Greenhouse Whitefly Management Program on Strawberries Jianlong Bi

Abstract California is a world leader in strawberry production. The greenhouse whitefly, Trialeurodes vaporariorum (Westwood), has recently been emerged as a major pest on strawberries in California. This insect ingests plant phloem fluid to cause yield and quality reduction while producing honeydew and associated sooty mold to decrease the market value of fruits. In addition, this insect transmits plant virus diseases. This chapter focuses on the development of an integrated greenhouse whitefly management program on strawberries largely based on the author’s research for over a decade. Keywords Strawberry, whitefly, integrated pest management

23.1

Introduction

Strawberries (Fragaria x ananassa Duch) are considered as one of the most delicious and nutritious fruits in the world. The United States is a world leader in strawberry production where California accounts for near 90% of the production, with 65–75% for fresh markets and the remainder processed. California grows over 20% of the world’s strawberry supply. In 2008, the total production acreage was 14,780 ha with a value exceeding $2.3 billion (PSABC 2008). There are five strawberry growing areas in California: Central Coast, Santa Maria, Oxnard Plain, South Coast and Central Valley (Fig. 23.1). About two-thirds of the total acreage is planted in the Central Coast and Santa Maria Valley and one-third is planted in the Oxnard Plain. The high productivity of strawberry fields is attributed to the high yield potential of the cultivars, mild coastal climates, intensive management of the crop, and effective pest management techniques (UC IMP 2008). The strawberry production uses clean nursery seedlings planted into fumigated soil. Jianlong Bi University of California Cooperative Extension, Salinas, CA 93901, USA E-mail: [email protected]


Development of an Integrated Greenhouse Whitefly Management Program on Strawberries 405

Fig. 23.1 Strawberry growing areas in California. Shaded areas are regions for strawberry production (UC IPM 2008).

The greenhouse whitefly, Trialeurodes vaporariorum (Westwood), is an important insect pest of horticultural crops in greenhouses and an increasing pest problem on outdoor crops such as strawberry, pepper (Capsicum annuum L.), tomato (Lycopersicon esculentum Mill), lima bean (Phaseolus lunatus L.), cucumber (Cucumis sativus L.), raspberry (Rubusarcticus L.), lettuce (Lactucasativa L.), citrus (Citrus spp), celery (Apium graveolens var dulce L.) and cut flowers. It causes economic damage to crops by ingestion of plant sap, contamination of crop products with honeydew which forms a substrate for the development of sooty molds and transmission of plant virus diseases (Omar et al. 1992; Johnson et al. 1992; Liu et al. 1993). Since 1998, strawberry crop has been increasingly under attack from the greenhouse whitefly, especially in the production area of Oxnard Plain. Prevention of economic losses requires development of an integrated greenhouse whitefly management program. This will require a thorough understanding of the developmental biology and population ecology, accurate assessments of economic thresholds and a detailed knowledge of the most effective means of control.

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23.2

Ecologically based management

The Oxnard Plain is in a major agricultural area with high crop diversity throughout the year. Major favorable alternative host plants in the area for the greenhouse whitefly include lima bean, pepper, tomato and cucumber. Lima bean was grown from early May until late August, pepper from June until October, tomato from June until November, and cucumber from May until late October. Strawberry production in this region follows two crop regimes. The majority of fields (about 70%) are planted in September or October (Fall) and terminated in the following June or July. A growing minority of fields (about 30%) are planted in late July or August (Summer) and terminated in December. We investigated the preference and developmental duration of the whitefly on strawberry, pepper, tomato, lima bean and cucumber in growth chamber studies (Fig. 23.2 and Fig. 23.3). The most preferred host for the adult whiteflies was cucumber, followed by tomato, strawberry, lima bean and pepper. Developmental time for eggs on the 5 host-plants species ranged from 9.6 to 10.1 days. Developmental duration for the first instar crawler was 7.4 days on strawberry, 6.1 days on cucumber, 6.2 days on lima bean, 6.6 days on pepper and 5.7 days on tomato. Developmental duration for the second instar was 5.4 days on pepper, 4.9 days on tomato, 4.8 days on strawberry, 3.6 days on cucumber and 3.3 days on lima bean. Developmental duration for the third instar was 6.4 days on strawberry, a 36%, 56% and 31% longer than these on cucumber, pepper and tomato, respectively, whereas the developmental time on cucumber, lima bean, pepper and tomato was similar. The fourth instar developmental time on strawberry, cucumber, lima bean and tomato ranged from 7.5 to 8.7 days, which were longer than that on pepper (5.4 days). The total developmental time from egg to adults was 35.3, 32.7, 31.8, 31.6 and 31.4 days on strawberry, cucumber, lima bean, tomato and pepper, respectively.

Fig. 23.2 Greenhouse whitefly host selection among strawberry, cucumber, lima bean, pepper and tomato.

Population ecology of the greenhouse whitefly was investigated in the Oxnard


Fig. 23.3 Greenhouse whitefly developmental duration on strawberry, cucumber, lima bean, pepper and tomato.

Plain by monitoring the seasonal population dynamics of the whitefly from April 2000 to April 2001 in commercial fields of strawberries, lima bean, pepper and tomato (Bi et al. 2002a). Result showed that whiteflies were most abundant during April and May on fall-planted strawberries and when the season of fallplanted strawberries was near over these whiteflies migrated to lima bean and tomato (Fig. 23.4). When the tomato season was close to end, they shifted their host to summer-planted strawberries and then returned to the fall-planted strawberries. For an effective ecologically-based management, this seasonally overlapping host circle must be broken. Growers are advised to avoid growing these favorable host species in proximity. Also, crop sanitation after the harvest is necessary.

Fig. 23.4

Greenhouse whitefly population dynamics in the Oxnard Plain.

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23.3

Jianlong Bi

Biological control

Encarsia Formosa (Gahan) is used world wide to control the greenhouse whitefly on various greenhouse crops by direct feeding and ovipositing in the whitefly nymphs (Hoddle et al. 1998). E. formosa prefers to feed on 2nd instar nymphs and pupae and to oviposit in the 3rd and 4th instar nymphs (Nell et al. 1976; Alphen et al. 1976; Nechols and Tauber 1977). The commercial use of E. formosa in Europe started in the 1920s. After World War II, the use of E. formosa declined due to the development of new insecticides which provided control on most greenhouse crops. However, resistance to insecticides was observed and interest in this biological control agent has increased since 1970. In 1983, the use of E. formosa to control whiteflies was reported from 15 countries (Van Lenteren 1983) and in 1999 E. formosa was used on 4800 hectares of greenhouse crops (Van Lenteren 1995). The potential of using parasitoids to control the greenhouse whitefly on outdoor crops was also investigated. Udayagiri and Bigelow (2000) released 30,000 E. formosa per acre in strawberry fields in the central coast area of California in January and February of 2000. They subsequently observed near 50% reduction in adult whitefly population, 53% parasitism of whitefly pupae and the parasitoid was recovered at all the releasing sites (Udayagiri and Bigelow 2000). In the Oxnard Plain, Philips et al. (1999) inundatively released 36,000 Eretmocerus emericus (Rose and Zolnerowich) per acre in commercial strawberry fields in April of 1999 (Philips et al. 1999). Their results indicated that E. emericus failed to become established and instead native Encarsia was found in the fields. We have also found that natural parasitism or predation in strawberry fields in the Oxnard Plain is very low. This may be largely caused by heavy pesticide applications in this intensive agricultural production area. These evidences suggest that using parasites and/or predators to control the whitefly in this area may not be feasible.

23.4

Chemical control

Insecticides are important components of integrated pest management systems designed to suppress whitefly populations on various crops (Liu et al. 1993; Toscano et al. 1998). Intensive research has been carried out in recent years for evaluating insecticides with novel modes of action against whiteflies. Imidacloprid and thiamethoxam are chloronicotinyl insecticides that act as nicotinic acetylcholine receptor agonists and are used to control homopteran pests (Leicht 1993; Elbert et al. 1998). Buprofezin is an insect growth regulator which inhibits chitin synthesis in several homopteran pests including whiteflies (De Cock et al. 1990). Pyriproxyfen is a juvenile hormone mimic affecting the hormonal balance in insects and resulting in strong suppression of embryogenesis and adult formation (Ishaaya and Horowitz 1994). Pyridaben is a mitochondrial electron transport inhibitor insecticide and acaricide (Denholm et al. 1998). Pymetrozine is a new insecticide which inhibits feeding behavior, leading to


mortality due to starvation or desiccation (Harrewijn and Kayser 1997; Fuog et al. 1998). It is highly active and specific against sucking insect pests such as aphids and whiteflies (Fuog et al. 1998). The separate modes of action of these compounds, together with their selectivity against targeted insect pests and relative safety to beneficial insects and other organisms, present an exciting opportunity for their effective integration into pest management strategies (Darvas and Polgar 1998; Ishaaya and Horowitz 1998). The availability of such chemical diversity enables the development of a management strategy which minimizes the threat of insecticide resistance (Denholm et al. 1998). In 2000, we evaluated the efficacy of these novel insecticides against the greenhouse whitefly on strawberries in both greenhouse and field experiments (Bi et al. 2002b). In greenhouse studies, 10, 20 and 40 mg systemic applications of imidacloprid per plant caused adult mortalities of 82%, 93% and 96%, respectively, at 48 h post-treatment. Residual activity was significant and resulted in 84 to 87% mortality after 40 days and 58 to 78% mortality after 60 days when insects were exposed to treated plants. Topical applications of pyriproxyfen at rates of 16.2, 32.4 and 64.8 (top label rate) mg AI/ml completely inhibited egg hatch, whereas rates of 0.05, 0.10 and 0.20 mg AI/ml resulted in 38% to 70% egg hatch. However, 84% to 94% of these hatched nymphs failed to develop into adults. Applications of buprofezin and pyridaben at 0.5, 1.0 and 2X their greatest label concentrations caused 100% mortality of nymphs whereas pymetrozine at 0.5, 1.0 and 2.0X its greatest label concentration only caused mortalities of 70%, 74% and 78%, respectively. Soil formulated thiamethoxam at 0.5 and 1.0X its greatest label concentration killed 67% and 90% of nymphs, respectively. In field experiments with fall-planted strawberries in the mid season (7 months after planting) in the Oxnard Plain, soil formulated imidacloprid applications decreased adult densities by 31% to 61% from 21 to 42 days post treatment compared with controls, and suppressed densities of third and fourth instars by 34% to 68% after 42 days of treatment. Pyriproxyfen and buprofezin had no significant effect on adult densities on most sampling dates. An application of buprofezin (on 20 April) followed by an application of pyriproxifen (on 4 May) decreased nymphal densities by 98% on the 4 and 11 May sampling dates and maintained 61%–85% control after the 1 June sampling dates. Applications of pyriproxyfen (20 April) followed by buprofezin (on 4 May) had no effect on the nymphal densities on 4 and 11 May but densities had fallen by 78%–90% from early to late June. Applications of pyridaben and pymetrozine had no effect on adult densities on any sampling dates. However, pyridaben suppressed densities of third and fourth instars by 39%–44% on 4 and 11 May sampling dates, and by 47%–73% in late June. Pymetrozine had no effect on the densities of nymphs. Applications of thiamethoxam had no effect on adult densities on most sampling dates and did not significantly change the immature densities compared to the control. The efficacy of imidacloprid, thiamethoxam, buprofezin and pyriproxifen against the greenhouse whitefly on summer-planted strawberries was also

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evaluated in field experiments in the Oxnard Plain (Be et al. 2002c). Imidacloprid applied in soil three weeks after planting decreased whitefly adult numbers by 58% to 90%, first and second instars by 78% to 93% up to 56 days post application, and third and fourth instars by 42% to 86% up to 77 days post application, whereas thiamethoxam applied similarly reduced adults by 58% to 80%, first and second instars by 78% to 93% up to 6 weeks post treatment, and third and fourth instars by 48% to 80% up to 10 weeks after initial application, compared to non-treated controls. Imidacloprid applied in soil immediately prior to planting further suppressed numbers of whiteflies by 71% to 83% (adults), 58% to 74% (first and second instars) and 52% to 74% (third and fourth instars), in comparison with the same compound applied through drip irrigation lines four weeks after planting. Buprofezin and pyriproxifen applied six weeks after planting reduced numbers of adult whiteflies by 25%–81% and 40%–73%, respectively, first and second instars by 61%–92% and 51%–100%, respectively, and third and fourth instars by 45%–100% and 37%–87%, respectively, on most sampling dates up to 7 weeks post application. These results supported the emergency registration of Admire (soil formulated imidacloprid) and Esteem (pyriproxifen) in 2003 on strawberries in California. Since then, almost all the fields infested with greenhouse whiteflies were treated with Admire.

23.5 Impact of the greenhouse whitefly management on strawberry fruit quality Strawberries are considered as one of the most delicious and nutritious fruits in the world. Strawberries contain soluble carbohydrates and organic acids, which are important dietary nutrients and make a significant contribution to the fruit flavor (Kader 1990; Cordenunsi et al. 2003). Soluble carbohydrates accumulate in fruits in the form of glucose, fructose and sucrose with hexose predominating (Woodward 1972; Ranwala et al. 1992; Hubbard et al. 1991). Soluble carbohydrate content is dependent upon genotype and prevailing climatic conditions (Humle 1970; Olsson et al. 2004). Strawberries also contain dietary antioxidants such as vitamin C and phenolic compounds (Wang et al. 1996; Heinonen et al. 1998; Wang et al. 2003). Antioxidants can scavenge reactive oxygen species or free radicals which have the potential to damage cell components such as lipids, nucleic acids and proteins, leading to cancer and other human diseases (Halliwell 1991; Halliwell and Aruomo 1992; Steinberg 1991). Strawberries have shown a high scavenging activity toward these radicals (Heinonen et al. 1998; Wang and Jiao 2000; Wang and Lin 2000). We tested the effects of whitefly management with insecticides on the quality of strawberry fruit (Bi et al. 2007). Applications of imidacloprid, thiamethoxam, buprofezin and pyriproxyfen decreased the mean adult whitefly numbers by 2.80-, 2.17-, 1.69- and 1.39-fold, respectively, compared to the untreated control. Similarly, the mean numbers of first and second instar whiteflies were reduced 4.36-, 2.20-, 1.90- and 2.02-fold, respectively, while the mean numbers of third


and fourth instars were reduced 5.48-, 2.28-, 2.71- and 1.43-fold, respectively, in plants treated with imidacloprid, thiamethoxam, buprofezin and pyriproxyfen. The mean soluble solids content in imidacloprid, thiamethoxam, buprofezin and pyriproxyfen treatments was 1.04-, 1.06-, 1.03- and 1.04-fold greater, respectively, than that in the control. The whitefly reduction enhanced the mean fruit titratable acidity by 4–6%. Mean glucose levels in imidacloprid and thiamethoxam treatments were significantly higher than in other treatments. However, the whitefly management did not affect the mean fructose levels. Imidacloprid, thiamethoxam and pyriproxyfen treatments boosted the ascorbic acid levels by up to 4%. It should be noted that the natural population of whiteflies involved in this study was relatively small (e.g. 8 adults per leaflet in the untreated plots). We observed a much greater whitefly population in some strawberry fields ( > 50 adults/leaflet) in southern California in 1999 and 2003 when the greenhouse whitefly was regionally more abundant (unpublished data). The higher numbers of whiteflies may further reduce the quality of strawberry fruits.

23.6

Combating insecticide resistance

Conventional insecticide classes such as chlorinated hydrocarbon, organophosphate, carbamate and pyrethroid are still being used for controlling several insect pests on various crops in Ventura/Oxnard area. At present, these insecticides are only recommended for limited use in rotation with neonicotinoids and/or insect growth regulators to control whiteflies (Liu and Meister 2001; Polumbo et al. 2001). On strawberries, foliar application of these insecticides is recommended to suppress the high whitefly populations in mid or late season when imidacloprid applied at planting or early season diminishes. Extensive reliance on chemical insecticides for whitefly control has resulted in whitefly resistance to almost all major classes of conventional insecticides throughout the world (Wardlow et al. 1972; Wardlow et al. 1975; Wardlow et al. 1976; Zou and Zheng 1988; Omer et al. 1992). Resistance monitoring can be an effective component of a resistance management program and detection of changes in resistance/susceptibility can facilitate use of alternate control measures (Prabhaker et al. 1992). We conducted studies to determine the status of the greenhouse whitefly susceptibility to neonicotinoids including imidacloprid, thiamethoxam, dinotefuran and acetamiprid, and commonly used conventional insecticides such as endosulfan (chlorinated hydrocarbon), chlorpyrifos and malathion (organophosphate), methomyl (carbamate), bifenthrin and fenpropathrin (pyrethroid) on strawberries in the Oxnard Plain (Bi and Toscano, 2007a). For bioassay tests, adult whiteflies were collected from commercial strawberry crop and immatures were directly developed from eggs laid by these adults (Bi and Toscano 2007a). LD50s of soil-applied imidacloprid, thiamethoxam and dinotefuran were 8.7-, 3.2- and 4.9-fold higher for the adults, 1.8-, 1.2- and

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1.5-fold higher for the first instar nymphs, and 89.4-, 390.3- and 10.4-fold higher for the third instar nymphs, respectively, than their top label rates. LC50s of foliarapplied imidacloprid, thiamethoxam and acetamiprid were 6.1-, 6.0-, 1.7-fold higher for the adults, 3.8-, 8.7-, and 4.4-fold higher for the second instar nymphs, respectively, than their top label rates. For the adults, LC90s of endosulfan, malathion, methomyl, bifenthrin, and fenpropathrin were 2.2-, 1.2-, 1.9-, 2.3- and 4.9-fold lower than their respective top label rates. Chlorpyrifos was not very effective against the adults as indicated by its LC90 being 120% higher than its top label rate. Our results strongly emphasize the need to develop resistance management strategies in the region. Introduction of novel insecticides with distinct modes of action into the current whitefly control program is a valuable tactic for resistance management (Denholm et al. 2002; Liu 2004). Spiromesifen is a novel insecticide and acaricide belonging to the new chemical class of spirocyclic phenyl-substituted tetronic acids (Nauen et al. 2002). This compound acts on interfering with insect/mite lipid biosynthesis (Nauen et al. 2002). Spiromesifen is especially active against whiteflies (Bemisia spp. and Trialeurodes spp.) and spider mites (Tetranychus spp.) in several cropping systems including cotton (Gossypium hirsutum L.), vegetables, and ornamentals (Nauen et al. 2002; Liu 2004; Polumbo 2004). We tested the efficacy of spiromesifen against the greenhouse whitefly on strawberries under both laboratory and field conditions (Bi and Toscano 2007b). Laboratory experiments showed that spiromesifen at 0.5 and 1.0 μg$mL–1 a.i. inhibited the egg hatching by 80% and 100%, respectively, while at concentrations of 3.1, 3.0, and 10.0 μg$mL–1 a.i., this insecticide respectively killed 100% of the first, second, and third instar nymphs (the label rate is 300 μg$mL–1 a.i.). Much lower toxicity to adults was observed. Field trials revealed that application of spiromesifen reduced the whitefly egg numbers by 61% to 80% from 2 to 3 weeks post-treatment in comparison with the pyriproxyfen treatment, whereas the application lowered the egg numbers by 34% to 73% from 2 to 5 weeks post-treatment compared to the buprofezin treatment. In comparison with pyriproxyfen treatment, spiromesifen application decreased the numbers of immature whiteflies by 29% to 92% from 1 to 6 weeks post-treatment. The effect of spiromesifen on reduction of immatures was similar to that of buprofezin. Also, the efficacy of spiromesifen on suppression of adult numbers was comparable to that of pyriproxyfen or buprofezin. Spiromesifen shows promise for inclusion in integrated greenhouse whitefly management programs and insecticide resistance management programs on strawberry. Acknowledgements The author wish to thank Dr. Nick Toscano (University of California Riverside) and the California Strawberry Commission for their support, Mr. G. R. Ballmer and Mr. X. F. Li for their assistance, and Drs. S. J. Castle, C. C. Chu, and Q. N. Cai for their critical review of an earlier version of the manuscript.


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Development of an Integrated Greenhouse Whitefly Management Program on Strawberries 415 storage. J. Agric. Food Chem., 2004, 52: 2490–2498. Omer A D, Leigh T F, Granett J. Insecticide resistance in field populations of greenhouse whitefly (Homoptera: Aleyrodidae) in the San Joaquin valley (California) cotton system. J. Econ. Entomol., 1992, 85: 21–27. Philips P A, Rogers J, Malone R. Biological control of greenhouse whitefly, Trialeurodes vaporariorum in Ventura county strawberries using Eretmocerus emericus. The Pink Sheet, 1999: 99. Polumbo J C. Comparative efficacy of Oberon (spiromesifen) against Bemisia whiteflies in spring cantaloupes. 2004 Vegetable Report, University of Arizona, college of Agriculture and Life Science (http://ag.arizona.edu/pubs/crops/az1348/az1348_2a.pdf). Polumbo J C, Horowitz A R, Prabhaker N. Insecticidal control and resistance management for Bemisia tabaci. Crop Prot., 2001, 20: 739–765. Prabhaker N, Toscano N C, Perring T M, et al. Resistance monitoring of the sweetpotato whitefly (Homoptera: Aleyrodidae) in the Imperial Valley of California. J. Econ. Entomol., 1992, 85: 1063–1068. Ranwala A P, Suematsu C, Masuda H. Soluble and wall-bound invertase in strawberry fruit. Plant Sci., 1992, 84: 59–64. Steinberg D. Antioxidants and atherosclerosis: a current assessment. Circulation, 1991, 84: 1420– 1425. Toscano N C, Prabhaker N, Zhou S, et al. Toxicity of Applaud and Knack against silverleaf whiteflies from southern California: Implications for susceptibility monitoring. // Dugger P and Richer D. Proceedings of Beltwide Cotton Research Conference. Memphis: National Cotton Council of America, 1998: 1093–1095. Udayajiri S, Bigelow R. Potential for biological control of the greenhouse whitefly in the field with Encarsia Formosa. // Hoddle M. California conference on Biological Control, 2000: 191– 194. Van Lanteren J C. Potential of entomophagous parasites for pest control. Agri. Econ. Environ., 1983, 10: 143–158. Van Lanteren J C. Integrated pest management in protected crops. // Dent D R. Integrated Pest Management: Principles and systems Developments, 1995, 12: 311–343. Wang S Y, Jiao H. Scavenging capacity of berry crops on superoxide radicals, hydrogen peroxide, hydroxyl radicals, and singlet oxygen. J. Agric. Food Chem., 2000, 48: 5677–5684. Wang S Y, Lin H S. Antioxidant activity in fruit and leaves of blackberry, raspberry, and strawberry is affected by cultivar and maturity. J. Agric. Food Chem., 2000, 48: 140–146. Wang H, Cao G, Prior R L. Total antioxidant capacity of fruits. J. Agric. Food Chem., 1996, 44: 701– 705. Wang S Y, Bunce J A, Maas J L. Elevated carbondioxide increases contents of antioxidant compounds in field-grown strawberries. J. Agric. Food Chem., 2003, 51: 4315–4320. Wardlow L R, Ludlam F A B, French N. Insecticide resistance in glasshouse whitefly. Nature, 1972, 239:164–165. Wardlow L R, Ludlam F A B, Hammon R P. A comparison of the effectiveness of insecticides against greenhouse whitefly (Trialeurodes vaporariorum). Ann. Appl. Biol., 1975, 81: 433–435. Wardlow L R, Ludlam F A B, Bradley L F. Pesticide resistance in glasshouse whitefly (Trialeurodes vaporariorum Westwood). Pestic. Sci., 1976, 7: 320–324. Woodward J R. Physical and chemical changes in developing strawberry fruits. J. Sci. Food Agric., 1972, 23: 465–473.

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Zou Y Q, Zheng B Z. The toxicity of some insecticides to greenhouse whitefly (Trialeurodes vaporariorum Westwood) and monitoring of resistance. Acta Phytophyl Sinica, 1998, 15: 277– 281.

CHAPTER 24

Advances in Research on the Venom Chemistry of Imported Fire Ants Jian Chen and Hanwu Shang

Abstract Workers of the imported fire ants,including red imported fire ants, Solenopsis invicta Buren, black imported fire ants, S. richteri Forel, and their hybrid (S. invicta S. richteri), are vicious stingers. Since the venomous sting is a significant medical problem to humans, the chemistry of imported fire ant venom has been a subject of rigorous research. This chapter summarizes the recent research on the chemistry of imported fire ant venom and focuses on several newly identified piperideine alkaloids. The potential pharmacologic application of piperidine alkaloids and updates on major allergens found in the imported fire ant venom are also reviewed. Keywords imported fire ants, venom, piperidine alkaloids, allergens, pharmacologic application

24.1

Introduction

Imported fire ants, including red imported fire ant, Solenopsis invicta Buren, black imported fire ant, S. richteri Forel, and their hybrid (S. invicta S. richteri) are significant pests in the United States, particularly the S. invicta. Both S. invicta and S. richteri were introduced into the United States from South America (Buren et al. 1972). S. invicta has been found in 13 states (Mobley and Redding 2005); however, S. richteri is confined to a very limited area in northern Mississippi, Alabama and Tennessee (Buren et al. 1974; Vander Meer et al. 1985; Oliver et al. 2009). The hybrid imported fire ant (S. invicta S. richteri) was found in Jian Chen Biological Control of Pests Research Unit, Agricultural Research Service, U.S. Department of Agriculture, Stoneville, MS, 38776, USA E-mail: [email protected] Hanwu Shang College of Life Sciences, China Jiliang University, Hangzhou, Zhejiang, 310018, China


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Mississippi (Menzel et al. 2008; Streett et al. 2006), Alabama (Diffie et al. 1988), Georgia (Gardner et al. 2008; Diffie et al. 1988) and Tennessee (Oliver et al. 2009; Gibbons and Simberloff 2005). S. invicta has also invaded Australia, New Zealand, Bahamas, British and U.S. Virgin Islands, Cayman Islands, mainland China, Hong Kong, Malaysia, Singapore, Taiwan, Trinidad and Tobago, Turks and Caicos Islands and Mexico (ISSG 2009; Sánchez-Peña et al. 2005). In agriculture, fire ants damage 57 species of cultivated plants (Adams 1986). They feed on various crop seeds (Morrison et al. 1997) and can cause loss in soybeans (Adams et al. 1993). Vinson (1997) has reviewed the biology, behavior, and impacts of imported fire ants in North America. It is believed that among the most important negative results of the presence of imported fire ants in North America is their ecological impact (Wojcik et al. 2001). After only 70 years of introduction, S. invicta has even altered morphologies of native lizards in the infested area (Langkilde 2009). Undoubtedly, the venom plays a critical role in the success of imported fire ants as invasive species. It is also the venom that makes the imported fire ants such a significant health hazard to humans. Approximately 30% of the human population in infested areas is stung each year and medical care is required for victims who develop hypersensitivity. Today, in the fire ant infested region, fire ant hypersensitivity has become the predominant cause of prescribed immunotherapy for hymenopterous allergy (Freeman 1997). The venom causes a burning sensation at the site of the sting. Soon after the attack, the site of the sting begins to itch. At the same time, a wheal-and-flare reaction develops and then subsides within 2 h. A sterile pustule may develop at the sting site over the next 12–24 h (Caro et al. 1957). In addition to local dermal reactions, the venom of imported fire ants is a major cause of serious anaphylactic reactions in the southern United States and other distributed areas (Hoffman 1995). Left untreated, anaphylactic shock can lead to death within minutes. The sting apparatus of the imported fire ants is a modified ovipositor. There are two glands that are associated with the sting apparatus: poison gland and Dufour’s gland (Callahan et al. 1959). The venom of imported fire ants is produced in the poison gland and stored in a venom sac that is attached to the stinger. Venom production is age dependent (Deslippe and Gao 2000; Haight and Tschinkel 2003). Venom synthesis is limited to early life, and the amount of venom injected is modulated (Haight and Tschinkel 2003). The highest rate of synthesis is in workers aged 1 day after adult eclosion (1.17 μg venom/day for workers with head widths of 1 mm). The rate of synthesis then declines by 75% at age of 15 days (0.30 μg venom/day). Amount of venom delivered per sting is also age dependent. Mid-age worker deliver more venom than older workers per sting. Venom production is also season dependent. Amount of venom in nest-defenders is 55% higher in the spring than in the rest of the year (Haight and Tschinkel 2003). In the act of stinging, fire ant workers anchor their bodies by grasping the skin with their mandibles, and then insert the stinger into the skin, and release the

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venom. Multiple stings can be delivered around a single grasping point, resulting in a circular pattern of stings. It is believed that many workers can sting at the same time; however, how ants achieve such synchronization is unknown. In addition to direct injection, venom is used in many other ways. Workers disperse venom through the air using gaster flagging behavior to repel heterospecifics encountered in the foraging arena or to dispense to the brood surface as an antibiotic (Obin and Vander Meer 1985). Venom alkaloids were also found in red imported fire ant nest material (Chen 2007). In contrast to the venoms of other insects in Hymenoptera, such as bees, hornets, and wasps, which are usually aqueous solutions containing proteins, 95 % of red imported fire ant venom consists of alkaloids with only a small fraction of proteins (0.1%). The alkaloid component of fire ant venom contains predominately 2-methyl-6-alkyl or alkenylpiperidines (Brand et al. 1972). One imported fire ant can deliver about 0.11 μl of venom per sting and they can deliver venom in 20 consecutive stings before depleting their venom storage (deShazo and Soto-Aguillar 1993). Each sting contains 10–100 ng water-soluble proteins (Stafford et al. 1989). The chemical structures of the piperidine alkaloids have been well defined (MacConnell et al. 1970, 1971, 1974; Brand et al. 1972; Jones et al. 1982; Blum et al. 1992; Leclercq et al. 1994). The proteins found in imported fire ant venom have also been well studied (Hoffman et al. 2008, 2005, 1990, 1988; Hoffman 1997, 1995, 1993; Padavattan et al. 2008; Park et al. 2008; Schmidt et al. 2003, 2001, 1996, 1993). Fire ant venom has been reviewed by several authors (Jones et al. 1982; Fitzgerald and Flood 2006; Hoffman 1995; Tschinkel 2006). This chapter summarizes the recent research on the chemistry of imported fire ant venom, focusing on several newly identified piperideine alkaloids. The potential pharmacologic application of piperidine alkaloids, and updates on major allergens found in the imported fire ant venom have also been reviewed.

24.2 New piperideine alkaloids found in venom of imported fire ants After isolating the extracts of whole worker bodies with silica gel column chromatography, several piperideine alkaloids were found in the imported fire ants, including seven Δ1,6-piperideines (Chen et al. 2009; Chen and Fadamiro 2009a, 2009b) and six Δ1,2-piperideines in S. invicta and S. richteri (Chen and Fadamiro 2009a, 2009b) (Table 24.1). Six Δ1,6-piperideines were purified from the extracts of poison glands and worker whole bodies of S. invicta (Chen et al. 2009) (Figure 24.1). Mass spectrum of each Δ1,6-piperideine had significant ions at m/z 96, 111, and 124. Three parent ions [M + H]+ were detected at m/z 306.3131, 308.3279, and 334.3440 respectively in the high-resolution MS analysis (HRMS). HRMS analysis showed that the molecular formula for compounds 3, 4 and 5 (Fig. 24.1) were C21H39N, C21H41N and C23H43N, respectively. Reduction of those samples using NaBH4 in ethanol generated

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Table 24.1 Piperideine alkaloids in imported fire ants (R: S. invicta, B: S. richteri, H: S. invicta S. richteri). Double bond position on the ring 1,6

Δ

Δ1,2

Chemical name

Ants R B H

2-methyl-6-undecyl-6-piperideine

Y Y Y

2-methyl-6-(cis-4-tridecenyl)-6-piperideine

Y Y Y

2-methyl-6-tridecyl-6-piperideine

Y Y Y

2-methyl-6-(cis-6-pentadecenyl)-6-piperideine

Y Y Y

2-methyl-6-pentadecyl-6-piperideine

Y Y Y

2-methyl-6-(cis-8-heptadecenyl)-6-piperideine

Y ─ Y

2-methyl-6-heptadecenyl-6-piperideine

Y ─ Y

2-methyl-6-undecyl-2-piperideine

Y Y u

2-methyl-6-(cis-4-tridecenyl)-2-piperideine

Y Y u

2-methyl-6-tridecyl-2-piperideine

Y Y u

2-methyl-6-(cis-6-pentadecenyl)-2-piperideine

Y Y u

2-methyl-6-pentadecyl-2-piperideine

Y ─

u

2-methyl-6-(cis-8-heptadecenyl)-2-piperideine

Y ─

u

Y: found; ─: not found; u: undetermined.

piperidine alkaloids found in the fire ant poison gland, resulting in both the cis and trans piperidine alkaloids (Fig. 24.2), indicating they are structurally related to piperidine alkaloids of the red imported fire ants. Due to numerous investigations on the chemistry of fire ant piperidine alkaloids, chemical structures of piperidine alkaloids in fire ant venom and their GC behavior were well defined (MacConnell et al. 1971; Jones et al. 1982). This work provided an excellent opportunity to aid in identifying these new piperideine alkaloids. The identification of piperidine alkaloids was achieved by comparing their retention and mass spectra to those of purified piperidine alkaloids from fire ant extracts or synthetic standards. Based on the information from mass spectra, GC-retention times, molecular weights obtained from high resolution mass analysis, and profiles of NaBH4 reduction products, these piperideine alkaloids were identified as 2-methyl-6-tridecenyl-6-piperideine, 2-methyl-6-tridecyl-6-piperideine, 2methyl-6-pentadecenyl-6-piperideine, 2-methyl-6-pentadecyl-6-piperideine, 2methyl-6-heptadecenyl-6-piperideine, and 2-methyl-6-heptadecyl-6-piperideine. Identification of Δ1,2-piperideines in S. invicta and S. richteri were solely based on conventional mass spectra (Chen and Fadamiro 2009a, 2009b). Since chemical profiles of venom alkaloids are correlated to the genetic makeup of imported fire ant species, they have been used as a taxonomic tool to separate S. invicta, S. richteri and their hybrid (Ross et al. 1987). Would the piperideine profile be helpful in separating fire ant species? Chen and Shang (unpublished data) have looked into this possibility based on the profile of Δ1,6piperideines. In S. richteri, 2-methyl-6-heptadecenyl-6-piperideine and 2-methyl-


421

Fig. 24.1 GC-MS-EI total ion chromatograms (TIC) of hexane extract of fire ant poison glands (A) and worker whole bodies (B) after column chromatography and peak assignment.

6-heptadecyl-6-piperideine were not detected in all samples, whereas, 2-methyl6-tridecenyl-6-piperideine, 2-methyl-6-tridecyl-6-piperideine, 2-methyl-6-pentadecenyl-6-piperideine, and 2-methyl-6-pentadecyl-6-piperideine were present in all samples. In the hybrid, 2-methyl-6-heptadecenyl-6-piperideine and 2-methyl6-heptadecyl-6-piperideine was detected in 2 of 9 samples, while 2-methyl-6tridecenyl-6-piperideine, 2-methyl-6-tridecyl-6-piperideine, 2-methyl-6-pentadecenyl-6-piperideine, and 2-methyl-6-pentadecyl-6-piperideine were detected in all samples (Fig. 24.3). In S. invicta, the peak of 2-methyl-6-pentadecyl-6piperideine or 2-methyl-6-pentadecenyl-6-piperideine was the greatest. The peak of 2-methyl-6-heptadecenyl-6-piperideine was always more intensive than

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Fig. 24.2 Typical chromatograms of the NaBH4 reduction products of piperideine alkaloids (A), piperidine alkaloids (B) and peak assignment.

2-methyl-6-heptadecyl-6-piperideine. In S. richteri and the hybrid, the peak of 2methyl-6-pentadecenyl-6-piperideine was always more intensive than 2-methyl-6pentadecyl-6-piperideine (Fig. 24.3). The means of the ratio between 2-methyl-6pentadecenyl-6-piperideine and 2-methyl-6-pentadecyl-6-piperideine were significantly different among two species and their hybrid (F = 67.44, df = 2.34, P


423

Fig. 24.3 GC-MS-EI total ion chromatograms (TIC) of hexane extracts of fire ant worker whole bodies after column chromatography (See figure 1 for peak assignment).

< 0.0001; Table 24.2). Although the difference of the means among species and hybrid was obvious, some data points were very close. For example, the maximum ratio for the hybrid was 2.61 and the minimum for S. richteri was 2.69. It is easy to separate S. invicta from S. richteri, since only S. invicta has 2-methyl6-heptadecenyl-6-piperideine and 2-methyl-6-heptadecyl-6-piperideine; however, the more challenging is to separate S. richteri from the hybrid, because they can sometime have a very similar Δ1,6-piperideine profile. The ratio of peak area of 2-methy-6-pentadecenyl-6-piperideine to that of 2methy-6-pentadecenylpiperidine was 0.0150.002 (meanSE, n = 6), indicating that these piperideine alkaloids are definitely minor components in the fire ant

424


Table 24.2 ideine.

Ratio of 2-methyl-6-pentadecenyl-6-piperideine to 2-methyl-6-pentadecyl-6-piper-

Species

Mean (SE)

Min

Max

S. invicta

0.69 (0.063) a

0.39

0.92

S. richteri

4.00 (0.22) c

2.69

9.96

S. invicta S. richteri

1.85 (0.16) b

1.14

2.61

Means followed by different letter are significantly different.

venom. However, from a biological standpoint, minor compounds do not necessarily have lesser functions. Although nothing is known about the function of these piperideine alkaloids in fire ants, result of preliminary tests on Pseudacteon phorid flies, the biological control agents of imported fire ants, is interesting. Chen and Fadamiro (2009b) reported that Pseudacteon tricuspis antennae responded to Δ1,6- piperideines, in addition to cis and trans piperidine alkaloids. Phorid flies may use these compounds to locate their hosts. This is an hypothesis that should be tested. Leclerq and colleagues proposed that reduction of piperideine to piperidine is the final step of biosynthesis of piperidine alkaloids (Leclercq et al. 1996). Discovery of piperideines in the venom may serve as evidence to support this hypothesis. Piperideine alkaloids have been found in other Solenopsis ants and in plants. For example, 2-methyl-6-undecyl-2-piperideine from S. xyloni (Brand et al. 1972); 2-(4-penten-1-yl)-piperideine from a Puerto Rican ant species (Jones et al. 1982); 2-methyl-6-(2-oxopropyl)-1,2-piperideine, 2-methyl-6-(2-hydroxypropyl)-1,2-piperideine, and 2-methyl-6-(1-propenyl)-1, 6-piperideine from several Pinus species (Gerson et al. 2004); and γ-coniceine from the poison Hemlock (Conium maculatum L.) (Reynolds 2005) and several Aloe species (Dring et al. 1984; Nash et al. 1992).

24.3

Piperidine alkaloids

Piperidine alkaloids, major components of imported fire ant venom, have been a subject of numerous investigations (see Tschinkel 2006 for a review). Their chemical structures have been well defined. All piperidines found in imported fire ant venom are 2-methyl-6-alkyl or alkenylpiperidines. Both trans- and cispiperidine ring configurational isomers exist for each piperidine and the trans isomer is the predominant one. The alkyl or alkenyl side chains have 11, 13, 15, and 17 carbons. If the side chain is an alkenyl group, there is a double bond on the chain and the double bond isomer was believed to be all cis until Chen and Fadamiro (2009b) reported trans isomers for trans-2-methyl-6-tridecenylpiperidine and trans-2-methyl-6-pentadecenylpiperidine in S. invicta. Two piperidines with a double bond at the end of the side chain, were found in S. invicta (Chen and


425

Fadamiro 2009b), including trans-2-methyl-6-(12-tridecenyl)piperidine and trans-2-methyl-6-(14-pentadecenyl)piperidine. It was found that the absolute configuration of the trans alkaloids in the fire ants is always (2R, 6R) while that of the cis alkaloids is (2R, 6S) (Leclercq et al. 1994). Piperidine alkaloids are summarized in Table 24.3. Table 24.3 2-methyl-6-alkyl or alkenylpiperidine in imported fire ants (R: S. invicta, B: S. richteri, R B: S. invicta S. richteri). Chemical name

Isomer of ring configuration

Ants R B H

2-methyl-6-nonylpiperidine

trans

─ Y

u

cis

─

u

2-methyl-6-(cis-2-undecenyl)piperidine

trans

─ Y

u

cis

─

u

trans

Y Y Y

2-methyl-6-undecylpiperidine

─ ─

cis

Y Y Y

2-methyl-6-(cis-4-tridecenyl)piperidine

trans

Y Y Y

cis

Y Y Y

2-methyl-6-(trans-4-tridecenyl)piperidine

trans

Y ─

cis

─

─

u

trans

Y ─

u

cis

─

u

trans

Y Y Y

2-methyl-6-(12-tridecenyl)piperidine 2-methyl-6-tridecylpiperidine

─

u

cis

Y Y Y

2-methyl-6-(cis-6-pentadecenyl)piperidine

trans

Y Y Y

cis

Y Y Y

2-methyl-6-(trans-6-pentadecenyl)piperidine

trans

Y ─

u

cis

─

─

u

trans

Y ─

u

cis

─

u

trans

Y Y Y

2-methyl-6-(14-pentadecenyl)piperidine 2-methyl-6-pentadecylpiperidine 2-methyl-6-(cis-8-heptadecenyl)piperidine 2-methyl-6-n-heptadecenylpiperidine

─

cis

Y Y Y

trans

Y ─ Y

cis

Y ─ Y

trans

Y ─ Y

cis

Y ─ Y

Y: found; ─: not found; u: undetermined.

The absence of trans 2-methyl-6-(8-pentadecenyl)piperidine has been considered a character of S. richteri (Ross et al. 1987). However, Chen and Fadamiro (2009a) reported both trans and cis 2-methyl-6-(8-pentadecenyl)

426


piperidine and 2-methyl-6-pentadecylpiperidine in S. richteri. It may indicate that there is possible variation among samples regarding the content of this piperidine alkaloid. Chen and Fadamiro (2009a) also found 2-methyl-6-nonylpiperidine in S. richteri workers. Both cis and trans 2-methyl-6-nonylpiperidine were found in the venom of female alates of S. richteri (MacConnell et al. 1974); however, cis2-methyl-6-nonylpiperidine had never been reported in the venom of S. richteri workers. Alumina column chromatography can be used to separate cis and trans of synthetic piperidine alkaloids (MacConnel et al. 1971). Chen and Fadamiro (2009a, 2009b) used silica gel column chromatography to separate cis and trans piperidine alkaloids. They found that the cis isomers were always eluted before their corresponding trans isomers. Unfortunately, the separation protocol was not reported. Leclerq et al. (1996) proposed the first pathway for biosynthesis of fire ant venom alkaloids. The authors hypothesized that fire ant piperidine alkaloids were formed by the linear combination of 9, 10, or 11 acetate units. Loss of the carboxyl group from the resulting long-chain acid, followed by addition of an amino group, intramolecular cyclization, and reduction of the imino group could produce the cis- and trans-piperidine alkaloids. The intermediate long-chain acid can be either a suitably functionalized fatty acid or a polyketo acid. This biosynthetic pathway would therefore be similar to the well-established biosynthesis of the hemlock alkaloid coniine, which is formed by the linear combination of four acetate units (Leete 1963). Chen and Fadamiro (2009a) have also proposed a biosynthetic pathway for cis- and trans-piperidine alkaloids, which provides more details on the step in which Δ1,2 and Δ1,6-piperideines are reduced to piperidines. They hypothesized that an enantioselective enzyme reduced Δ1,2-piperideines exclusively into trans- piperidines (2R, 6R), whereas, it reduced Δ1,6-piperideines to both trans (2R, 6R) and cis (2R, 6S). Imported fire ant venom is toxic to many biological systems (see Tschinkel 2006; Fitzgerald et al. 2006 for reviews). They are allergenic, bactericidal, cytotoxic, fungicidal, hemolytic, herbicidal, insecticidal, necrotoxic and repellent (Tschinkel 2006). Recently, Greenberg et al. (2008) used natural venom collected from S. invicta to determine the LD50 of venom on Argentine ants, and several other ant species. They found Argentine ants were most susceptible to fire ant venom. Apparently, it is the venom that makes imported fire ants such a great threat to public health. However, several recent publications indicate that venom alkaloids of imported fire ants may have potential medical applications. Decentralized organisms such as bacteria use quorum sensing (QS) as a type of decision-making process. In that process, bacteria release signaling chemicals into the environment. They measure the concentration of the signaling chemicals within a population and change behavior and physiology in response to the concentration of the signaling chemicals. Many species of bacteria use quorum sensing to coordinate their gene expression, in response to the local density of


427

their population. In Pseudomonas aeruginosa, a common bacterium which can cause disease in animals and humans, quorum-sensing signaling regulates the expression of virulence factors and thus represents an attractive new target for anti-infective therapy. Park et al. (2008) evaluated the modulation of virulence factor expression and transcriptional levels of QS-regulated genes in P. aeruginosa by solenopsin A (trans-2-methyl-6-undecylpiperidine) and found that solenopsin A disrupted quorum sensing signaling. Even more interesting was the observation that the addition of C4-homoserine lactone, but not 3-oxo-C12homoserine lactone, restored P. aeruginosa quorum-sensing signaling, suggesting that solenopsin A targets the C4- homoserine lactone-dependent rhl quorumsensing system. Both C4-homoserine lactone and 3-oxo-C12- homoserine lactone are signaling molecules involved in quorum sensing of P. aeruginosa. Arbiser et al. (2007) found trans-2-methyl-6-undecylpiperidine had antiangiogenic activity by inhibiting the phosphatidylinositol-3-kinase (PI3K) signaling pathway. Although angiogenesis is a normal physiological process, it is also a fundamental step in the transition of tumors from a dormant state to a malignant state. PI3K and its downstream effector Akt, or proteinkinase Bα(PKBα) regulate apoptosis, proliferation, mad angiogenesis. PI3K and Akt are over-expressed in many cancers, such as sarcomas (cancer of connective tissue), ovarian cancer, multiple myeloma (cancer of plasma cells), and melanoma (a skin cancer). The production of the angiogenic factor VEGF (vascular endothelial growth factor) was also controlled by the PI3K/Akt pathway. VEGF protects cancer cells from both chemotherapy and reactive oxygen–induced apoptosis through phosphorylation of substrates such as apoptotic peptidase–activating factor-1 (APAF-1), forkhead proteins, and caspase 9. Compounds that suppress the PI3K/Akt pathway may have potential medical application as angiogenesis inhibitors and antineoplastic agents. Consistent with inhibition of the activation of PI3K, trans2-methyl-6-undecylpiperidine prevented the phosphorylation of Akt and the phosphorylation of its substrate forkhead box 01a (FOXO1a), a member of the forkhead family of transcription factors. Interestingly, trans-2-methyl-6-undecylpiperidine also inhibited Akt-1 activity in an ATP-competitive manner in vitro without affecting 27 of 28 other protein kinases tested. Arbiser et al. (2007) believed that trans-2-methyl-6-undecylpiperidine could be easily conjugated to other molecules for targeted delivery, because it was a free secondary amine. Conjugation of piperidine alkaloids to molecules that are specific for tumors may provide a novel and specific therapy for advanced neoplasms (abnormal proliferation of cells), both in terms of treatment of patients directly and purging of autologous bone marrow for transplantation.

24.4

Proteins

Allergic reaction to stings of the imported fire ants is common in fire ant infested areas. Piperidine alkaloids are responsible for the typical pustule at the sting sites, but they are not allergenic. Four allergens, Sol i I, Sol i II, Sol i III, and Sol i IV,

428


have been identified in the water soluble portion of S. invicta venom. Homologous proteins to Sol i I, Sol i II and Sol i III, which are named Sol r I, Sol r II, and Sol r III respectively, were also identified in S. richteri venom (Hoffman et al. 1990); however, no Sol i IV equivalent was found in S. richteri venom. These allergens cause anaphylactic reactions in highly sensitive individuals. Hoffman et al. (1988) isolated and purified these four allergens using gel filtration and high-performance cation exchange chromatography. Basic information of those allergens is summarized in Table 24.4. Table 24.4

Basic information on four allergens in imported fire ants venom Sol i I

Sol i II

Sol i III

Sol i IV

MW (da)

37,000

13,217

24,040

13,340

PI

8.51

9.63

8.24

10.08

No.of amino acid

148

119

212

117

Content (%)

2–4

50–67

15–25

8–10

Sol i I is a lipase of the GX class, lipoprotein lipase superfamily, pancreatic lipase homologous family and RP2 subgroup phospholipases. Vespid wasp venom phospholipases belong to the same subgroup. Sequence alignment with eight vespid venom phospholipases showed conservation of the regions on the enzyme active site area, this may explain why Sol i I exhibits some crossreactivity with IgE antibodies from patients sensitized to other Hymenoptera venoms. Sol i I is the only major fire ant allergen exhibiting cross-reactivity with wasp venom proteins (Hoffman et al. 1988; Nordvall et al. 1985). Sol i II is a disulfide-linked homodimer and each subunit containing seven cysteine residues (Hoffman 1993). No structurally related proteins were found in the Protein Identification Resource or Swiss-Prot databases. Sol i II has an acid labile aspartyl-prolyl bond at position 70 and is susceptible to proteolysis. Since each subunit contains only one tyrosine and one tryptophan, Sol i II has a relatively low absorbance at 280 nm. The highly purified Sol i II does not have phospholipase activity. Sol i II is the most variable among species (Hoffman 1997). Sol i III is a single chain protein, containing eight cysteine residues. It belongs to the PR-1 protein family (Henriksen et al. 2001) and is about 44%–50% identical to five antigen 5 molecules from Vespula, Dolichovespula, and Vespa. Despite the significant amino acid sequence homology, no consistent immunologic cross-reactivity was found between Sol i III and vespid antigen 5 (Hoffman et al. 1988; Padavattan et al. 2008). Sol i IV is also a single chain protein, containing six cysteine residues. Its sequence is 35% identical to Sol i II, but it is not significantly related to other proteins. Sol i IV has no tryptophan residues and has two variant forms differing by two amino acids. Sol i IV is not immunologically cross-reactive with Sol i II (Hoffman 1993).


429

Sequences of all four proteins have been obtained (Hoffman et al. 2005; Hoffman 1993; Schmidt et al. 1993). In order to understand why these proteins are such potent allergens, the molecular features of these proteins must be determined. The sequence information is a prerequisite to such an attempt. Sequence information is also useful in studies to characterize the T- and B-cell determinants. These determinants can be synthesized and used to modulate the specific immune response to each allergen by reducing specific IgG blocking antibody production (Hoffman 1993). The sequence is also useful in potential production of nonallergenic vaccines for specific immunotherapy (Baldo 1991). Whole body extracts have been used in diagnosis and immunotherapy of imported fire ant stings. Whole body extracts may contain non-venom proteins and quantities of allergens may vary among preparations. Although allergen specificity and activity can be improved using pure venom (Stafford et al. 1992), piperidine alkaloids in pure venom can cause an immediate wheal and flare reaction. One solution is to express allergens as recombinant proteins. Production of recombinant proteins overcomes the difficulty of collecting large quantities of allergens from ants. The use of a hexahistidine tag simplifies purification. All four allergens have been cloned and expressed as recombinant proteins (Han et al. 2009; Hoffman et al. 2008; Han et al. 2007; Schmidt et al. 2003; Schmidt and Hoffman 2001; Schmidt et al. 1996; Schmidt et al. 1993). As mentioned above, although Sol i III has significant amino acid sequence homology with vespid wasp antigen 5, no consistent immunologic crossreactivity has been found between them. Padavattan et al. (2008) determined the crystal structure of recombinant (Baculovirus) Sol i III to a resolution of 3.1 Å by the method of molecular replacement. They found that secondary-structure elements of Sol i III is an α-β-α sandwich folding consisting of a central antiparallel β -sheet surrounded on both sides by a α-helices. The overall structure of Sol i III is very similar to that of the homologous wasp venom allergen Ves v 5. The major difference resides in the solvent-exposed loop regions that contain amino acid insertions. The limited conservation of surface chemical properties and topology between Sol i III and Ves v 5 may explain the lack of relevant crossreactivity revealed by immunochemical and serological studies with human IgE antibodies (Padavattan et al. 2008). Understanding the chemical components of the venom is the fundation for studies about its functions. In contrast to the significant progress in understanding the protein allergens in imported fire ants venom, few efforts have been made to further identify chemical compounds in the water-insoluble phase of the venom. Much of our knowledge on the toxicity of imported fire ant venom was from the research on piperidine alkaloids. The identification of piperideines indicates that there may be many other important compounds in the venom. Future research may help improve our understanding of the chemistry of imported fire ant venom and enable us to better understand its functions.

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Acknowledgements The authors thank Abner M. Hammond, Jr. at Department of Entomology, Louisiana State University, Baton Rouge, Louisiana and Douglas A. Streett and Eric W. Riddick at Biological Control of Pests Research Unit, USDA-ARS, Stoneville, Mississippi for their comments and suggestions on this chapter.

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Alabama. J. Entomol. Sci., 1988, 23: 187–191. Dring J V, Nash R J, Roberts M F, et al. Hemlock alkaloids in Aloes. Occurrence and distribution of γ-coniceine. Planta Med. 1984, 50: 442–443. Fitzgerald K T, Flood A A. Hymenoptera stings. Clin. Tech. Small Anim. Pract., 2006, 21: 194–204. Freeman T M. Hymenoptera hypersensitivity in an imported fire ant endemic area. Ann. Allergy Asthma Immunol. J. Nat. Toxins, 1997, 78: 369–372. Gardner W A, Diffie S, Vander Meer R K. Distribution of the fire ant (Hymenoptera: Formicidae) hybrid in Georgia. J. Entomol. Sci., 2008, 43: 133–137. Gerson E A, Kelsey R G. Piperidine alkaloids in North American Pinus taxa: Implication for chemosystematics. Biochem. Syst. Ecol., 2004, 32: 63–74. Gibbons L, Simberloff D. Interaction of hybrid imported fire ants (Solenopsis invicta x S. richteri) with native ants at baits in southeastern Tennessee. Southeast. Nat., 2005, 4: 303–320. Greenberg L, Kabashima J N, Allison C J, et al. Lethality of red imported fire ant venom to Argentine ants and other ant species. Ann. Entomol. Soc. Am., 2008, 101: 1162–1168. Haight K L, Tschinkel W R. Patterns of venom synthesis and use in the fire ant, Solenopsis invicta. Toxicon, 2003, 42: 673–682. Han X Q, Lin X M, Chen H J, et al. The Prokaryotic expression and bioactivity of the recombinant red fire ant venom allergen Sol i 4. Agricultural Sciences in China, 2009, 8: 182–187. Han X Q, Lin X M, Zhang Y G, et al. Expression and activity analysis of the recombinant Solenopsis invicta Buren venom allergen Sol i 1. Acta Entomol. Sin., 2007, 50: 655–661. Henriksen A, King T P, Mirza O, et al. Major venom allergen of yellow jackets, Ves v 5: structural characterization of a pathogenesis-related protein superfamily. Proteins, 2001, 45: 438–448. Hoffman D R, Peroutka C M, Schmidt M. Imported fire ant venom phospholipase, Sol i 1, is a protein allergen. J. Allergy Clin. Immunol., 2008, 121: S30. Hoffman D R, Sakell R H, Schmidt M. Sol i 1, the phospholipase allergen of imported fire ant venom. J. Allergy Clin. Immunol., 2005, 115: 611–616. Hoffman D R. Reactions to less common species of fire ants. J. Allergy Clin. Immunol., 1997, 100: 679–683. Hoffman D R. Fire ant venom allergy. Allergy, 1995, 50: 535–544. Hoffman D R. Allergens in hymenoptera venom XXIV: The amino acid sequences of imported fire ant venom allergens Sol i II, Sol i III, and Sol i IV. J. Allergy Clin. Immunol., 1993, 91: 71–78. Hoffman D R, Smith A M, Schmidt M, et al. Allergens in hymenoptera venom XXII: Comparison of venoms from two species of imported fire ants, Solenopsis invicta and richteri. J. Allergy Clin. Immunol., 1990, 85: 988–996. Hoffman D R, Dove D E, Jacobson R S. Allergens in Hymenoptera venom. XX. Isolation of four allergens from imported fire ant (Solenopsis invicta) venom. J. Allergy Clin. Immunol., 1988, 82: 818–827. ISSG. Solenopsis invicta (insect). Global invasive species database. http://www.issg.org/database/ species/distribution.asp?si = 77&fr = 1&sts = &lang = EN (April 28, 2009). Jones T H, Blum M S, Fales H M. Ant venom alkaloids from Solenopsis and Monomorium species. Recent developments. Tetrahedron, 1982, 38: 1949–1958. Langkilde T. Invasive fire ants alter behavior and morphology of native lizards. Ecology, 2009, 90: 208–217. Leclercq S, Braekman J C, Daloze D, et al. Biosynthesis of the solenopsins, venom alkaloids of the fire ants. Naturwissenschaften, 1996, 83: 222–225.

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Leclercq S, Thirionet I, Broeders F, et al. Absolute configuration of the solenopsins, venom alkaloids of the fire ants. Tetrahedron, 1994, 50: 8465–8478. Leete E. The biosynthesis of Coniine from four acetate units. J. Am. Chem. Soc., 1963, 85: 3523– 3524. MacConnell J G, Williams R N, Brand J M, et al. New alkaloids in the venoms of fire ants. Ann. Entomol. Soc. Am., 1974, 67: 134–135. MacConnell J G, Blum M S, Fales H M. The chemistry of fire ant venom. Tetrahedron, 1971, 26: 1129–1139. MacConnell J G, Blum M S, Fales H M. Alkaloid from fire ant venom: Identification and synthesis. Science, 1970, 168: 840–841. Menzel T O, Cross D C, Nebeker T E, et al. A survey of imported fire ant (Hymenoptera: Formicidae), species and social forms across four counties in east-central Mississippi. Midsouth Entomol., 2008, 1: 3–10. Mobley D, Redding J. USDA amends imported fire ant quarantine. USDA-APHIS press releases. http://www.aphis.usda.gov/lpa/news/ 2005/08/ fireantq_ppq.html. 2005. Morrison J E, Williams D F, Oi D H, Potter K N. Damage to dry crop seed by red imported fire ant (Hymenoptera: Formicidae). J. Econ. Entomol., 1997, 90: 218–222. Nash R J, Beaumont J, Veitch N C, et al. Phenyethylamine and piperidine alkaloids in Aloe species. Planta Med., 1992, 58: 84–86. Nordvall S L, Johansson S G O, Ledford D K, et al. Allergens of imported fire ant (IFA), Solenopsis invicta. J. Aller. Clin. Immunol., 1985, 75: 122. Obin M S, Vander Meer R K. Gaster flagging by fire ants (Solenopsis spp.): Functional significance of venom dispersal behavior. J. Chem. Ecol., 1985, 11: 1757–1768. Oliver J, Vander Meer R K, Ochieng S, et al. Statewide survey of imported fire ant (Hymenoptera: Formicidae) populations in Tennessee. J. Entomol. Sci., 2009, 44: 149–157. Padavattan S, Schmidt M, Hoffman D R, et al. Crystal structure of the major allergen from fire ant venom, Sol i 3. J. Mol. Biol., 2008, 383: 178–185. Park J, Kaufmann G F, Bowen J P, et al. Solenopsin A, a venom alkaloid from the fire ant Solenopsis invicta, inhibits quorum-sensing signaling in Pseudomonas aeruginosa. J. Infect. Dis., 2008, 198: 1198–1201. Reynolds T. Hemlock alkaloids from Socrates to poison aloes. Phytochemistry, 2005, 66: 1399– 1406. Ross K G, Vander Meer R K, Fletcher D J C, Vargo E L. Biochemical phenotypic and genetic studies of two introduced fire ants and their hybrid (Hymenoptera: Formicidae). Evolution, 1987, 41: 280–293. Sánchez-Pena R, Patrock R J W, Gilbert L E. The red imported fire ant is now in Mexico: Documentation of its distribution along the Texas-Mexico border. Entomol. News, 2005, 116: 363–366. Schmidt M, McConnell T J, Hoffman D R. Immunologic characterization of the recombinant fire ant venom allergen, Sol i 3. Allergy, 2003, 58: 342–349. Schmidt M, Hoffman D R. Expression of recombinant fire ant venom allergen Sol i 4. J. Allergy Clin. Immunol., 2001, 107: S112–S113. Schmidt M, McConnell T J, Hoffman D R. Production of a recombinant imported fire ant venom allergen, Sol i 2, in native and immunoreactive form. J. Allergy Clin. Immunol., 1996, 98: 82–88.


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Schmidt M, Walker R B, Hoffman D R, et al. Cloning and sequencing of cDNA encoding the fire ant venom protein Sol i II. FEBS Lett., 1993, 319: 138–140. Stafford C T, Wise S L, Robinson D A, Hoffman D R. Safety and efficacy of fire ant venom in the diagnosis of fire ant allergy. J. Allergy Clin. Immunol., 1992, 90: 653–661. Stafford C T, Hoffman D R, Rhoades R B. Allergy to imported fire ants. South Med. J., 1989, 82: 1520–1527. Streett D A, Freeland T B, Vander Meer R K. Survey of imported fire ant (Hymenoptera: Formicidae) populations in Mississippi. Florida Entomol., 2006, 89: 91–92. Tschinkel W R. The fire ants. Cambridge: Harvard University Press, 2006: 365–382. Vander Meer R K, Lofgren C S, Alvarez F M. Biochemical evidence for hybridization in fire ants. Fla. Entomol., 1985, 68: 501–506. Vinson S B. Invasion of the red imported fire ant (Hymenoptera: Formicidae): Spread, biology, and impact. Am. Entomol., 1997, 42: 23–39. Wojcik D P, Craig R A, Brenner R J, et al. Red imported fire ants: impact on biodiversity. Am. Entomol., 2001, 47: 16–23.

CHAPTER 25

Utilization of Lignocellulose-feeding Insects for Viable Biofuels: an Emerging and Promising Area of Entomological Science Jianzhong Sun and Xuguo Joe Zhou

Abstract Most insects are unable to use plant cell walls as their main food sources, but some insects subsist on lignocellulosic biomass from agricultural crops to forest woody substrates as their only foods, such as in the case of termites (all seven families), wood-feeding roaches (Blattidae, Cryptoceridae), beetles (Anobiidae, Buprestidae, Cerambycidae, Scarabaeidae), wood wasps (Siricidae), leaf-shredding aquatic insects (Pteronarcidae, Limnephilidae, Tipulidae), silver fish (Lepismatidae), leaf-cutting ants (Formicidae), etc. Cellulose digestion has been demonstrated in more than 20 families representing ten distinct insect orders, e. g. Thysanura, Plecoptera, Dictyoptera, Orthoptera, Isoptera, Coleoptera, Trichoptera, Hymenoptera, Phasmida, and Diptera. The ability of these insects to feed on wood, foliage and detritus has recently stimulated extensive investigations into the mechanisms of how these insects digest the structural and recalcitrant lignocellulose in their foods. With these studies, scientists would possibly advance biofuel technologies with the discovery of novel lignocellulolytic enzymes and a better understanding of the bioconversion mechanisms that breakdown plant cell walls inside the insect’s gut. Producing monomeric sugars from cellulose or hemicellulose with high yields and low cost is far more difficult than deriving them from sugar- or starch-containing crops. This difficulty is primarily due to a lack of efficient and economic lignocellulolytic enzymes that convert rigid plant cell walls to their monomeric pentose and hexose sugar subunits. However, termites, especially wood-feeding termites (including lower and higher termites), are a unique group of lignocellulose-feeding insects exhibiting incredible wood degradation capabilities, which accomplish lignocellulose digestion using specialized gut physiology, endogenously produced Jianzhong Sun School of the environment, Jiangsu University, 301 Xuefu Rd., Zhenjiang, Jiangsu, 212013 China E-mail: [email protected] Xuguo Joe Zhou Insect Integrative Genomics, Department of Entomology, University of Kentucky, Lexington, Kentucky, KY, 40546, USA


Utilization of Lignocellulose-feeding Insects for Viable Biofuels

435

digestive enzymes, and via their specialized association with prokaryotic and eukaryotic gut symbionts. It is believed that the guts of these lignocellulosefeeding insects harbor diverse symbiotic microbes and endogenous enzymes that could be used as a rich source of lignocellulases as well as functional gene resources for improving the conversion of wood or waste plant biomass to valuable biofuels. Recent studies showed that lignocellulose-feeding insects and their symbionts have not only cellulolytic or lignin decomposition activity, but also aromatic hydrocarbon degradation. Thus, as an emerged new area of entomological science, utilization of lignocellulose-feeding insects would be very valuable for viable biofuels production made from lignocellulosic biomass. Clearly, understanding the mechanisms of the biomass digestion in these insect guts could potentially shed light on efficient, low cost, lignocellulose-based biofuel production systems. This review addresses various lignocellulolytic systems, the potential values, various challenges, and opportunities that exist for investigating lignocellulose-feeding insects in biofuels production, as well as possible future research directions. Keywords lignocellulose, insect, lignocellulase, lignocellulolytic system, biofuels, bioreactor

25.1

Introduction

Global warming and the depletion of fossil oil are creating an urgent need to find renewable and ecologically friendly fuels, which have received tremendous attention, making many headlines in the public media. As the world reserves of fossil fuels and raw materials are consumed, development of biofuels, such as bioethanol, biodiesel, biohydrogen, etc., is accelerating worldwide. The production and use of biofuels have entered a new era of global growth, experiencing acceleration in both the scale of the industry and the number of countries involved. Brazil, the United States, many European countries, and a growing number of countries in Southeast Asia are now pinning their hopes on biofuels. Of these countries, Brazil, the United States, and China rank as the top three bioethanol producers in the world (Worldwatch Institute 2007). However, almost all bioethanol today is produced from sugar- or starch-based agricultural crops, using so called first-generation technologies (Mcmillan 1997; Otero et al. 2007). Bioethanol accounts for over 90% of the total biofuels production worldwide. In the United States, it currently only contributes less than 5% to the total transportation fuels mix. However, despite a huge market demand for bioethanol or other forms of biofuels, the limited availability of food-based commodities, and their price competition with traditional food, animal food, and other related industries, inevitably restricts the scale of the first generation of biofuel industries and their sustainable development. Plant material is the most abundant lignocellulosic biomass on Earth,

436

Jianzhong Sun and Xuguo Joe Zhou

consisting mainly of cellulose, hemicellulose and lignin. Of these three components, cellulose and hemicellulose comprise the most abundant and renewable source of sugars (pentose and hexose) on the planet and their microbial degradations are of considerable biological and industrial importance. The lignocellulosic biomass, including agricultural and forestry waste residues, has the potential to become a major source of fermentable sugars for biofuels in the future. It is estimated that in the USA alone, more than one billion tons (dry ton) per year of biomass could be sustainably harvested in the form of crop and forestry residues, replacing as much as 30% of the total US gasoline consumption (Merino and Cherry 2007). Therefore, lignocellulosic biomass such as agricultural and forestry residues and herbaceous energy crops (i.e. switchgrass, Panicum virgatum; Miscanthus, Miscanthus sinensis, Chinese silvergrass ) can serve as a large scale, low cost feedstock for production of fuel ethanol and other value-added commodity chemicals. However, the challenge is getting energy out of lignocellulosic biomass efficiently and economically in the format of a sugar-platform (various monomer sugars converted from different polysaccharides of plant cell walls). The lignocellulose-based biofuels are so-called second-generation technology; but currently, it is an expensive process, limited to a handful of pilot plants in the US and the rest of world (Schubert 2006; Merino and Cherry 2007; Jørgensen et al. 2007). Comparing first and second generation technologies, it takes anywhere from 40–100 times more cellulases to break down cellulose than starch if measured on a protein weight basis, yet the cost of enzymes being used in the second generation technology is not substantially different (Merino and Cherry 2007). Thus, there is an acute demand for identification of novel biomass degrading microorganisms and enzymes at higher efficiencies and reduced costs. This has catalyzed an interest in utilizing lignocellulose-feeding insects, such as termites ¾ the world’s smallest, and the most efficient lignocellulosic bioreactors (Brune 1998), to discover the novel lignocellulolytic microorganisms/lignocellulases and the associated encoding genes for viable biofuels (Brune 2007; Warneche et al. 2007; Sun 2008; Matsui et al. 2009). Insects are the largest taxonomic group of animals on Earth and most of them rely on non-cellulosic material as their main food sources. However, cellulose digestion has been demonstrated in some representatives of widely different taxonomic groups of insects, such as lower and higher termites, various beetles, wood wasps, etc (Martin 1983; Prins and Kreulen 1991; Varma et al. 1994). Among them, wood-feeding termites have the unique ability to digest woody material with extremely high efficacy, often using it as a sole food source. The ability of many insects to thrive on wood, dead foliage and detritus with various digestive mechanisms, such as degradation with the symbiotic intestinal microorganisms, acquired microbial cellulases by ingesting fungi, as well as secreting the lignocellulases by themselves, has recently stimulated many investigations for how these insects to digest the structural polysaccharides in their food and efficiently convert the recalcitrant lignocelluloses to the monomer


437

sugars. Further, these insect guts may be remodeled as an efficient bioreactor/ fermentator for a large scale lignocellulose bioconversion by biofuels industry. Given the relative novelty of this field in entomology and its interdisciplinary nature, this chapter will give an overview of the general concepts involved in the deconstruction of lignocellulose by insects, potential values and distinct roles of lignocellulose-feeding insets for viable biofuels. The various mechanisms in lignocellulose degradation by these insects will also be discussed in terms of the recent efforts made by some pioneer scientists from various disciplines. Finally, this chapter summarizes the various challenges and opportunities that exist for the cellulose-feeding insects contributing to present biofuels industry, microbiology, bioengineering, and entomological sciences as well.

25.2 Chemical nature of the lignocellulosic biomass and the enzymatic degradations 25.2.1

Composition of lignocellulosic biomass

Lignocellulosic biomass is a heterogeneous material that includes almost all kinds of living and dead plants. Therefore, the earth’s estimated annual production of lignocellulose would range from 2 to 5 ´1012 metric tons (Glazer and Nikaido 2007). It is composed primarily of cellulose, hemicellulose, lignin and small amounts of extracts (Baeza and Freer 2001). These three main elements represent over 90% of the dry weight of a plant cell wall and bind strongly to each other by non-covalent forces as well as by covalent cross-links, making a composite material. The quantity of each of the polymers, i.e. cellulose, hemicellulose, and lignin, varies considerably with plant species (Table 25.1). Usually, softwoods have a higher content of lignin (25%–35%) than do hardwoods (18%–25%). Hemicellulose is relatively higher in grasses (25%–50%), i.e. switchgrass, than that of wood plants (23%–38%). For agricultural crops, especially the plants in Poaceae, they have a relatively lower fraction of lignin when compared to the woody lignocellulosic biomass (Table 25.1). Dedicated energy crops, such as switchgrass and miscanthus, will follow as a feedstock supplement once a market for cellulosic biomass develops further (Worldwatch Institute 2007). In the future, some bioengineered energy plants, such as poplar tree (Populus trichocarpa), could genetically bring a new composition of plant cell walls that would significantly reduce the recalcitrant extent in enzymatic biodegradation, as well as the biomass yield of these energy crops (US DOE 2006). Table 25.1

Major chemical constituents of different lignocellulosic materials

Lignocellulose biomass Softwoods b

a

Cellulose (%)

Hemicellulose (%)

Lignin (%)

37–50

25–35

25–35

Picea glauca

41

31

27

Abies balsamea

42

27

29

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Jianzhong Sun and Xuguo Joe Zhou (Continued)

Lignocellulose biomass

Cellulose (%)

Hemicellulose (%)

Lignin (%)

Pseudotsuga menziesii

39

23

29

Pinus strobus

41

27

29

Pinus radiata

37

29

27

Tsuga canadensis

41

23

33

Juniperous communis

33

27

32

Thuja occidentalis

41

26

31

b

41–55

23–38

18–25

Acer rubum

45

29

24

Acer saccharum

41

27

25 24

Hardwoods

Ulmus americana

51

23

Populus tremuloides

48

27

21

Betula papyrifera

42

38

19

Betula verrucosa

41

30

22

Eucalyptus globules

51

21

22

Fagus grandifolia

45

29

22

Hybrid poplar

49

16

22

Agricultural crops

27–38

23–28

11–25

Corn stover

35

28

16–21

Rice straw

34

25

12

Wheat straw

38

23

23

Sweet sorghum

27

25

11

Bagasse c

40

24

25

Bamboo

41–49

24–28

24–26

Grasses

25–51

25–50

10–30

Swichgrass

44–51

42–50

11–17

Miscanthus

44

24

26

Reed canary grass

26

12.3

15.6

Weeping lovegrass

36.7

21.9

21.2

36

25

18

Office paper

69

20

11

Municipal solid waste paper

56

14

30

Big bluestem Others

a) All values are given as percentage of the wood dry weight. Data were adapted from Wiselogel et al. 1996; Ragauskas et al. 2006a; Worldwatch Institute 2007; Jørgensen et al. 2007; and Glazer and Nikaido 2007. b) For commercial purposes, wood are divided into hardwoods and softwoods. Hardwoods (angiosperm) are the woods of dicotylendons irrespective of whether they are hard or soft. Dicotylendons are a class of flowering plants generally characterized as having two cotylendons (seed leaves). Softwoods (gymnosperm) are the wood of conifers. c) Bagasse is the residue after the juice is extracted from sugar cane.


25.2.2

439

Cellulose

The predominant polysaccharide in wood or other herbaceous plants is cellulose, which, in situ, is a homo-polysaccharide composed entirely of Dglucose linked together by β - (1, 4)-glycosidic bonds and with a degree of polymerization (DP: number of monomeric residues per polymer molecule) of up to 10,000 or higher (Fig. 25.1A). The linear structure of the cellulose chain enables the formation of both intra- and intermolecular hydrogen bonds resulting in the aggregation of chains into elementary crystalline fibrils of 36 cellulose chains (so-called microfibrils) (Jørgensen et al. 2007). The microfibrils (250Å wide) are combined to form larger fibrils. The basic structure of an elementary fibril is crystalline; but some parts of the chain surface could be viewed as amorphous (Ward and Moo-Young 1989; Ding and Himmel 2006). Therefore, cellulose fibrils have regions of high order, crystalline regions, and regions of less order, amorphous regions (more accessible for cellulases) (Fig. 25.1B). Nevertheless, there is no sharp boundary between these regions. The fraction of crystalline cellulose and the chain length of cellulose vary from source to source. In higher plants, the DP values appear to generally range from 7,000 to 14,000 (Richmond 1991).

Fig. 25.1 Structure of cellulose (reprinted from Béguin and Aubert 1994). (A) β-Glucosidic bonds. (B) Schematic structure of a microfibril.

The structure of cellulose along with the intermolecular hydrogen bonds gives cellulose high tensile strength when intertwined with hemicellulose and lignin components, thus making cellulose insoluble in most solvents (Word and MooYoung 1989). The cellulose exists in the form of microfibrils that is also deeply embedded in hemicellulose and lignin, and is consequently well protected from enzymatic or chemical attack. Therefore, it is much more resistant to microbial degradation than are other glucose-based polymers, such as starch. In current bioconversion processes to ethanol or other chemicals, a thermochemical

440


pretreatment step is required to disrupt the lignocellulosic structure of biomass. This makes the cellulose more accessible to hydrolytic enzymes and allows partial decomposition of the highly crystalline structure of the cellulose fibers. 25.2.3

Hemicellulose

Hemicelluloses are complex heterogeneous polysaccharides composed of monomeric residues: pentoses (D-xylose, L-arabinose), hexoses (D-glucose, Dgalactose, L-galactose, D-mannose, L-rhamnose), and uronic acids (D-glucuronic acid and 4-O-methyl-D-glucuronic acid) (Glazer and Nikaido 2007; Jørgensen et al. 2007) (Fig. 25.2). These residues are variously modified by acetylation or methylation. Unlike cellulose being a linear homopolymer (glucose) with little variation in structure from one plant species to another, hemicelluloses are highly branched, generally non-crystalline heteropolysaccharides. They show a much lower degree of polymerization (DP < 200 sugar residues) than cellulose.

Fig. 25.2 Monomeric building blocks of lignocellulose for carbohydrates.

The main hemicellulose wall components in the majority of plants are xyloglucans (Xy-Glc) and their basic function relies on cross-linking ability, which can complex cellulose microfibrils, to form together a large threedimensional skeleton framework (Kqczkowski 2003). There are three major hemicelluloses in softwoods: glucomannan, galactoglucomannans and arabinoglucuronoxylan. The two mannose-containing polymers differ greatly in galactose content. Their approximate sugar compositions (galactose: glucose: mannose) are 1∶1∶4 and 1∶1∶3, respectively (Glazer and Nikaido 2007). Their backbone consists of 1, 4-linked β-D-glucopyranose and β-D-mannopyranose


441

units. As contrast, the major hardwood hemicellulose is glucuronoxylan, with a backbone of 1, 4-linked β-D-xylopyranose units. Generally, the predominant hemicelluloses for hardwoods are glucomannan and glucuronoxylan, with lesser amounts of galactans and glucans (Ragauskas et al. 2006a). Similar to hardwoods, xylan is also the predominant hemicellulose component for both agricultural residues and herbaceous grasses. Hemicelluloses are composed of neutral sugars, uronic acid and acetyl groups, and all represented as their respective anhydrides, i.e. xylan, araban, galactans, and mannan (Table 25.2). Three examples of hemicellulose structures, xyloglucan, glucuronoarabinoxylan, and galactomannan, are presented in Fig. 25.3 (Pauly and Keegstra 2008). The further discussion on hemicellulose structure is also available from the reviews by Brigham et al. (1996) and Glazer and Nikaido (2007). 25.2.4

Lignin

Lignin is the most variable, abundant aromatic polymer in the nature, and it represents a recalcitrant part of lignocellulose and comprises 20–30% of the dry weight of soft and hard woods (Table 25.1). Lignin is found in the cell walls of higher plants (gymnosperm and angiosperm), ferns, and club mosses, predominantly in the vascular tissues specialized for water transport. In structure, lignins are the diverse group of three-dimensional polymers of aromatic alcohols that include three direct precursors of lignin, i.e. coniferyl, sinoapyl, and pcoumaryl (Higuchi 1990; Lewis and Yamamoto 1990) (Fig. 25.4). Lignins are also described as softwood lignin, hardwood lignin, or grass lignin, on the basis of the relative amounts of these building blocks (aromatic alcohols, or monolignols) (Glazer and Nikaido 2007). A typical softwood (gymnosperm) lignin consists of building blocks originating mainly from coniferyl alcohol, some from p-coumaryl alcohol, but none from sinapyl alcohol. While hard wood (angiosperm) lignin contains equal amounts (46% each) of coniferyl- and sinapylpropane units and a minor amount (8%) of p-hydroxyphenylpropane units (derived from p-coumaryl alcohol). Grass lignin is composed of coniferyl-, sinapyl-, and p-hydroxyphenylpropane units, with p-coumaric acid (5%–10%) (Higuchi 1990; Kirk 1987). Most lignin is found within the cell walls, where it is intimately interspersed with the hemicelluloses, forming a matrix that surrounds the orderly cellulose microfibrils. In wood, lignin is in high concentration in the glue that binds contiguous cells, forming the middle lamella. Unlike cellulose or hemicellulose, lignin contains no chains with repeating subunits, thereby making the enzymatic hydrolysis of this polymer extremely difficult (Kirk 1987). 25.2.5

Lignocellulose degradation and depolymerization

Currently, there are three main industrial biorefinery processes used to deconstruct lignocellulosic biomass that are usually categorized into the

442


Fig. 25.3 Three examples of hemicellulose structures commonly present in plant cell walls, including monosaccharides, and their linkage and ester constituents. Easily fermentable hexoses are distinguished by their square shape. Other sugars are shown in


443

Percentage of sugars in hemicellulose from various lignocellulosic sources

Table 25.2

Lignocellulose sources

Xylan

Mannan

Araban

a

Galactan

Softwoods Pine

8.8

11.7

2.4

–

Radiata pine

5.9

11.3

1.6

2.5

Douglas fir

2.4

12.2

1.1

3.5

Spruce

8.6

16.3

2.3

3.7

Hardwoods Maple

17.3

2.9

2.8

–

Poplar

17.4

4.7

1.8

1.2

Hybrid poplar

14.6

0.5

0.3

0.3

White oak

18.0

2.9

2.4

0.4

Black locust

16.2

1.0

0.4

0

Beech

20.8

0.9

1.5

– 1.0

Agricultural crops b

18.0

0.6

3.0

Rice straw

24.5

–

–

–

Wheat straw

21.2

0.3

2.5

0.7

21.1

0.3

1.9

0.5

Corn stover

Bagasse

c

Grasses Swichgrass

20.4

0.3

2.8

0.9

Tall fescue

13.8

0.3

2.9

1.1

Weeping lovegrass

17.6

–

2.6

1.7

Reed canary grass

9.8

0

2.4

0.1

Weeping lovegrass

17.6

0

2.6

1.7

Flatpea hay

7.4

0.1

2.0

1.5

Big blue stem

21.2

0.3

2.7

0.8

Office paper

12.4

7.8

–

–

Municipal solid waste paper

8.3

5.6

–

14

Others

a) All values are given as percentage of the dry weight compositions in hemiceluloses. Data were adapted from McMillan 1997; Wiselogel et al. 1996; Ragauskas et al. 2006a; Kumari et al. 2006; and Glazer and Nikaido 2007. b) Corn stover includes corn stalks and cobs as they come out of the combine. c) Bagasse is the residue after the juice is extracted from sugar cane.

different shapes (Glc: glucose, Gal: galactose, Man: mannose, Xyl: xylose, Ara: arabinose), (reprinted from Pauly and Keegstra 2008).

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Fig. 25.4 The three common monolignols in lignin structure: (A) r-coumaryl alcohol, (B) coniferyl alcohol, (C) Sinapyl alcohol, and an example of a possible lignin structure.

thermochemical (pyrolysis, gasification) and biochemical pathway (hydrolysis) (Lange 2007; Gomez et al. 2008). The thermochemical pathway needs an intensive energy input ( > 500–800°C heating) to facilitate the processing that inevitably lead to an increase in both cost and carbon footprint. However, the biochemical pathway mainly involves converting biomass into sugars by enzymatic hydrolysis with much less energy requirement during the processing. It is believed that the cellulases and their encoding genes derived from some


445

natural life systems, such as lignocellulose-feeding insects, are highly valuable for the application in enzymatic hydrolysis of biochemical processing (Scharf and Tartar 2008; Sun 2008; Matsui et al. 2009). It will be helpful to understand some essential enzymes involved in biological degradation to cellulose, hemicellulose and lignin prior to the discussion on the digestive systems of lignocellulose-feeding insects. It is currently accepted that at least three different types of enzymes are involved in the complete hydrolysis of native cellulose to glucose: exo-1,4-βglucanases (cellobiohydrolases: CBHs) (EC 3.2.1.91), which specifically cleave cellobiosyl units from the non-reducing ends of the cellulose chains; endo1,4-β-glucanases (EC 3.2.1.4), which cleave internal cellulosic linkages; and βglucosidases (EC 3.2.1.21), which specifically cleave glucosyl units from the nonreducing ends of cello-oligosaccharides. The CBH has been widely described in the literature in terms of its activity on microcrystalline cellulose (Avicel) with cellobiose (G2 units) as the main product and its inability to hydrolyze the internal cellulose linkages of carboxymethyl cellulose (CMC). The term ‘‘Avicelase’’ is commonly regarded as synonymous with exoglucanases or CBH (Beguin and Aubert 1994; Lynd et al. 2002; Rabinovich et al. 2002). It is assumed that endoglucanases can only attack the amorphous regions of cellulose molecules (Fig. 25.1B). The degradation of the crystalline regions is thought to occur as a result of CBH action catalyzing the release of G2 residues from the nonreducing ends of cellulose chains. In fact, complete or nearly complete hydrolysis of cellulose typically requires synergistic collaboration by each of these three types of cellulases (Fig. 25.5). To date, most studies of enzymatic hemicellulose degradation have focused on xylans, to the virtual exclusion of galactoglucomannans and glucomannans. Xylan is the most abundant hemicellulose and xylanases are one of the major hemicellulases which hydrolyze the β-1, 4 bond by endo- and exo-β-1, 4xylanases in the xylan backbone yielding short xylooligomers that are further hydrolyzed into single xylose units by β-xylosidase. The complete conversion of a glucuronoxylan into its building blocks requires the combined activity of endoand exo-β-1, 4-xylanases, β-xylosidases, α-arabinofuranosidaes, α-uronidases, and esterases such as acetylxylan esterase, ferulic acid esterase. Based on the amino acid or nucleic acid sequence of their catalytic modules, hemicellulases are either glycoside hydrolases (GHs) that hydrolyze glycosidic bonds or carbohydrate esterases (CEs), which hydrolyze ester linkages of acetate or ferulic side groups (Shallom and Shoham 2003). For details on the types, structure, function, classification of microbial hemicellulases, as well as the catalytic mechanisms refer to some review articles by Merino and Cherry (2007), Shallom and Shoham (2003), Howard et al. (2003), and Rabinovich et al. (2002). The lignin biodegradations are primarily modeled with white rot fungi that represent more than 90% of wood-decaying fungi (white/brown/soft rot fungi > 2000 species) .White rot fungi significantly deconstruct lignin either

446


Fig. 25.5 Synergism of endoglucanases, cellobiohydrolases (exoglucanases) and βglucosidases in fungal cellulase systems. Glucose residues are indicated by hexagons, reducing ends are shown in black (adapted from Béguin and Aubert 1994).


447

with “sequential decay” or a simultaneous degradation of lignin and polysaccharides. These fungi (Basidiomycetes) breakdown lignin aerobically through the use of a family of extracellular enzymes collectively termed “lignases”. Two families of lignolytic enzymes (lignases) are widely considered to play a key role in the enzymatic degradation: phenol oxidase (laccase) and peroxidases (lignin peroxidase: LiP; and manganese: MnP) (Kirk 1987; Howard et al. 2003; Malherbe and Cloete 2003). Successful degradation of lignin requires attacks on both the phenolic and nonphenolic lignin components. Degradation is usually initiated by lignin peroxidase and the phenol oxidases, which catalyze the oxidative degradation of the polymer to oligolignols and then to mono-and dilignols (Glazer and Nikaido 2007). Other seemingly involved enzymes may include H2O2 producing enzymes, glyoxal oxidase, glucose oxidase, veratryl alcohol oxidase, methanol oxidase, alcohol dehydrogenase, etc. (Howard et al. 2003). In nature, hundreds of fungi and bacteria are able to degrade lignocellulose. These organisms include aerobes and anaerobes, mesophiles and thermophiles. However, although many microorganisms can grow on cellulose-rich substrates or produce enzymes that can degrade amorphous cellulose, relatively few produce the entire complement of extracellular lignocellulases able to degrade crystalline cellulose and lignin in vitro (Glazer and Nikaido 2007). The development of cost-effective lignocellulosic biocatalysts for the industrial utilization will require continued efforts to find the efficient natural life systems that can efficiently deconstruct plant biomass, such as termites or other lignocellulose-feeding insects.

25.3 Diversity of lignocellulose-feeding insects and their lignocellulolytic systems Cellulose digestion has been demonstrated in the insect orders Thysanoptera, Plecoptera, Dictyoptera, Orthoptera, Isoptera, Coleoptera, Hymenoptera, Trichoptera, Phasmida, and Diptera (Martin 1983; Prins and Kreulen 1991). However, considering the number and diversity of insects that thrive on diets consisting largely of plant tissue, it is surprising how few are able to digest plant cell walls (lignocelluloses), the most abundant non-fossil carbon source on Earth (Martin 1991). In fact, the ability to digest lignocellulose is quite rare in insects and is limited to a small number of insect families (~20) (Prins and Kreulen 1991). Although the capability of lignocellulose digestion in insects is most often attributed to symbiont-mediated decomposition (a so-called symbiont-dependent system) a variety of these insects also uniquely possess a capability to produce their own lignocellulases from various gut tissues, such as saliva glands and the midgut, to assist lignocellulose digestion (a so-called symbiont-independent system). Both lignocellulolytic systems may occur in the same insect gut to synergistically process lignocellulosic diets (Nakashima et al. 2002a; Scharf and Tartar 2008). In addition, the sophisticated and well-coordinated cooperation

448


between some insect species, such as termites or ants, and the symbiotic fungi growing in their nests, such as Termitomyces albuminosus, enables efficient degradation of lignocellulose (a so-called acquired enzyme system: enzymes are mainly obtained from the cultivated symbiotic fungi). 25.3.1

Cellulose-feeding insects and their digestibility

Although most phytophagous insects lack the capability to utilize plant cell wall from their plant tissue diets (refer to cytoplasm consumers), cellulose digestion has been observed in representatives of diverse taxonomic groups: various woodfeeding insects (woodroaches – Dictyoptera, lower and higher termites –Isoptera, various beetles – Coleoptera, and wood wasps – Hymenoptera), detritus/litterfeeding insects (e.g. leaf shedding aquatic insects at the immature stage – Trichoptera, Diptera, and Plecoptera, silverfishes – Thysanura, crickets – Orthoptera), and forage-feeding insects (beetles – Coleoptera). It is estimated that there are more than 100 insect species reported to have the capability of cellulolytic activity. Table 25.3 lists most of reported lignocellulose-feeding insects, representing various insect taxa in order and family, as well as the evidence to be presented for the utilization of lignocellulose via various means. Table 25.3

Lignocellulose-feeding insects

a

Evidence for capability to digest cellulose b

Order

Family

Representative species

Thysanura

Lepismatidae

Ctenolepisma lineata

Orthoptera

Gryllidae

Acheta domesticus

Acrididae

Schistocerca gregaria

Cryptocercidae

Cryptocercus punctulatus

EN

Periplaneta americana

14

Periplaneta australasia

EN

Pycnoscelus surinamensis

EN

Blaberidae

Gromphadorrhina portentosa

EN

Phasmida

Phasmatidae

Eurycantha calcarata

Isoptera

Mastotermitidae

Mastotermes darwiniensis

AD (72–87), 14C, EN

Lepisma sp.

Dicyoptera

Blattidae

Kalotermitidae

EN AD (41) EN

C

EN EN, 14C

Kalotermes flavicollis

AD (74–91)

Cryptotermes brevis

AD (86), 13C

Neotermes bosei Hodotermitidae

Zootermopsis angusticollis

Rhinotermitidae

Coptotermes formosanus

EN AD (82), EN, 13C 14

C, EN, AD (75), DNA

C. acinaciformis

14

C. lacteus

14

C C


449 (Continued)

Order

Family



Reticulitermes flavipes

AD (91), 14C, EN, DNA

R. lucifugus

AD (96–99), EN

R. speratus

EN, DNA 14

R. santonensis Heterotermes indicola Termitidae

C

AD (78–89)

Macrotermes natalensis

EN

M. bellicosus

EN

M. subhyalinus

EN

M. michaelseni M. gilvus

EN

Odontotermes formosanus

EN, DNA

N. exitiosus N. nigriceps Nasutitermes sp. Cubitermes orthognathus Pteronarcyidae

Pteronarcys proteus

Trichoptera

Limnephilidae

Pycnopsyche luculenta

Hymenoptera

Siricidae

Sirex cyaneus

Formicidae

Coleoptera

Anobiidae

AD (91–97) EN, 14C 14

C

DNA (hindgut) 14

C

AD (11),

14

AD (12),

14

AD (22)

S. phantoma

AD (31)

Atta colombica

EN

A. cephalotes

EN

A. sexdens

EN

Acromyrmex octospinosus

EN

A. crassispinosus

EN

A. echinator

EN

A. subterraneus

EN

Anobium punctatum Ernobius mollis Ptilinus pectinicornis Xestobium rufovillosum

C C

EN

S. gigas

A. striatum

Buprestidae

C, DNA

Trinervitermes trinervoides Nasutitermes ephratae

Plecoptera

EN 13

AD (33), EN AD (31) EN EN AD (68),

Agrilus fuscicollis

EN

Anthaxia corinthia

EN

14

C

450


(Continued) Order

Family

Cerambycidae

Representative species Capnodis miliaris

EN

Capnodis sp.

EN

Chalcophora mariana

EN

Melanophila picta

EN

Acanthocyinus aedilus

EN

Aegosoma scabricornae

EN

Anoplophora glabripennis

EN,

13

Callidium sanguineum

EN

Cerambyx cerdo

EN

Derobrachus brunneus

EN

Ergates faber

EN

Gracilia minuta

C

AD (33)

Hylotrupes bajulus

AD (12–21), EN

Isotomus speciosus

EN

Leptura rubra Leptura sp. Macrotoma palmata Monochamus marmorator

AD (33) EN AD (14–47) AD (27),

14

C, EN

Morimus funereus

EN

Oxymirus cursor

AD (49), EN

Phymatodea testaceus

EN

Physocnemum brevilineum

EN

Plagionotus detritus

EN

Rhagium bifasciatum

EN

R. inquisitor

EN

R. mordax

EN

Saperda populnea

EN

Smodicum cucujiforme

EN

Stromatium barbatum

AD (30–57), EN

S. fulvum Xylotrechus rusticus Apriona germari Scarabaeidae


Oryctes nasicornis

EN EN EN, DNA AD (68),

14

Sericesthis geminata

EN

Pachnoda marginata

EN, DNA

C


451 (Continued)

Order

Diptera

Family


Coccinellidae

Epilachna varivestis

Passalidae

Odontotaenius disjunctus

Curculionidae

Pissodes notatus

Tipulidae

Tipula abdominalis

Evidence for capability to digest cellulose b AD (17–47), 14C, EN EN (by AFM and HPLC) EN AD (19), EN, DNA

a) This Table is primarily modified from Martin (1987). b) AD = cellulose digestion demonstrated by deter mination of assimilation efficiency (value given in 13 14 parentheses) of ingested cellulose. C or C = cellulose digestion demonstrated by detection of the 14 13 incorporation of carbon-14 or 13 into tissue or respiratory CO2 following ingestion of [U- C or d C] cellulose. EN = gut fluid demonstrated to possess enzymatic capability to degrade filter paper, cotton, Avicell, or some other form of insoluble crystalline cellulose. DNA = cellulolytic genes identified from different symbiont metagenomes/transcriptomes and termite genomes.

A huge difference has been reported in cellulose utiliz-ation by various insect species from 11%–99% (Table 25.3). Generally, cellulose digestion in foliage feeders is very rare (Martin 1991). The digestibility that has been determined to date for these insects suggests that termites are the most efficient cellulose digesters in terms of not only their efficiency in cellulose degradation (Fig.25.6), but also the total amount of lignocellulose consumed in each year. Termites have spread throughout the world and now cover 68% of the earth’s landscapes. The large termite population, which has been estimated to be 2.4 ´1017 individuals (Zimmerman et al. 1982), plays a critical role in recycling wood or other lignocellulosic biomass. It has been reported that these termites would consume about 3 – 7 ´1015 g of lignocellulose annually (Breznak and Brune 1994; König et al. 2006). Although the size of a single termite is small ( < 5 mg), the biomass density of termites in tropical and subtropical regions can be so large ( > 50 g/M2) that their impact often surpasses that of grazing mammals (0.013–17.5 g/M2) (Breznak 1984; Breznak and Brune 1994). As one of the main lignocellulosic biomass decomposers on Earth, termites, especially wood-feeding termites, can be inadvertently distributed by human activities, resulting in expanded pantropical/subtropical or even wider distribution (Sun et al. 2007) and further attack urban or rural wood properties. The subterranean termite, Coptotermes formosanus Shiraki, causes more than $1.2 billion in damage annually in the United States (Su and Scheffrahn 1998). Some termites, thus, are well recognized as pests for wooden structures because of their economic impacts. Indeed, only < 4% of total termite species (~2700) in the world can attack man-made products and structural materials. The large majority of termite species are not pests under any circumstance; instead, they carry out wholly beneficial activities. The extent of their benefits is only just beginning to gain recognition and has scarcely been economically evaluated. At present, termites are classified into seven families (with 14 subfamilies) under the order Isoptera, which include 283 genera with approximately 2700

452


Fig. 25.6 The capability of cellulose digestion in some insects. Approximate cellulose digestibility determined by comparing the cellulose contents of food and feces (adapted from Prins and Kreulen 1991; Martin 1983; and Itakura et al. 1995).

species (Krishna 1970; Abe et al. 2000; König et al. 2006) (Fig. 25.7). A large majority of known genera (~85%) are included in one apical family, Termitidae, representing ~72% of all termite species (Abe et al. 2000). Termites are also ordinarily divided into two groups, so-called lower (more primitive) and higher termites, based on their symbiotic system with microorganisms (Fig. 25.7). Lower termites harbor symbiotic cellulosic protists (or flagellates) in their

Fig. 25.7 Phylogenic scheme of termite evolution showing the presumed relationship of the seven termite families, adapted from Higashi and Abe (1997) and Pester 2006. Numbers superimposed on the single branches represent currently recognized genera/ species of the respective families (Abe et al. 2000).


453

hindgut (~28% of total termite species), whereas higher termites do not (one family: Termitidae). The higher and lower termites represent quite different symbiosis in terms of digestion of lignocellulose. Lower termites usually feed on wood, but higher termites show considerable variation in their feeding habits and food substrates that include wood, soil, grass, fungi, and heterogeneous litters. The wood-feeding termites predominantly employ wood as well as other sources of lignocellulose as their main and original food diets. These include the fungus-growing termites (~330 species) that culture the basidiomycete Termitomyces on a substrate of feces produced from ingested wood, litters, or grass, i.e. various lignocellulosic materials. The fungus cultured in termite nests is nitrogen-rich; and it directly decomposes lignocellulose from termite feces. Most importantly, fungi also allow termites to obtain external lignocellulolytic enzymes from ingested fungus comb for efficient lignocellulose degradation (Abo-Khatwa 1978; Martin and Martin 1977; Waller and La Fage 1987). In the real world, the direct and indirect woodfeeding termites would approximately account for > 80% of total termite species, leading to a distinct role in recycling wood or other lignocellulosic biomass on our planet (König et al. 2006). They can be either beneficial or destructive to man, and this is determined to a large degree by which species are present and their feeding and nesting behavior. In contrast to wood-feeding termites, the only detritus feeders from Table 25.3, the immature forms of the stonefly Pteronarcy proteus, the Caddisfly Pycnopsyche luculenta, and crane fly Tipula abdominalis, demonstrate a low rate of cellulose assimilation at 11, 12, and 19%, respectively. The xylophagous larvae of siricid woodwasps as well as anobiid, buprestid and cerambycid beetles are somewhat more efficient, with assimilation rate from 12-68%. The silverfish Ctenolepisma lineate, the American cockroach PeriplanetaAmericana, and the house cricket Acheta demosticus are omnivorous insects that readily incorporate cellulose and non-cellulosic materials in their diets, although they are efficient assimilators of celluloses (41%–87%). It is interesting that most lignocellulose-feeding insects are unable to fully deconstruct the aromatic polymer lignin from woody or herbaceous biomass despite the fact that some disruption or modification processing on the lignin has been observed in various investigations, such as with termites and beetles (Katsumata et al. 2007; Geib et al. 2008). Without this disruption on lignin, the insects would not be able to further biodegrade the polysaccharide components of lignocellulose substrate, because lignin structures serve to protect cellulose and hemicellulose from biodegradation. A study by Itakura (1995) with the lower termite, Coptotermes formosanus, indicated that this species assimilates < 25% of total lignin polymers (Fig. 25.8). However, in contrast to this data, the fungusgrowing termites (Family: Termitidae) can completely degrade lignin polymers and the polysaccharides with help from an exosymbiotic fungus, Termitomyces, cultured in their nest (König et al. 2006). Clearly, some lignocellulose-feeding insects particularly termites have

454


Fig. 25.8 Wood components digestibility (%) by a lower termite species, Coptotermes formosanus, (adapted from Itakura et al. 1995). The letters above each bar represent mean separation and bars with same letter did not differ significantly at α = 0.05, Tukey HSD. Error bars indicate 1 SEM.

developed unique and super-potent cellulolytic systems needed to breakdown various lignocellulosic substrates. Of the lignocellulose-feeding insects we discussed, termites certainly play a significant and indispensable role in carbon recycling on the Earth. 25.3.2

Lignocellulolytic systems in insects

In nature, each lignocellulose-feeding insect has to acquire or develop a cellulolytic system to cope with the recalcitrant plant cell walls. In most cases, this requires a sophisticated cooperation between insects and symbiotic microorganisms either inhabiting their guts (endosymbiosis) or their habitats (exosymbiosis). No evidence has been reported for an insect species that possesses a complete set of insect-derived lignocellulases independently capable of degrading plant cell walls. It is well known that many types of microorganisms, from bacteria, fungi to protozoa, have developed various independent lignocellulolytic systems capable of degrading a variety of the polymers of plant cell walls, such as cellulose, hemicellulose and lignin (Bignell 2000; Jørgensen et al. 2007; Demain 2009; Douglas 2009). Thus, insects must have direct or indirect associations with microorganisms, in a form of endosymbiosis or exosymbiosis, to deconstruct lignocellulosic biomass. These symbiotic relationships between host insects and the microbiota are often obligated. In order to successfully utilize lignocellulosic substrates as their food, insects have


455

to balance the C/N ratio of their food by either adding N (ingesting fungal rotted wood/fixing N from atmosphere) or electively eliminate C (i. e. concentrate N in their food) because many species of termites thrive on sound, decay-free wood which contains as little as 0.03–0.1% nitrogen (dry-weight basis, Cowling and Merrill 1966). Only microbes have the enzymes capable of this, and insects cannot do it alone. As a matter of fact, the C-to-N ratio of sound wood (no microbial decay) is up to 100 fold higher than that of the insect body. Moreover, a lignocellulosic diet typically lacks most of the essential nutrients required by the insect, such as amino acids, vitamins, and sterols (Brune 2003). That is another important reason lignocellulose-feeding insects have to be associated with microorganisms for partial or complete utilization of plant cell walls (Abe and Higashi 1991). Based on the origins of lignocellulases and the relationship between insect hosts and the microorganisms hosted by insects, three lignocellulolytic systems are possessed by insects: symbiont-dependent (microorganism-affiliated, e.g. gut endosymbiosis), symbiont-independent (insect tissue-affiliated), as well as the acquired enzyme system via food ingestion (e. g. fungus-growing: exosymbiosis). The various lignocellulolytic systems may possibly be incorporated into an insect gut system to synergize the processing for the conversion of rigid plant cell walls (a dual lignocellulolytic system). Numerous investigations have indicated that both host insect tissues, such as saliva glands, midgut, and the symbiotic microbiota in insect guts, would produce an array of different complex cellulases that differ in their functions, but complement one another, for an efficient degradation of cellulose (O’Brien et al. 1979; Itakura et al. 1997; Nakashima and Azuma 2000; Nakashima et al. 2002a, 2002b; Watanabe et al. 1998; Li et al. 2006; Slaytor 2006; Zhou et al. 2007; Douglas 2009). More and more evidence via recent research on termite gut cellulases and their encoding genes supports a hypothesis that both gut symbiont-dependent and-independent lignocellulolytic systems in a termite gut digestive system work together through a well coordinated collaboration (Nakashima et al. 2002a; Inoue et al. 2005; Tokuda and Watanabe 2007; Scharf and Tartar 2008). More detailed evidence and a continuous discussion on this subject will be offered in the latter part of this chapter (see section 5). The third important lignocellulolytic system in some insects is mainly referred to as an acquired enzyme system, in which insects regularly encounter fungi and consume fungal tissues to obtain essential lignocellulases (an exosymbiotic association). This mechanism is commonly demonstrated by the fungus-growing insects either as an obligated association with the symbiotic fungi cultured in their nests (i.e. termites, Attine ants, and the siricid woodwasps) or in a facultative association with the fungi encountered in their environments (e.g. cerambycid beetles, the detritus feeders). With this exosymbiosis system between insects and fungi, insects would sustainably obtain a diverse of active and lignocellulolytic enzymes by ingesting fungal fruiting body and the comb substrate (Martin 1987; Abo-Khatwa 1978, 1989; Okuma et al. 2001), combining with insect-derived

456


cellulolytic enzymes in their guts to accomplish a complete degradation of lignocelluloses. The fungus-growing termites are a group of lignocellulose-feeding insects characterized with a highly coevolved and sophisticated association between the insects and the cultured fungi in their nests (Abo-Khatwa 1978, 1989; RoulandLèfevre 2000; Ohkuma et al. 2001). These fungi (genus Termitomyces) are completely obligated and no evidence shows that they occur elsewhere in nature. The mature fungus combs are firm, highly convoluted, sponge-like structures that are constructed from termite fecal pellets with partially digested plant debris. The surfaces of combs are covered with a growth of mycelium and numerous small white spheres, the so-called nodules, that consist of an aggregation of conidiophores and conidia (asexual spores). To construct the fungus comb, different fungus-growing termite species feed on different plant materials; usually wood, leaf litter or grass. For example, Macroterme carbonarius feeds on leaf litter while Odontotermes spp are likely to be predominantly wood-feeders. It has been reported that, in Nigeria, the consumption by fungus-growing termites represented about 25% of the litter production annually generated (RoulandLefèvre 2000); and the extensive degradation of plant material by these insects is due in large part to the genus Termitomyces which serves as the main source of digestive enzymes. The combs are made of plant materials that are collected by the so-called old workers of the nests. Then, the plant litter is chewed by young workers, exchanged between workers, mixed with a liquid that is probably of salivary origin, passed rapidly through the gut, and deposited on the combs in the form of semi-liquid brown or greenish fecal pellets. The symbiotic fungi grow on this fresh comb medium. The fungal nodules then form on the surface of the combs. The lignin content has also been reported progressively decrease as the fungus comb matured (Hyodo et al. 2000), where the laccase (phenol oxidase) was identified as the main lignase in the fungus, the constructed combs, as well as the alimentary tract of the termites (Pan et al. 2009). The fungus nodules are usually ingested by young workers, while the old combs are consumed by old workers to produce final feces. But, final fecal pellets are rarely found in the nests (Ohkuma et al. 2001). This may indicate that a high extent of decomposition and complete biorecycling of plant litter. This lignocellulolytic mechanism is one of the most efficient natural systems and quite unique among the wood-feeding insects due to the degree of lignin degradation, which means cellulose can more easily be attacked by the cellulases produced by the termites (symbiont-independent). There have been several suggestions or hypotheses for the roles of the symbiotic fungi in fungus-growing termite nutrition: (1) decomposition of lignin; (2) supply of cellulase and xylanase to work synergistically with the enzymes produced by the termite; and (3) concentration of nutrients as nitrogen for termites (Ohkuma et al. 2001). Among these hypotheses, most current investigations are focused on the “acquired enzyme system” of which termites exclusively rely on the fungus to digest lignocellulose. However, since some


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endogenous cellulases originated from some termite gut tissues have recently been recognized in wood and litter-feeding termites (Watanabe et al. 1998; Nakashima and Azuma 2000; Scharf and Tatar 2008), the fungus-growing termites may also possess their own cellulases to form a dual lignocellulolytic system. The siricid woodwasps (Hymenoptera, Siricidae) are a group of insects that use the cultured symbiotic fungi in their nests to decompose woody lignocellulose. The damage to a wood tree by woodwasps occurs at the larvae stage as the wood borers and the ovipositing female adults mainly play a role as the vectors of symbiotic fungi. It is reported that two pear-shaped structures at the base of the egg-laying apparatus are filled with fragments of fungal mycelium suspended in a mucous and these mycelium fragments, called arthrospores, are implanted in the wood with an egg during oviposition. Following implantation, the fungus produces vegetative hyphae that colonize the wood surrounding the oviposition hole (Spradbrey 1977; Martin 1987). The fungal associates of siricid woodwasps are basidiomycetes that can also occur as free-living species. The fungal symbionts of species that attack hardwoods differ markedly from those of species that invade softwoods. The fungal associate of Tremex columba, which invades beech, is Daedalea unicolor (Stillwell 1964); whereas the fungal symbionts of the woodwasps attacking conifer trees are in the genus Amylosterum, such as A. chailletii and A. sanguine (Stillwell 1966; Gaut 1969). It has been confirmed that the fungal symbionts of woodwasps are cellulose and hemicellulose digesters and that 22% of the cellulose in fir wood and 31% of the cellulose in spruce wood is assimilated during passage through the guts of Sirex gigas and S. phantoma, respectively (Table 25.3). The midgut of woodwasps is the major site of digestion, where it consists of almost all sets of enzymes required for degrading cellulose, xylan and pectin. With a series of wellconceived and well-executed comparison studies, for the cellulolytic enzyme identities between larval midgut and the symbiotic fungi collected from the larval galleries, the digestive enzymes in woodwasp midgut are primarily derived from the fungal symbionts they ingested (Martin 1987), which indicates that woodwasps primarily possess an acquired enzyme system through their food ingestion. The acquisition of digestive capability through the ingestion of fungal enzymes is also demonstrated in other lignocellulose-feeding insect groups that may have an obligated (leaf-cutting ants) or a facultative association (aquatic leaf-shedding insects) between insects and the symbiotic fungi. The presence of these mechanisms for digesting plant structural polysaccharides in two distantly related taxa, Isoptera and Hymenoptera, suggest that they might have evolved independently. Indeed, the acquired enzymes system has also been combined with another lignocellulolytic system (symbiont-independent) to enhance the efficiency in wood degradation by fungus-growing termites (Rouland et al. 1988; Rouland 1995).

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No doubt, lignocellulose-feeding insects have to possess at least one of these three lignocellulolytic systems to cope with cellulose-rich diets and most of them use a combination of the above mentioned mechanisms for the efficient digestion of lignocellulosic diets. Indeed, there is no evidence that any lignocellulosefeeding insect is completely independent from symbionts for cellulose degradation. However, it is not always clear which mechanism prevails over another and how well a dual lignocellulolytic system is being coordinated, and complemented one another, on each essential process performed in insect guts. 25.3.3

Insect guts and their microhabitats

The insect guts are the only location for the digestive processing of lignocellulosic food. The diverse gut systems serve as a continuous bioreactor or fermentator, the smallest one in the world, to liberate sugars from various polysaccharides obtained from their food. Despite their small volume of about 0.5–10 ml gut fluid, the hindguts of termites are morphologically complex systems. Indeed, the gut of termites and other cellulose-feeding insects is highly specialized for housing gut microbiota that aid in digestion of cellulose obtain from wood, grass, plant litters, fungi, or other sources, depending on their lifestyle. Thus, the termite gut will be an ideal model to discuss in detail with respect to intestinal structures and associated gut microhabitats. Termite gut structure is an elongated tube differentiated into the foregut, midgut, and hindgut (Fig. 25.9). The fore gut includes the esophagus, crop, and gizzard (muscular proventriculus). The midgut is a simple tube of uniform diameter distally inserted by 4–12 Malpighian tubules (Waller and La Fage 1987; Bignell et al. 1983). All higher termites possess a prominent hindgut which could be further divided into five successive segments: the proctodeal segment (P1), the enteric valve (P2) controlling the entrance of food, the paunch (P3) harboring symbiotic flora, as well as colon (P4) and rectum (P5) (Noirot and NoirotTimothee 1969) (Fig. 25.9). Highly developed the hindgut, at least in xylophagous termites, represents > 40–60% of the termite’s weight (Slaytor et al. 1997). It serves as a reservoir for most of the symbiotic microorganisms (bacteria, fungi and protozoa) and as the primary place of nutrient absorption. However, hindguts among termites, such as wood-feeders, grass-feeders, and fungus-feeders, would vary considerably in shape, size, and the degree of differentiation largely depending on their feeding diets and their gut symbiotic microorganisms. Unlike termite guts, the alimentary tract of woodwasps (immature stage) is a simple tube with no blind sacs or enlarged region in the hindgut that might serve as a fermentation chamber. Thus, the midgut is the largest segment in its gut system and serves as the major site of digestion due to an acquired enzyme mechanism by food ingestion. Similarly, the midgut, a so-called caeca (mycetomes), in the larvae of the cerambycid beetles also serves as the main digestive region with cellulolytic activities, where it also hosts the majority of the


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Fig. 25.9 Morphology of termite guts. (a) Generalized gut in the lower termites. (b) Nasutitermes costalis. (c) Microcerotermes arboreus. (d). soil feeders Cubitermes severus and Procubitermes aburiensis. The division of the hindgut of higher termites into five successive segments follows Noirot and Noirot-Timothée (1969) and Johnson (1979). C: crop; M: Midgut; MX: mixed segment; CE: cecum; P1–P5: proctodeal segments. (from Bignell and Anderson 1980).

symbiotic microbes. The pH of the midgut in termite is about neutral and that of the hindgut is in the range of 6.0–7.5, which are, in general, similar for lower and higher termites although exceptions have been found among soil-feeding higher termites, such as Odontotermes obesus, where pH values could be as high as 10.4–12 (Noirot and Noirot-Timothée 1969; Bignell and Anderson 1980). As another case, the pH values in the midgut of larval scarab beetles are reported to be highly alkaline (pH 11–12) (Wiedemann 1930; Bayon 1980). Indeed, a near neutral pH (6–7.5) is often found in the regions where microbes are found (hindgut or midgut). The termite hindgut, of both higher and lower termites, was assumed to be anaerobic, but it could best be described as an anaerobic gradient system, which is constantly supplied with oxygen via the gut epithelium. Studies using microeletrodes have shown a sophisticated spatial distribution in the termite hindguts with respect to pH and axial or radial gradients of oxygen or hydrogen concentrations (Fig. 25.10, Brune and Kühl 1996; Brune 1998; Schmitt-Wagner and Brune 1999; Brune and Friedrich 2000). Hindgut regions such as the paunch, which are heavily colonized by symbiotic microbiota, appear anaerobic with an Eo usually in range of – 230 to – 270 mV. By contrast, midgut sites appear to be aerobic (especially when not an important site for microbes). The lignin content of the ingested cellulosic diets is particularly important for an efficient cellulose and hemicellulose utilization by insects because it is tightly

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Fig. 25.10 Axial profiles of O2 and H2 partial pressure, and pH in the highly compartmentalized guts of soil-feeding Cubitermes spp. The photograph shows the unraveled gut embedded in agarose, with a microsensor inserted into the P1 region; C, crop; M, midgut; ms, mixed segment; P1–5, proctodeal regions. (from Brune and Friedrich 2000).

associated with cellulose and other plant polysaccharides. Access of cellulases to the cellulose substrate is aided by the physical disruption of lignocellulose by insect mastication (chewing to break plant material into smaller particles) and also by chemical disruption. As discussed by Watanabe and Tokuda (2001), termites grind and crunch their ingested material, which may enhance the amount of surface that can be accessed by the cellulolytic enzymes. The chemical disruption is an oxidative process by lignases (e.g. laccase, peroxidase) requiring molecular oxygen, which is in conflict with the process of microbial fermentation of polysaccharides in the hindgut because high titers of oxygen are harmful to the anaerobic microorganisms, such as cellulolytic protozoa and methanogenic and acetogenic bacteria. However, most insect species that utilize insect-derived cellulases to degrade lignin use one of several strategies to cope with this challenge: 1) have relatively insignificant microbiota in the hindgut, 2) support spatial separation of lignin and cellulose degradation into oxic and anoxic gut regions, respectively, 3) use oxygen consuming bacteria (facultative and strict anaerobic) for maintaining an anoxic microhabitat (Veivers et al. 1982, Adams and Boopathy 2005), and 4) to have an anaerobic gradient system. In many termite species investigated, they use strategies (3) and (4) to create both oxic and anoxic microhabitats in the hindgut. Oxygen penetrates 50–200 mm into the periphery of the hindgut lumen, leaving only the central portion of the dilated compartments anoxic (Brune and Stingl 2005). Cellulosic diets are first chewed by termite mandibles and then continuously ground to a very small particles by the cuticular spines of the proventriculus


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(gizzard in the foregut) prior to entering foregut posterior regions (approximately 20–100 mm in size with average 50 mm for a lower termite, such as C. formosanus) (Yoshimura 1995). These wood fragments generally showed fibrous or chip-like form without noticeable degradation in foregut. Then, wood particles enter the midgut with a partial and noticeable degradation by those cellulolytic enzymes primarily secreted by termite various tissues, such as saliva glands and midgut. Finally, the digesta (particles) are subjected to microbial attack in the hindgut, from where the fermentation products (mainly acetate) are absorbed. Lignin, indeed, is not substantially degraded in most of insect guts, though it may be modified by side-chain oxidation, demethylation of ring methoxyl group, and ring hydroxylation; consequently, the feces of wood-feeding species are characteristically enriched in lignin (Itakura et al. 1995; Hopkins et al. 1998; Geib et al. 2008). This modification of lignin is a critical and necessary process (similar to a so-called "pretreatment" on biomass in industry) for subsequent depolymerization of cellulose by the cellulases in insect guts. Because of the small scale of the gut system in termites (0.5–10 ml gut fluid), the food substrates have to be passed rapidly through their gut systems to obtain adequate energy. The mean retention time of digesta in the hindgut is estimated to be around 24–26 h (Breznak 1984). These facts have significant influences on the composition of the microbial flora and the distribution of cellulolytic flagellates in the lower termite hindguts (Yoshimura 1995; König et al. 2006). It has been proposed that there are four distinct microhabitats in the termite gut: the gut lumen, the surface and the cytoplasm of flagellates, as well as the gut epithelium (König et al. 2006). Many investigations with electron microscopy on termite guts have shown that prokaryotes occur either suspended in the gut fluids, or are located within or on the surface of symbiotic protists, as well as are attached to the gut wall (Breznak and Pankratz 1977). All of these gut prokaryotic symbionts in various gut micro-habitats are considered to play indispensable roles in host nutrition and the lignocellulolytic activities. Further information regarding this subject is available from the recent review work by Ohkuma (2008) and König et al. (2006), and need not be repeated in detail here.

25.4 Cellulolytic symbionts from insect guts and their living environments Digestive symbioses are very common among lignocellulose-feeding insects that may involve both obligate and facultative microbes in their guts. Most gut symbionts are characterized as obligate nutritional mutualisms (cultureindependent) between the host and various microorganisms and belong to three domains: Archaea, Eubacteria, and Eucarya (protozoans and fungi) (Bignell 2006). However, there is a diversity of culture-dependent microorganisms, including bacteria, fungi, and yeast, which reside in insect guts to aid nutrition utilization, cellulose degradation, as well as functioning of the gut microhabitats (König et al. 2006). The microbial flora, either culture-dependent or-independent,

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is not distributed randomly in the gut, but plays certain indispensable roles in distinct microhabitats (Fig. 25.11). Wood-feeding termites to date are the only group of insects whose interactions with intestinal microorganisms has been systematically studied. Thus, in the following discussion, we will mainly focus on termite microbial symbionts that colonize the intestinal tract and are directly involved in lignocellulose digestion regardless of a variety of functions played by other symbionts residing in insect guts.

Fig. 25.11 Example pictures of the symbiotic bacteria and flagellates presented in different microhabitats in the hindgut of a lower termite worker, C. formosanus. (a) bacterial ectosymbiosis on the surface of a flagellate, Pseudotrichonympha grassii, (b) endosymbiosis of Bacteroidales bacteria within a cell of P. grassii (left: a phase-contrast image, right: the FISH image of the endosymbionts in green color), (c) spirochete bacteria as free-swimming symbionts in hindgut fluid, (d) Bacteria residing in the surface of epithelium cells in hindgut, (e) The largest flagellate species, P. grassii, in the hindgut of C. formosanus, (f) main parts of the gut compartments from a worker of C. formosanus. Pictures (a), (c), (d), (e), and (f) are reprinted from Jianzhong Sun unpublished data; and (b) is reprinted from Ohkuma 2008.

25.4.1

Culture-dependent/independent symbionts from insect guts

A variety of microbes, mainly bacteria, have been isolated from insect guts; and they were characterized with various cellulolytic functions in vitro (Paul et al.


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1986; Schäfer et al. 1996; Wenzel et al. 2002; König et al. 2006; Kuhnigk and könig 1997). However, a major of microbial species has not been isolated yet although more than hundreds of microbial strains have been obtained in pure culture, mainly from termite guts. The hindgut is the last, as well as the largest gut compartment in the termite digestive system. The hindgut finally decomposes partially degraded digesta passed from midgut, where the symbiotic microbiota, including culture-dependent and -independent, in hindgut plays the most important role. Although some bacteria colonize the fore- and midgut, the bulk of the intestinal microbiota is found in the hindgut, especially in the paunch, the region immediately posterior to the enteric valve (Fig. 25.9, Breznak and Brune 1994). In lower termites, the ingested wood-particles entering the hindgut are immediately endocytosed (ingestion of wood particles) into their food vacuoles by the cellulolytic protozoa for a substantial degradation. There are no special feeding apparatuses and the ingestion can take place throughout the plasma membrane except the flagellated body surfaces (Kiuchi et al. 2004; Radek 1999). Some ectobiotic bacteria on the surface of protozoa may be engulfed with wood particles and remain attached to the membrane of the digestion vacuole where the wood particles are stored and ultimately degraded (Radek et al. 1992). These anaerobic flagellates (sensitive to oxygen) that occupy a large fraction of the hindgut (1/7–1/3 of total body weight) represent a major source of cellulolytic and xylanolytic activities in termite digestive system (Yoshimura 1995; Brune 2003). By direct microscopic count, the protozoa community are about 1 – 4 ´ 104 per gut (~ 107/ml) in most of lower termite hindguts (Breznak 1984). In general, there are few different flagellate species residing in a single termite’s hindgut (varied from 3–20 species). The different flagellates are functionally and nutritionally specialized and each species may intend to fill a specific niche (Yashimura 1992). The mechanism for a flagellate distribution in the hindgut of a lower termite is not yet known. Given that 60% of the hindgut is oxic to some degrees, flagellate distribution may be related to oxygen tolerance (Brune et al. 1995). A total of ~500 species and subspecies of trichomonad (~175 species), oxymonad (~68 spp.), and hypermastigote (~191 spp.) flagellates (protozoa), within 80 genera, examined from 205 termite species have been reported from the intestines of six families of lower termites (Yamin 1979). Probably many further flagellate species exist, since only about less half of the known species of lower termites have been examined to date (Radek 1999). The majority of the symbiotic protozoa are likely to have cellulolytic activity critical to termite survival. These species seem not to occur elsewhere in nature except in wood-eating roaches of the geneus Cryptocercus. To date, we are unable to obtain axenic cultures of these cellulolytic protozoa, except for three species obtained in culture: Trichomitopsis termopsidis (Yamin 1978, 1980), Trichonympha sphaerica (Yamin 1981) from the termite Zootermopsis sp. and Trichomitus trypanoides from the termite Reticulitermes santonensis (Berchtold et al. 1995). By using autoclaved rumen

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fluid as a supplement in media also containing antibiotics, cellulose, and serum, the flagellate, Trichomitopsis termopsidis, can be cultured (Yamin 1978). The further investigation also indicated that the flagellates did not grow with clarified rumen fluid or with heat-killed cells of several known bacterial species as a supplement. When cellulose was replaced by other carbohydrates no growth occurred, suggesting that cellulose is the main substrate flagellates used for energy. However, whether lower termites obligatorily depend on these flagellates is still arguable (König et al. 2006). Although the lower termites and wood-feeding roaches are believed to depend nutritionally on their protozoa fauna, the precise involvement of each protozoa species and their interactions among the flagellate members are still obscured by the complexity of the protozoa fauna in the hindgut and their non-culturing nature. However, recent advances in selective removal of certain flagellate species from termite hindgut have allowed identification of specific lignocellulosic degradation functions of a protist (Tanaka et al. 2006). For example, when workers of C. formosanus were forced to feed on cellulose substrates with various degrees of polymerization, each protozoa species showed characteristic responses for its utilization. Protozoa, Pseudotrichonympha grassii, could use high DP cellulose, but not low-molecular weight cellulose (LC) as diet. Instead, Holomastigotoides hartmanni and Spirotrichonympha leidyi utilized LC only, but not high DP cellulose (Yoshimura 1995; Tanaka et al. 2006). These results indicate that the degree of polymerization of cellulose might be a major factor for sharing the nutritional sources among various flagellates in the hindgut of C. formosanus and possibly be the same factor for other lower termite species as well. Both the bacteria and protozoa are found in the hindgut of lower termites, but the bacteria usually have a higher density than that of protozoa (e.g. in the termite R. flavipes, 4 ´104 protozoa and 3 ´106 bacteria per termite gut). Prokaryotic symbionts in the guts of lower and higher termites also play a significant role both in lignocellulose digestion and termite nutrition. It has been estimated that the total number of prokaryotes in the hindgut of termites could range between 107 – 1011/ml (Cazemier et al. 1997; Berchtold et al. 1999). Detailed descriptions of the in situ morphology and arrangement of prokaryotes in the hindgut have been made for lower termites (R. flavipes and C. formosanus (Breznak and Pankratz 1977), and higher termites (Nasutitermes exitiosus) (Czolij et al. 1985), Odontotermes formosanus (Yara et al. 1989). However, symbiotic prokaryotes may also be present in the foregut and midgut at different densities depending on host species. For example, the counts of total bacteria for the house cricket, Acheata domestica, are 2.1 ´109, 2.8 ´1010, and 1.9 ´1011 in fore-, mid-, and hindgut, respectively (Cazemier et al. 1997). It seems that most gut bacteria are not directly involved in cellulose degradation in lower termites; while in higher termites, symbiotic gut bacteria are the main decomposers responsible for the degradation of lignocellulose, including various cellulolytic-bacteria both culture-dependent and-independent (O’Brien et al. 1979; Warneche et al. 2007).


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Recently, there has been an increase in the number of studies on the gut microbiota of lignocellulose-feeding insects like termites, with the object of determining their role in the digestive processes of lignocellulose-feeding insects (Wenzel et al. 2002; Cazemier et al. 2003; Adams and Boopathy 2005; König et al. 2006) and their potential uses for generating viable biofuels (Gidh et al. 2006; Warneche et al. 2007; Scharf and Tartar 2008; Sun 2008; Sun and Scharf 2010). The efforts involved in isolating culture-dependent microbes from termite guts have been well reviewed by Varma et al. (1994), Radek (1999), and König et al. (2006). To date, cellulolytic bacteria have been often recovered from the lower and higher termites. The bacteria groups obtained in pure culture primarily belong to the branch of Gram positive bacteria, proteobacteria, as well as Bacteroides, Fusobacterium branch. These culture-dependent bacteria are believed more or less to be involved in the degradation of cellulose, hemicellulose, and aromatic compounds as well as nitrogen fixation. A new facultative bacterial species, Promicromonospora pachnodae sp. nov., with a capability of degrading both cellulose and xylose was successfully isolated from the larvae of the rose chafer (a scarab beetle), Pachnoda marginata, (Cazemier et al. 2003). Cellulolytic activities have also been demonstrated for some bacterial strains isolated from the lower termite hindguts (Wenzel et al. 2002), indicating that bacteria may play a role in cellulose digestion in addition to the cellulolytic flagellates. Other than culture-dependent microorganisms, by using 16S rDNA approach, a diversity of non-cultivable bacteria that mainly belong to the spriochetes, proteobactia, Gram positives and Bacteroides/Flavobacterium branch have also been identified from termite guts (Berchtold et al. 1999; Berchtold and König 1996; Ohkuma et al. 1999; Tokuda et al. 2000; Watanabe et al. 2003; Husseneder et al. 2005; Fisher and Brewste 2007). Recently, the most astonishing progress has been made by metagenomic research on a higher termite hindgut, Nasutitermes sp., which revealed a broad diversity of bacteria that included 12 phyla and 216 phylotypes, approximately 250 bacterial species (Warneche et al. 2007). As a special group of symbiotic bacteria characterized with the filamentous and Gram-positive, actinomycetes were also isolated from the hindguts of some higher termites, Macrotermes, Armitermes, Odontotermes, Microcerotermes spp, Procubitermes aburiensis, and Cubitermes severus, which would grow on cellulosic substrates with a very high cellulolytic activity (Pasti and Belli 1985; Bignell et al. 1991). The isolated gut actinomycetes (such as Streptomyces sp., Micromonospora sp. and Thermomonopora sp.) from these termites have been reported to produce active cellulases. Their endoglucanase levels were indeed higher than those found in Trichoderma reesei cellulase complex (Pasti and Belli 1985). But inhibition by glucose was a common feature for almost all isolates identified. Most of Archaea identified from lower and higher termite guts are anaerobic and culture independent, which belong to the families Methanobacteriaceae,

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Methanosarcinaceae, and Methanomicrobiales (a so-called methanogenic bacteria) (Breznak and Brune 1994; Braumann et al. 2001; Tokuda et al. 2000; Deevong et al. 2004). These symbionts mainly function in methanogenic metabolisms (fermentative process using CO2 and the methyl group of acetate as electronic acceptors) (Breznak and Brune 1994) and may reside within the protozoa cells as an endosymbiotic association (Ohkuma et al. 1999; Hara et al. 2004; Ohkuma 2008). A diversity of yeasts occurs in the gut of the lower termites Mastotermes darwinensis, Zootermopsis angusticollis, Z. nevadensis, Reticulitermes santonensis, Heterotermes indicola, and the roach Cryptocercus punctulatus (Prillinger et al. 1996). Most of the yeast isolates belong to the genera Candida, Cryptococus, Debaryomyces, Pichia and Sporothrix (König et al. 2006). Some yeast isolates can degrade cellulose and xylan (Schäfer et al. 1996; Wenzel et al. 2002). In addition, 14 various yeasts were also isolated from the gut of a variety of insects, including beetles, lacewings, fish flies, crane flies, and a cockroach. These yeasts were identified as Candida membranifaciens, C. tenuis, Pichia nakazawae, and nine un-described taxa in Saccharomycotina (Suh et al. 2005). Another investigation by the same authors also indicated that the yeast, Enteroramus dimorphus, from the gut of the passalid beetle, Odontotaenius disjunctus, demonstrated a cellulolytic activity on xylose (Suh et al. 2004). It is also important to note that some aerobic bacteria isolated from the hindgut of xylophagous termites, including the lower termites, Mastotermes darwiniensis, Reticulitermes santonensis, Zootermopsis angusticollis and higher termite, Nasutitermes nigriceps would potentially possess a capability of degrading dimeric lignin model compounds (Kuhnigk et al. 1994; Kuhnigk and König 1997; Harazono et al. 2003). The bacterial strains that utilize lignin model compounds as substrates were identified as Bacillus sp. Acinetobacter sp., Alcaligenes sp., Comamonas sp., Nocardia sp., Ochrobactrum sp., Pseudomonas sp., and Streptomyces sp. These findings indicate that the hindgut symbiotic bacteria of termites are potentially able to utilize the aromatic extractives from wood when oxygen titer in the specific gut habitat is satisfied (oxygen is required for the cleavage of the aromatic rings). 25.4.2

Lignocellulolytic microbes from insect nest environments

The symbiotic associations between insects and microorganisms are also observed outside of the termite gut digestive systems. The fungi are a special group of symbionts often used by many insects to help them digest lignocellulosic diets through the unique exosymbiotic relationships established between host insects and the fungi that are cultivated in their nests and the associated environments (Martin 1987; Wood and Thommas 1989; RoulandLefèvre 2000; Rouland-Lefèvre et al. 2006). As we have mentioned in the previous discussion, these symbiotic fungi are mostly obligated and exclusively found in their nests, such as Termitomyces. Indeed, symbiotic relationships with


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fungi have been observed in many groups of insects (Kendrick 1991), including scale insects (Hemiptera), gall midges (Diptera), wood-wasps (Hymenoptera), beetles (Coleoptera), aphids (Homoptera), and ants (Hymenoptera). Among these insects, the fungus-growing ants/wood-wasps/termites are well known using a sophisticated exosymbiotic association with fungi to cope with the recalcitrant cellulose-rich diets and thus demonstrated a major impact on tropical ecosystems (Wood and Thommas 1989; Rouland-Lefèvre 2000). Most of these fungi utilized by wood-feeding insects, such as termites, woodwasps, and leaf-cutting ants, belong to Basidiomycetes that make cellulosic biomass easily digestible by insect hosts. The symbiotic association between each of these insect groups and the ‘fungus-garden’ has been discussed early and need not be repeated here. However, it is noteworthy to emphasize that symbiotic fungi may play different roles in the processing of wood degradation among different fungus-growing insects. For example, in Macrotermes spp., the main role of symbiotic fungi is to degrade lignin, so that termites can utilize cellulose more efficiently, whereas in Odontotermes spp., Hypotermes makhamensis, Ancistrotermes pakistanicus, and Pseudacanthotermes militaris, the fungi are to serve as a food source (Hyodo et al. 2003). Termite soils (the soils from termite nest environments) provide a very distinct ecological environment which harbors and promotes very specialized cellulolytic and hemicellulolytic microorganisms. It has been reported that the cellulolytic microbiota are uniquely rich in the termite nests, mounts (hill), as well as other environments associated with their living habitats (i.e. termite body surface, infested wood, etc.), which include bacteria, actinomycetes, protists, as well as fungi that are more numerous than all other groups of microbes (non-cellulolytic) examined and more abundant in mound/nest soils than in adjacent soils (Keya et al. 1982; Paul et al. 1985; Paul and Varma 1992; Varma et al. 1994; Kumari et al. 2006). The estimates of microbial population in termite-infested soil mounds in Kenya (Africa) showed that bacteria and actinomyces were in the most abundant during the wet season with the density up to 106 and 105/g of dry soil for bacteria and actinomyces, respectively (Keya et al. 1982). The buildup of large numbers of specialized microbes in an ecosystem plays a vital role directly or indirectly in maintaining symbiotic balances and regulates the ecology of microorganisms. In another investigation, a strain of Bacillus sp. isolated from termite infested soil demonstrated a broad spectrum of enzymes responsible for the degradation of cellulosic and hemicellulosic components of agricultural residues (Paul and Varma 1992). To date, more than one hundred strains of various cellulolytic microbes (bacteria and fungi) have been isolated from various termite habitats, including termite mounds/nests (Zoberi 1979; Paul and Varma 1990; 1993; Sarkar 1991), termite body surfaces (Zoberi and Grace 1990), and termite infested wood (Paul and Varma 1993). Indeed, these unique habitats may potentially serve as a pool for screening novel cellulolytic microbes.

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25.5


Insect lignocellulases

Through photosynthesis, plants convert solar energy into thermochemical energy and store them in the form of carbohydrates (lignocelluloses), the universal fuel and building blocks for all living organisms. To exploit the stored energy efficiently, cellulolytic insects have evolved different digestion systems and strategies including specific lignocellulases. In the following section, we will review 1) enzymatic machineries which can depolymerize cellulose, hemicellulose and lignin, respectively; 2) distributions of these highly specialized enzymes, 3) major strategies that are deployed by insects to exploit lignocelluloses, and 4) recent progress on insect lignocellulose degradation through genomic and metagenomic research. 25.5.1

Lignocellulose degradation machinery

Lignin As we have discussed, the heterogeneity and stereoirregularity of lignin polymers protect cellulose and hemicellulose from chemical and/or enzymatic degradations (Reid 1989). Clearly, lignin has been the first layer of defense for the plant cell wall and the toughest substrate for either thermal-chemical or enzymatic pretreatments (Higuchi 1990; Lewis and Sarkanen 1998; Cullen and Kersten 2004; Rubin 2008). No organisms are known to utilize lignin as their sole carbon and energy source, although a few microbial taxa such as white rot fungi and cellulolytic insects develop systems to break down lignin (Cullen and Kersten 2004). For these organisms, however, depolymerization of non-carbohydrate lignin polymers is the critical first step to gain access to the carbohydrate polymers (hemicelluloses and celluloses) to eventually release thermal-chemical energies stored in the lignocellulosic biomass. Unlike hemicelluloses and celluloses, lignin degradation requires an aerobic oxidation process rather than hydrolysis. The lignin degradation machinery in insects is mainly composed of oxidative enzymes such as peroxidases, oxidases, and laccases. In addition, a suite of auxiliary enzymes may contribute to the complete mineralization of monomers to CO2 and H2O and to the generation of secondary metabolites. These supporting casts include enzymes typically involved in the xenobiotic detoxification and aging process (e. g. antioxidants). For example, alcohol dehydrogenase, catalase, superoxide dismutase, methyltransferases, cytochrome P450, cytochrome P450 co-factor, epoxide hydrolase, reductase, glutathione-S-transferase, and esterase (Matsumura 1986; Cullen and Kersten 2004; Feyereisen 2005; Zhou et al. 2006; Scharf and Tartar 2008; Table 25.4). Ohkuma (2003) believes that oxidative nature of lignin degradation is likely one of the evolutionary origins for the degrading pathway of aromatic xenobiotics and/or environmental pollutants. Polychlorinated biphenyl (PCB) is a notoriously persistent organic pollutant which bioaccumulates in animals. In search of a biodegrading agent of PCB,

Utilization of Lignocellulose-feeding Insects for Viable Biofuels Table 25.4

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Lignocellulose degradation machinery in insects

Degradation Machinery Lignin

Lignocellulose Components Hemicellulose

Cellulose

endo-β-1,4-xylanases exo-β-1,4-xylanases β-xylosidases

endo-β-1,4-glucanases exo-β-1,4-glucanases β-glucosidases

Secondary alcohol dehydrogenase catalase Enzymes superoxide dismutase methyltransferases cytochrome P450 epoxide hydrolase reductase glutathione-S-transferase esterase

α-arabinofuranosidases α-glucuronidases acetylxylan esterase ferulic acid esterase p-coumaric acid esterase

NA

Degradation oxidation

hydrolysis

hydrolysis

Primary Enzymes

peroxidases oxidases laccases

Chung et al. (1994) isolated an aerobic, non-sporulating bacteria, Rhodococcuserythropolis, from a lower termite, Reticulitermessperatus. This particular bacteria species and other strains from the same Rhodococcus genus effectively metabolized aromatic organic pollutants, suggesting a potential in bioconversion and bioremediation of hazardous waste materials (Maeda et al. 1995; Kosono et al. 1997). Hemicellulose As a heteropolymer of pentoses (xylose, arabinose), hexoses (mannose, glucose, galactose), and sugar acids, hemicellulose is the second barrier protecting cellulose from enzymatic depolymerization. Complete biodegradation of hemicelluloses requires the combined efforts of endo-β-1, 4-xylanases, exo-β1, 4-xylanases, β-xylosidases, and several accessory enzymes, including αarabinofuranosidases, α-glucuronidases, acetylxylan esterase, ferulic acid esterase, and p-coumaric acid esterase (Saha 2003; Table 25.1). Specifically, conversions accomplished by each of these enzymes are: (1) endo-xylanases hydrolyze the β-1, 4-xylose linkages in the xylan backbone, (2) exo-xylanases hydrolyze reduced β1, 4-xylan linkages releasing xylobiose, (3) β-xylosidases act on xylobiose to liberate xylose and other short chain oligosaccharides, (4) α-arabinofuranosidases hydrolyze terminal non-reducing α-arabinofuranose from arabinoxylans, (5) αuronidases release α-glucuronic, -mannuronic and-galacturonic acids, and (6) esterases hydrolyze phenolic ester linkages between xylose units of the xylan and acetic acid or between arabinose side chain residues and phenolic acids, such as ferulic acid and p-coumaric acid (Sara 2003; Scharf and Tartar 2008; see Fig. 25.3). Cellulose Cellulose depolymerization requires the collaborative action of three soluble extracellular enzymes including endo-β-1, 4-glucanases, exo-β-1, 4-glucanases, and β-glucosidases (Breznak and Brune 1994; Ragauskas et al. 2006b; Table 25.4). Specifically, (1) endoglucanases internally hydrolyze β-1, 4-glucosyl linkages along the primary cellulose backbone to release shorter oligomers (4-5 carbon sugars) and/or glucose, (2) exoglucanases or “cellobiohydrolases” target

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the terminal regions of polymeric chains to liberate either glucose or cellobiosyl units, and (3) β-glucosidases, then, hydrolyze cellobiose and cellotriose to liberate glucose monomers. Liberation of glucose from cellulose appears to require synergistic collaboration by three groups of cellulases (Breznak and Brune 1994; Zhou et al. 2007). 25.5.2

Distribution of lignocellulose degradation machinery

Generally, there are two major factors to determine the distribution pattern of lignocellulose degradation machinery: 1) the structural and physical properties of three major polymer components of lignocelluloses, lignin, hemicelluloses, and cellulose, and 2) the physiochemical conditions of the microhabitats of an insect digestive tract. The degradation and fermentation of carbohydratic celluloses and hemicelluloses require anaerobic hydrolysis, and consequently the hydrolytic cellulose and hemicelluloses degradation machineries are likely confined to the anaerobic section of insect digestive tract (hindgut; Fig. 25.12). The hindgut

Fig. 25.12 A schematic drawing of insect lignocellulose degradation processes. Photo (top) showing a mushroom-blooming fungus comb of the termite Odontotermes sp. in Thailand, and (bottom) different regions of the digestive tract of a lower termite, R. flavipes, SG (salivary gland); FG (foregut); MG (midgut); HG (hindgut); and FC (fermentation chamber). Fungus photo is reprinted from Ohkuma 2003; Termite gut photo was taken by J.A. Smith.


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consists of a fermentation chamber or “paunch” that is generally anaerobic, but does possess a micro-oxic zone (oxygen sink) around its periphery (Brune et al. 1995; Breznak and Brune 1994; Leadbetter et al. 1999). The hindgut houses the majority of gut endosymbionts, and is the location where most cellulose and hemicellulose degradation occurs, as well as fermentation (Scharf and Tartar 2008). In contrast, the disruption of lignin matrix requires aerobic oxidation, thus the degradation of lignin likely takes place in aerobic environments, such as oxic section of insect digestive tract (e.g. foregut, Fig. 25.12). In case of fungusgrowing insects, lignin degradation likely occurs inside the fungal exosymbionts rather than the digestive tract of host insects Fig. 25.12). 25.5.3

Strategies to exploit Lignocellulases

Although cellulolytic insects account for only a small fraction of the insect community, they have tremendous impacts on our economy and ecosystem (Su and Scheffrahn 2002; Sugimoto et al. 2000; Ohkuma 2003). The intimacy between cellulolytic insects and their endo- or exo-symbionts has been a text book example of symbiosis/mutualism, and these highly specific microbial floras are critical for the survival of insect hosts because of their cellulolytic capacity. In previous section (3.2), we discussed three lignocellulolytic systems in insects: symbiont-dependent, symbiont-independent, and acquired enzyme system. Here, we further introduce the four possible strategies that insects have developed from these lignocellulolytic systems to maximize their lignocellulolytic capability: (i) exploitation of eukaryotic endosymbionts in the hindgut microbiota; (ii) exploitation of prokaryotic endosymbionts in the hindgut microbiota; (iii) exploitation of eukaryotic exosymbionts; and (iv) utilization of endogenous lignocellulases secreted from insect tissues (Fig. 25.12 and 25.13). Exploitation of eukaryotic endosymbionts in the hindgut microbiota (protozoa origin) The remarkable mutual symbiosis between termites and their eukaryotic symbionts have recently been traced back to at least 100 million years ago. The discovery by Poinar (2009) in an Early Cretaceous Burmese amber that contained a new termite species, Kalotermes burmensis n. sp., and its 100 million-year old hindgut residents, flagellate protists, is the oldest example of "mutualism" ever discovered between an animal and microorganism. Eukaryotic endosymbionts from insect hindguts has long been considered as the primary source of lignocellulases (Cleveland 1923, 1924; Sugimoto et al. 2000) until the validation of existence of endogenous insect cellulases in termites (Watanabe et al. 1998). The representative insect species utilizing this strategy are the eusocial lower termites, e. g. Reticulitermes flavipes (Rhinotermitidae) (Cleveland 1924; Zhou et al. 2007) and subsocial wood-eating cockroach, Cryptocercus punctulatus (Cryptocercidae) (Cleveland 1934). These cellulolytic symbionts are anaerobic eukaryotes from specific genera of flagellates restricted to the hindguts of the lower termites and wood roaches (see 4.1 section for details).

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Fig. 25.13 Strategies to exploit lignocellulases by wood-feeding termites. Termites have developed all four degradation strategies to exploit the utility of lignocelluloses. Phylogenetically termites have seven families in which six of them belong to the lower termites (harboring cellulolytic flagellates in hindgut), and the other one belongs to the higher termites (harboring cellulolytic bacteria in hindgut). Lower termites rely on the combination of lignocellulases from the termite and eukaryotic endosymbionts that reside in their hindguts. Higher termites (Termitidae), account for 75% of termite species, have seven subfamilies in total. Higher termites utilize lignocellulases from the termite and prokaryotic endosymbionts that reside in their hindguts. However, in the subfamily Macrotermitinae, these termites have developed a unique mutualistic relationship with eukaryotic exosymbionts that reside in the vicinity of their nest to assist their lignocellulose degradation, especially with the lignin degradation.

Exploitation of prokaryotic endosymbionts in the hindgut microbiota (bacterialorigins) Higher termites which harbor diverse prokaryotic symbionts without cellulosedegrading eukaryotic protozoan in their hindguts are the main examples of this strategy. A recent metagenomic survey of an arboreal Nasutitermes species phylogenetically related to N. ephratae and N. corniger revealed an astonishing array of genes and gene fragments ( > 200) involved in cellulose and/or hemicellulose hydrolysis from 45 glycosyl hydrolase families (GHFs) (Warnecke et al. 2007). It is worth noting that no lignases were identified in this dataset,


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supporting the contention that lignin degradation are likely contributed by exosymbiont and/or insect-derived enzymes rather than endosymbionts (Scharf and Tartar 2008). Hindgut prokaryotic endosymbionts have been shown to contribute to cellulose digestion in several other cellulolytic insects including the American cockroach, Periplaneta americana (Blattidae) (Bignell 1977; Cruden and Markovetz 1979), the rhinoceros beetle, Oryctes nasicornis (Scarabaeidae) (Bayon 1981), and an aquatic crane fly, Tipula abdominalis (Tipulidae) (Sinsabaugh et al. 1985; Griffiths and Cheshire1987; Cook et al. 2007). Exploitation of eukaryotic exosymbionts (fungal-origins) The fungus-growing insects are another fascinating yet classical example of mutual symbiosis, and belong to the acquired enzyme system (see section 3.2. for details). This well-developed collaboration between insect hosts and their exosymbiotic fungi enables efficient degradation of lignocelluloses (Hinze et al. 2002). Fungus-growing termites belong to a unique subfamily of higher termites (Termitidae: Macrotermitinae; Fig. 25.13). With the help of their basidiomycete fungi from the genus Termitomyces (Agaricales, Tricholomataceae), these termites (e.g. Termitomyces albuminosus) can consume more than 90% of the dry wood in some arid tropical areas (Abe et al. 2000). This exosymbiontmediated lignocellulose degradation strategy has also been shown in other species of fungus-growing termites, such as Macrotermes natalensis (Martin and Martin 1978) and Macrotermes mulleri (Rouland et al. 1988), the wood-boring larvae of siricid woodwasp, Sirex cyaneus (Siricidae) (Kukor and Martin 1983), the wood-boring larvae of five species of cerambycid beetles (Kukor and Martin1986; Kukor et al. 1988), and the detritus-feeding nymphs of stonefly (Sinsabaugh et al. 1985; Martin1992). Utilization of endogenous lignocellulases secreted from insect tissues (insectorigin) The fourth and last strategy has nothing to do with symbionts but the insects themselves. Treves and Martin (1994) suggested that symbiont-independent (insect-derived) cellulose digestion system is a primitive trait whereas symbioticdependent system is a derived condition. Molecular analyses of an endogenous endoglucanase from many cellulolytic insects validate the contention (Watanabe et al. 1998; Tokuda et al. 1999; Watanabe and Tokuda 2001; Nakashima et al. 2002a; Zhou et al. 2007). This evolutionarily conserved endogenous endoglucanase belongs to the glycosyl hydrolase family 9 (GHF9). GHF-9 homologues have been identified in lower termites, higher termites, cockroaches, beetles, and crayfish, suggesting the presence of an ancestral enzyme before the symbiotic relationship with gut protists was established in termites (Lo et al. 2000). A recent survey on the endogenous lignocellulases from a lower termite, R. flavipes, identified over 100 genes and gene fragments involved in lignin oxidation, hemicellulose and cellulose hydrolysis from 21 glycosyl hydrolase families (GHFs) (Tartar et al. unpublished). The complexity and diversity of insectderived lignocellulases suggest the importance of symbiont-independent system in the insect lignocellulose degradation process.

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25.5.4


Genomics and metagenomics: a gateway to insect lignocellulase research

From the previous sections, we now know the general processes and complexity of lignocellulose degradation occur in the lignocellulose-feeding insects. Traditional “forward genetic” research can only provide a snapshot of lignocellulose degradation, whereas, current “reverse-genetic” approach can offer a global view through genomics and metagenomics analyses. In the following section, we summarize the latest progress made in genomic and metagenomic research on wood-feeding termites. Termites are the most efficient cellulose degraders with the assimilation rate up to 99%, whereas other cellulolytic insects vary from 10%–90% (Martin 1991 and references therein). This suggests that wood-feeding termites have developed a unique yet highly efficient cellulolytic system for the utilization of lignocellulosic materials. All the lignocellulolytic systems and strategies we have discussed in proceeding sections can be found in various types of wood-feeding termites (Fig. 25.13), which make them the ideal model systems for the study of lignocellulose digestion in insects. Metagenomics has been defined as the genomic analysis of microorganisms by direct extraction and cloning of DNA from an assemblage of microorganism (Handelsman 2004), such as from an insect hindgut. The intrigue associated with metagenomics is that it allows us to study the genomic material of many individual species simultaneously, without having to first do a culturing step (Nelson 2008). In the real world, we can only culture 1% of the microbial species in nature (Aman et al. 1995) and the majority of symbionts presented in the digestive tracts of lignocellulose-feeding insects are indeed culture independent. To date, there have been three examples of genomic and metagenomic research in wood-feeding insects: (i) meta-transcriptome sequencing of hindgut symbionts from a lower termite, R. speratus (Todaka et al. 2007); (ii) meta-transcriptome sequencing of termite digestive tract and hindgut symbionts, respectively, from a lower termite, R.flavipes (Zhou et al. 2007; Tartar et al. unpublished); and (iii) metagenome sequencing of hindgut symbionts from a higher termite, an unidentified species of Nasutitermes (Warnecke et al. 2007). Enzymes responsible for hydrolyzing carbohydrates are generally referred to as glycosyl hydrolases (GH). GH enzymes play important roles in plant cell wall metabolism, the biosynthesis of glycans, plant defense, signaling, and the mobilization of storage reserves (Minic 2008 and references therein). To date, GH enzymes have been classified into 113 glycosyl hydrolase families (GHFs) according to the Carbohydrate-Active Enzymes (CAZy) database (URL: http:// www.cazy.org/) (Cantarel et al. 2009). Because hemicelluloses and celluloses are bona fide carbohydrates, a wealth of information on GHFs is starting to come out following the genomic and metagenomic analyses of lignocellulose degradation in termites (Zhou et al. 2007; Warneche et al. 2007; Tartar et al. unpublished). Although R. flavipes and R. speratus datasets are partial metagenomic sequencing (i.e. ESTs), a comparison with the complete metagenome data from Nasutitermes


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reveals some interesting but serendipitous results. Firstly, at transcriptome level, 45.3% of GHFs were bacterial specific whereas only 3.8% were protozoa specific and 7.5% were termite specific (Fig. 25.14). This result supports the contention that termite guts are reservoirs for novel lignocellulases, and it is especially true for the rarely studied prokaryotic symbionts in higher termites.

Fig. 25.14 Summary of glucosyl hydrolase families (GHF) identified from different termite genomes and symbiont metagenomes. We compared 45 GHFs from the Nasutitermes bacterial symbionts, combined 20 GHFs from the Reticulitermes protozoan symbionts(R. speratus and R. flavipes), and 24 GHFs from R. flavipes digestive tract without symbionts. “Specific GHFs” denote GHFs only appeared once among three databases; “ubiquitous GHFs” refer to GHFs well represented in three databases; and “other GHFs” represent GHFs appeared in two databases.

Secondly, current annotation of GHFs was based on the sequence similarity (homology) with the reference genome/metagenome at the primary structure level. However, similar primary structures do not warrant similar functions. Because the fold of proteins (secondary or tertiary structures) is better conserved than their amino acid sequences (primary structure), some of the GHFs now are grouped into GH 'clans' to better predict their potential functions. Among the 53 GHFs identified from three termite species, 23 did not fall into any existing GH clans, whereas the remaining 30 GHFs have been categorized into 11 known GH clans: GH-A (11 GHFs: 1, 2, 5, 10, 26, 30, 35, 39, 42, 51, and 53), B (2 GHFs: 7 and 16), C (1 GHF: 11), D (3 GHFs: 27, 31, and 36), F (2 GHFs: 43 and 62), G (1 GHF: 37), H (3 GHFs: 13, 70, and 77), K (3 GHFs: 18, 20, and 85), L (1 GHF: 65), M (1 GHF: 8), and N (1 GHF: 28)(Fig. 25.15). The three GH clans lacking of termite representatives are GH-E, I, AND J. Such paucity, however, could be the result of incomplete sequencing in the lower termite R. flavipes and R. speratus. All these GH clans correspond to single or multi-hydrolytic function(s). A more

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Fig. 25.15 Summary of glucosyl hydrolase clans identified from different termite genomes and symbiont metagenomes. A total of 53 GHFs from both lower and higher termites were categorized into the GH clan based on their structural (protein folding) similarities. “Others” denotes GHFs cannot be categorized into any existing clans. The potential biological functions of various families and clans are described in details at the CAZy database (http://www.cazy.org/).

thorough investigation of these GH clans might shed light on the interactions between termites and symbiont cellulolytic systems. Finally, no lignases have been identified from either eu- or prokaryotic symbiont metagenomes. In contrast, lignin degradation machinery (e.g. laccases and peroxidases) has been found in the termite gut library (Tartar et al. unpublished). These results are congruent with the hypotheses that 1) oxidative degradation of lignin must first be degraded and/or at least partially depolymerized before cellulose and hemicellulose digestion carries out, and 2) lignin degradation likely takes place in the aerobic foregut and/or midgut using termite-derived lignases.

25.6

Termite hydrogen production and its potential

Of the metabolic reactions occurring in lower termite hindguts, molecular hydrogen (H2) is constantly produced by the fermentation process, either as a byproduct emitted to the atmosphere or recycled within the hindgut by prokaryotes (Kawaguchi et al. 2006; König et al. 2006; Cao et al. 2010). The metabolic hydrogen in termite guts, so-called biological hydrogen (biohydrogen), is converted from lignocellulosic biomass by symbiotic protozoa. It represents a unique mechanism distinct from other biohydrogen systems currently under evaluation, such as light-driven processes by green algae or cyanobacteria, and dark fermentation by anaerobic bacteria (Hallenbeck and Benemann 2002). Since


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biological H2 emission is extremely limited in natural ecosystems, H2 production that converts from the cellulosic diets by insects is very valuable. As we have known, hydrogen is widely recognized as a clean and efficient energy resource (Abraham 2002; Hoffmann 2001) and has the highest energy content per unit weight of any known fuel (143 GJ/tonne) (Boytles 1984). Therefore, the interests have been renewed in recent years to search the advanced ecosystems/ mechanisms to produce biohydrogen derived from a renewable source, such as lignocellulosic biomass. The potential value of biohydrogen system in the hindgut of wood-feeding termites would not have been recognized until recently due to the fact that hydrogen is one of the degradation products from lignocellulose (Carey and Adam 2006; U.S. DOE 2006; Kawaguchi et al. 2006; Pest and Brune 2007; Sun 2008; Cao et al. 2010). Thus, there is a need to have a brief discussion on termite gut H2 and its potential for future hydrogen economy. 25.6.1

Capability of energy gas emission (H2 and CH4)

It has been reported that termites may emit large quantities of H2 and CH4, the two types of energy gases, into atmosphere and the global annual emissions from their guts calculated from laboratory measurements are strikingly high with 1.51014 grams of methane (CH4) and 21014 grams of molecular hydrogen (H2) (Zimmerman et al. 1982; König et al. 2006). In lower termites, the wood particles ingested by termite workers are first depolymerized into sugars (pentoses and hexoses) and then the various sugars within the protozoan cells will be primarily fermented to H2, CO2, CH4, and acetate by diverse prokaryotes (Yamin 1981; Tsunoda et al. 1993; Zimmerman et al. 1984, 1982; Inoue et al. 2007). Although some symbiotic bacteria may also play a positive role in producing hydrogen (Taguchi et al. 1992, 1993), cellulolytic protists are considered to be the major producers of H2 in the termite gut where H2 partial pressures are the highest found (Pester and Brune 2007). However, molecular H2 produced by cellulolytic protozoa in the hindgut may serve as an intermediate product because it could be consumed by gut prokaryotes/Archaea, particularly acetogens and methanogens (both strictly anaerobic). Both metabolic reactions catalyzed by these symbionts in the hindgut are denoted as follows (Breznak and Switzer 1986; Slaytor 2006): Acetogenesis: 4H2+ 2CO2® CH3COOH + 2H2O (Acetate is utilized by termites) Methanogenesis: 4H2+ CO2® CH4+ 2H2O (CH4 is mainly emitted to the atmosphere) Thus, hydrogen emission from termite guts is postulated to be insignificant due to the fact that molecular H2 is largely transferred from H2-producing microorganisms to H2-consumers, such as acetogens or methanogens within the termite’s hindgut. Although an unknown fraction of the H2 produced by termites may be used for the metabolic reactions listed above, many

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investigations have often reported a significant H2 production released from a variety of lower termite species as a byproduct of the wood digestion (Yamin 1981; Odelson and Breznak 1985; Ebert and Brune 1997; Sugimoto et al. 1998; Inoue et al. 2007; Kawaguchi et al. 2006; Pester and Brune 2007; Cao et al. 2010), where the emission rates could vary from 122–5883.3 mmol/h/g termites. An intrinsic capability of hydrogen production in termite hindgut depends on termite species and it can be ranged from 15 to 470 nmol H2 per hour per termite gut (prior to being emitted to the atmosphere), which corresponds up to 34 liters of hydrogen per liter of termite gut content each day (Pester 2006). The hydrogen partial pressure in the termite hindgut could reach a maximum of 72.2 28.5 (kPa) (Pester and Brune 2007). Stoichiometrically, the evaluation of H2 gas from termites is equivalent to approximately 10% of the CO2 they produce (Zimmerman et al. 1982). In higher termites, the sources of hydrogen are not known but maybe due to lack of flagellated symbionts. The amount of molecular hydrogen to be emitted to the atmosphere by termites depends on the termite species being investigated, regardless of how much of the total H2 production generated within termite hindgut. As pointed out by Pester (2006), almost no molecular hydrogen was lost by emission from the termite guts for the three lower termite species being evaluated, e. g. Reticulitermes santonensis, Zootermopsis nevadensis, and Cryptotermes secundus, which indicated an efficient recycling by prokaryotes/Archaea within hindgut. However, in another group of lower termites Coptotermes formosanus, Reticulitermes flavipes and Reticulitermes virginicus, the rates of hydrogen emission were estimated at 1.37 0.15, 3.07 0.23, and 4.78 0.15 mmol /h/g body weight, respectively (Cao et al. 2010), implying that these termite species possess a relatively low levels of acetogenesis and methanogenesis. In addition to hydrogen emission, methane is emitted from lower termite guts at same time and often reported from some higher termite species as well, such as fungus-growing and soil-feeding termites (Zimmerman et al. 1982; Tsunoda et al. 1993; Sugimoto et al. 1998; Schmitt-Wagner and Brune 1999; Kawaguchi et al. 2006; Slaytor 2006; Cao et al. 2010). Methane emission rate from a termite gut may be recorded as equal, higher, or lower if compared to that of hydrogen emission from same termite species gut. The pronounced differences in the degree of hydrogen accumulation in termite hindgut and the relative amounts of methane/hydrogen emission to the atmosphere are often demonstrated in various lower termite species tested (Sugimoto et al. 1998). Recently, three distinct emission patterns representing three lower termite species were reported by Cao et al. (2010) using a modified CH4/H2 index, which we can use to approximate the relative potential for energy gas emission by a termite species (Fig. 25.16). A termite species with a greater negative index is desirable for H2 production system, reflecting a low level of homoacetogenesis and methanogenesis in hindgut. It also has been reported that, in wood-feeding lower termites, methanogenesis activity is outcompeted by acetogenesis from CO2 and H2 (Breznak 1994),


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Fig. 25.16 Comparison of the profiles of H2 and CH4 emission with the modified index of CH4/H2 for R. flavipes, R. virginicus and C. formosanus during the period of 72 h incubation. The index values varied from negative to positive and were defined as: more CH4 emission was measured than H2 when index was > 1 (denoted as CH4 > H2); CH4 emission was equal to H2 when index was 1 (denoted as CH4 = H2); less CH4 was produced than H2 when index was from 0 to 1 (denoted as CH4 < H2); and only H2 was detected when index was negative (denoted as H2 only).

whereas in the fungus-growing and soil-feeding termites, methanogenesis activity normally outcompetes acetogenesis (Brauman et al. 1992). As the consequences of these differences at least partially resulting from their various feeding diets, the hydrogen emission from wood-feeding termites may demonstrate a higher emission rate than that from higher termite guts. On the other hand, methane emission is predominant in the fungus-growing and soilfeeding termites (Slaytor 2006). It is also estimated that an intact field nest of fungus growing termites Macrotermes jeanneli would emit 0.5–1 liter of CH4 per day to the atmosphere (Darlington et al. 1997). In addition to termites, hydrogen emission were also reported in other groups of insects, including cockroaches (Blaberus sp., Blattella sp., Gromphdorhina sp., Leucophaea sp., Periplaneta sp. and Pycnoscelus sp.), beetles (Dynastes sp., Pachnoda sp., Cetonia sp., Dicronorrhina sp., Eudicella sp., Geotrupes sp., Leptinotarsa sp., Phaedon sp.), weevils (Otiorrhynchus sp), Caterpillars (Aphomia sp., Danaus sp.), and Flies (Tipula sp.) (Hackstein and Stumm 1994). Methane emission was also simultaneously detected from most of these insects tested and the emission rates varied from 46–255 mmol /h/g body weight (Hackstein and Stumm 1994). But, termites are a unique group of insects and the highest methane emission rate could be recorded at 380 317 mmol /h/g body weight of termites (Hackstein and Stumm 1994).

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25.6.2


Gut symbionts for hydrogen evolution and the mechanism insights

The cellulolytic flagellated protozoa in lower termites are mainly responsible for digesting cellulose-rich diets and then producing H2 by fermentation of depolarized carbohydrates. It has been reported that the H2 evolution activity in lower termites primarily occurs within the hydrogenosomes of cellulolytic protist cells, such as the protist Pseudotrichonympha grassii, where the hydrogenosomes are H2-producing and membrane-bound organelles that also play an important role in energy metabolism (Yarlett and Hackstein 2005). The recent research on a lower termite species, Coptotermes formosanus, has revealed that two genes, cloned from a large symbiotic protist, Pseudotrichonympha grassii, are responsible for encoding the proteins homologous to irononly hydrogenases (Fe hydrogenases) (Inoue et al. 2007). By comparative analyses of EST (Expressed Sequence Tag) of the mixed protist population from the hindgut of C. formosanus, four cDNA clones were identified with respect to proteins homologous to known Fe hydrogenases. After a survey was analyzed on corresponding enzyme sequence encoded by these clones and the associated enzymatic characteristics, it was concluded that the symbiotic protists from this termite species represented a rich reservoir of novel Fe hydrogenase biodiversity. In addition, a significant H2 emission was also observed from the gut of this termite species at 4.3 mmol per gram of cellulose consumption that was equivalent to ~0.75 mol of H2 conversion from 1 mole of glucose (4 M H2 conversion in theoretical maximum). Actually, the hydrogen production produced from cellulosic protists has been largely underestimated due to the fact that some free hydrogen could be recycled within the termite hindgut prior to the emission. The production of molecular hydrogen emission by lower termites is largely managed and balanced by an obligated symbiotic association between hydrogenproducing and hydrogen-consuming microorganisms in their hindguts. As many investigations have reported, associations of prokaryotes with gut protists are frequently observed and gut protists themselves are the hosts to prokaryotic symbionts (Ebert and Brune 1997; Noda et al. 2005; Inoue et al. 2007; Ohkuma 2003, 2008). In deed, the protist associated prokaryotes (mainly living in the cytoplasm of protist cells) consist of a main proportion of the gut bacterial community and may account for about 71% of the total bacterial cells in the gut community (Noda et al. 2005). Thus, the reported endosymbionts of hydrogen producing protist (i.e. P. grassii, from termite C. formosanus), such as methanogens and acetogens, may play a critical role in the uptake of hydrogen due to their abundance in number. However, some non-endosymbiontic prokaryotes residing outside of protist cells may also be involved in the activities of methanogenesis and acetogenesis. For example, 50% of total bacterial community colonizing lower termite hindguts may consist of spirochetes that are one of the acetogenic prokaryotes (Leadbetter et al. 1999). The antibiotic treatment on cellulose diet of C. formosanus demonstrated a significant enhancement of hydrogen emission, up to 2.7–6 folds depending on the dose


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applied and the feeding time on treated diets (Kawaguchi 2006, unpublished data by authors). Clearly, the amount of free hydrogen available for release is directly related to the populations of hydrogen-consuming prokaryotes, either residing as endo-symbionts or as ecto-symbionts to the flagellate protists. Although the gut protists are considered to have great potential for hydrogen production, some facultative bacteria in termite hindgut are also involved. Several strains of gut facultative bacteria have been successfully isolated from a lower termite species, C. formosanus, with respect to the capability in catalyzing hydrogen evolution (Taguchi et al. 1992, 1993; Kawaguchi 2006; Wen 2007). These hydrogen-producing bacteria mainly belong to Bacillus sp, Enterobacter cloacae, Clostridium beijerinckii, and C. xylanolyticum, among which only the last one would potentially possess an ability to directly use cellulose or xylose substrate and produce significant amounts of molecular hydrogen (Wen 2007). Using termite guts as bioreactors to produce biological hydrogen has several advantages over other biological processes under evaluation (Sun 2008). Although hydrogen is considered as a pollution free fuel for the future, the methane produced by termite guts is also a potential and valuable energy biogas. Termites can convert abundant cellulose sources, which are currently wasted, into an efficient non-polluting energy resource. The processing that converts cellulose-rich diets to molecular hydrogen is very efficient in lower termites with a 2.1–3.9 mmol H2/g cellulose conversion rates in terms of H2 emission detected (Cao et al. 2010). The maximum hydrogen partial pressure in lower termite hindguts could reach as high as 72 kPa although it can be partially recycled by the prokaryotes within the termite hindguts prior to a release (Pester 2006). The presence of cellulolytic protists is crucial for highly efficient degradation of cellulose and the fermentation for hydrogen production in the gut of lower termites. Metabolic activities of the parabasalian protists in the hindguts of lower termites include a hydrogenosome function that may be similar to those of a wellstudied cultivable representative of parabasalian, Trichomonas vaginalis, whose genome sequence has been studied recently (Carlton et al. 2007). However, the cellulose-degrading system is completely absent in Trichomonas vaginalis (Ohkuma 2008). Thus, the cellulolytic protists in lower termite hindguts are indeed multi-functional and value-added with respect to lignocellulose utilization. Well understanding of the complicated protist cell structure and its mechanism with respect to wood degradation and hydrogen evolution would potentially shed a new light on hydrogen economy and other relevant industrial applications.

25.7 Challenges and opportunities in utilization of lignocellulose-feeding insects for biofuels World-wide demands for biofuels have been remarkably stimulated after entering the 21st century, primarily by the concerns pertinent to 1) global warming from

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using fossil oil, 2) depletion of fossil fuels that are non-renewable, 3) national energy security, 4) increasing energy demands in the world, and 5) development of rural economy. According to an estimate, the global reserves will last another 218 years for coal, 41 years for oils, and 63 years for natural gas, under a business-as-usual scenario (Agarwal 2007). The growth in demand for oil and gas is rising exponentially due to the expansion of the economy and population. It also has been reported that the fossil fuel burning is now responsible for ~82% of net greenhouse gas emission (Lal 2004). Hence, there is an urgent need to understand the world energy crisis and the underlying science behind it, and of course, to transition to sustainable and renewable energy sources. The biofuels from feedstock are apparently the only foreseeable alternative sources of energy that can efficiently replace petroleum-based transportation fuels in the long term. It has been believed that lignocellulose-feeding insects may potentially play a role in advancing the technologies to accomplishing this energy transition. 25.7.1

The present state of art for biofuels and their main challenges

At present, bioethanol and biodiesel represent the main biofuel production in the world and both are produced from commodities that are also used for food. These are referred to as the first generation of biofuels in terms of the technology and feedstocks used (Fig. 25.17). Bioethanol, accounting for more than 90% of total biofuels currently used, is mainly produced by the fermentation of monosaccharides (sugars) from sugarcane (saccharum officinarum) or sugar beet (beta vulgaris), or derived from the hydrolysis of starch-rich crops (e. g. corn). With current technology, production of fuel ethanol depends almost entirely on corn grain in many countries, such as the US and China. The demands of this biofuel production are rapidly expanding in recent years and beginning to increase prices of corn for feed and the downstream animal products. This has resulted in a concern that food-based feedstocks will not ultimately solve our energy problem in a long term. Second-generation biofuels are produced from whole plant biomass or trees, dedicated energy crops (still under development), and those materials often classified as wastes, such as agricultural or forestry residues, which is truly carbon neutral or even carbon negative in terms of its impact on CO2 concentration. Currently, there are two different methods of producing second generation biofuels: thermochemical (including biomass gasification and pyrolysis) and biochemical processing (with enzymatic hydrolysis) (Gomez et al. 2008) (Fig. 25.17). Thermochemical processing defines the conversion of biomass into a range of products, by thermal decay and chemical reformation, and essentially involves heating biomass in the presence of differing concentrations of oxygen (Gomez et al. 2008). This processing requires an extensive energy input and a large quantity of feedstocks for operation, therefore, leading to an increase in cost and carbon footprint. In contrast to this pathway, biochemical processing deconstructs cellulose and hemicellulose by enzymatic hydrolysis to achieve the


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Fig. 25.17 Biofuel production from lignocellulosic biomass (adapted from Gomez et al. 2008).

fermentable sugars and then ferment them into various biofuel products by microorganisms (Fig. 25.17). This method provides a number of potential advantages in economy, efficiency, and environmental compatibility. Thus, most current investigations and efforts are focused on the biochemical processing of lignocellulosic biomass. At present state of art, the production of the second generation biofuels is not cost-effective and highly efficient because there are a number of technical barriers that need to be overcome prior to achieve their potential. Here, two major challenges can be identified as follows. First, cellulose is very difficulty to hydrolyze from the lignocellulosic feedstocks because 1) associated with hemicellulose, 2) surrounded by a lignin seal which protects from the cellulase attacks, and 3) much of cellulose has a crystalline arrangement that gives it a highly ordered, tightly packed structure (Weil et al. 1994; Demain 2009). Thus, as pointed out by Demain (2009), an important requirement for the cellulosic biofuels is to unlock the fermentable sugars from cellulosic and hemicellulosic polysaccharides which are associated with and surrounded by the lignin seal. Pretreatment technology is an important step to deconstruct highly packed crystalline and amorphous cellulose regions, which will help to disrupt crystallinity, remove the lignin seal, increase pore volume, and solubilize cellulose and hemicellulose, thus make target polymers susceptible to enzymatic attack (Kumar et al. 2009). However, current technology in pretreatment processing on various lignocellulosic feedstocks is not always cost-effective, environmentally benign, as well as highly efficient to achieve the fermentable hexose and pentose sugars from polysaccharides. It has been

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estimated that the pretreatment is projected to be the single, most expensive processing step in cellulosic ethanol, represents ~20% of the total cost (Yang and Wyman 2008), suggesting that pretreatment technology must be advanced to reduce the key cost as well as improve the performance in liberating fermentable sugars. Second, how to technically and economically overcome the saccharification barriers (biological conversion of cellulose and hemicellulose to sugars) after biomass pretreatment is another following key challenge in the second generation of biofuels, a so-called enzymatic hydrolysis in biochemical processing (Fig. 25.17). This processing represents the major technical bottleneck to achieving cost-effective production of liquid biofuels from plant biomass. Using enzyme mixtures in saccharification of biomass is widely accepted to be the most efficient means of obtaining fermentable sugars from lignocellulose. However, the problem still remains on the efficiency and the cost of enzymes due to the plant cell walls as structures (e.g. cellulose crystalline) that are extremely resistant to enzymatic digestion and our poor understanding of mechanisms and dynamics of enzyme reactions. Although two of the largest industrial enzyme producers in the world, Novozymes and Genercor, have brought costs down in recent years by ~10-folds (Zhang et al. 2006), these biocatalysts are still prohibitively expensive. Thus, the discovery of a novel and highly efficient lignocellulase-system from nature with a variety of viable biochemical properties for saccharification processing has become a critical step to achieve a possible industrial success. Clearly, with respect to the biochemical pathway, the development of processes for converting lignocellulosic biomass to diverse biofuels is mainly hampered by the lack of energy-efficient and cost-effective processes, including pretreatment technology, efficient biocatalysts, as well as the efficient and continuous bioreactor system for conversion of various lignocellulosic feedstocks. All of these challenges are primarily associated with the discovery and application of novel biocatalyst technology. However, many lignocellulose-feeding insects may uniquely serve as an ideal research model for scientists to investigate what digestive mechanisms, enzymes, and associated encoding genes may engage in an efficient natural degradation system. Then, we may possibly apply them to improve current biochemical processing of lignocellulosic biomass. 25.7.2

Advantages of lignocellulolytic systems in insects and their potentials

Based on the previous discussion, there are a number of insects that can truly and efficiently digest cellulose-rich biomass from woody to herbaceous substrates. These insects often utilize two of the three lignocellulolytic systems (symbiontdependent, symbiont-independent, and acquired enzyme system) through a sophisticated cooperation to enhance the degradation efficiency. Thus, as the most efficient degradation systems existed in nature, the lignocellulose-feeding insects, especially the wood-feeding termites, may hold a key to developing viable biofuels (Chaffron and Mering 2007; Emily 2007; Sun 2008; Matsui et al.


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2009). In fact, the more closely one examines the termite guts, the more insights/ potentials in the functions of the lignocellulolytic systems one may find. In this regard, some unique advantages of these insects may be highly valued for a potential application to the cellulose-based biofuels production. First, screening of the genes/enzymes from insect guts or their living habitats showing suitable properties for the use of biochemical processing, including pretreatment, saccharification, and fermentation, is one of the achievable and promising solutions to overcome the current technical barriers to realize the industrial competitive biofuels production. It has been showed that the lignocellulose-feeding insects could indeed serve as a rich resource of novel microbes, lignocellulosic biocatalysts, as well as the associated encoding functional genes, which may directly or indirectly contribute to the biofuel refinery with the help of the current biotechnologies (Watanabe et al. 1998; Nakashima et al. 2002a; Tokuda and Watanabe 2007; Warneche et al. 2007; Zhou et al. 2007; Scharf and Tartar 2008; Sun 2008; Matsui et al. 2009). Other than the bacterial isolation by classical enrichment culture or specific-gene targeting cloning, high-throughput screening from metagenomic library is now on progress for a couple of lignocellulose-feeding insects, including wood-feeding termites and Asian longhorn beetles. Following with the application of gene discovery technology, phylogenetic analysis for cellulosic symbionts in insect guts, isolation of culture-independent microorganisms, and the subsequent modification and transferring of these functional genes for protein expression in other cultivable bacteria has become a reality, which will most likely bring a significant improvement to the current biotechnologies used in cellulosic-biofuels (Warneche et al. 2007; Sun 2008; Matsui et al. 2009). For example, heterologous expression of termite-derived cellulase in Escherichia coli has been achieved with aid of family shuffling of the β-1,4-endoglucanase genes (Ni et al. 2005), which has improved thermostability of the enzyme (Ni et al. 2007a) for industry use. In addition, termite β-glucosidase is also easily expressed in E. coli system and characterized for potential refinery application (Ni et al. 2007b). Second, a significant property of lignocellulose-feeding insects is the dense and diverse gut microbial symbionts engaged in the degradation and fermentation of lignocellulosic materials. The symbiotic microorganisms would cover from Archaea, bacteria, actinomycetes, yeasts, protists as endo-symbionts within insect guts, to the fungi primarily cultivated in their nest environments as exosymbionts. As the sources of novel microorganisms, these insect symbionts constitute a unique and highly valuable microbial-pool for gene/enzyme screening that may meet a variety of demands in biochemistry and industrial processing for various biofuel products, such as bioethanol, biohydrogen, etc. Although a large fraction of the microbial symbionts may be culture-independent, recent advances in culture-independent molecular techniques (e.g. metagenomics) would make it possible to retrieve and use some functional genes buried in insect guts or stored in their living environments (Todaka et al. 2007; Warneche et al. 2007; Scharf and Tartar 2008; Matsui et al. 2009).

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Third, most animals are unable to digest the lignin polymers from the lignocellulosic diets; but distinctively, some insects, such as wood-feeding termites, longhorn beetles, are able to deconstruct lignin to varying degrees from partial disruption/modification (Itakura et al.1995; Kyou et al. 1996; Geib et al. 2008) to complete decomposition via a fungus-growing symbiotic system, such as fungus-growing termites (Ohkuma et al. 2001). Recent research has indicated that these exceptional insect-microbial digestive systems display some similarities to the pathways described for white-, brown-, and soft-rot fungi; but, these insects are much more efficient in the degradation of lignin than that of the action from a single fungal species. Further examination, via gene discovery technology to a lower termite species, Reticulitermes flavipes, has revealed that a variety of genes that encode an array of ligninases, such as peroxidase, laccase, esterase, and other oxidases, are almost entirely termite tissue-origins (Scharf and Tartar 2008). Another comprehensive genome study with a wood-feeding higher termite, Nasutitermes sp., has found the similar results that no ligninases encoding genes were identified from hindgut symbionts (Warneche et al. 2007), thus, suggesting that the termite itself may actually play a sole role in lignin deconstruction. Clearly, the extent in lignin removal ( < 25%, Fig. 25.6) by termite-derived ligninases is quite similar to the current pretreatment processing (~20%) used by industry to expose celluloses. This processing will dramatically reduce the degree of the feedstock resistance and facilitate the subsequent saccharification processing by an array of cellulases in insect guts. How termites and other insects achieve dietary "pretreatment" to expose cellulose for enzyme attack is still a mystery and a challenge to scientists. Once we understand it, the current pretreatment processing used for biofuel refinery may possibly be advanced or optimized. Fourth, the termite guts are referred to as the world’s smallest bioreactors (Brune 1998) due to the unusual micro-scale gut structures (0.5–10 sml volume) with distinct functions presented in their digestive systems. The efficiency in wood degradation is exceptionally characterized by a continuous processing pattern and a high cellulose and hemicellulose assimilation rate (74–99% and 65– 87%, respectively) (Ohkuma 2003; Sun 2008). In this regard, their gut microhabitats in each compartment section show a specific requirement in pH, oxygen, as well as other relevant biochemistry factors at a specific arrangement in situ, both in axial and radial distribution. No doubt, the gut microhabitats are functionally fundamental to ensuring the diverse tasks accomplished by an array of biocatalysts as well as the gut symbionts. A better understanding of gut microhabitats in situ would remarkably update our knowledge for how lignocellulolytic systems actually work in termite guts, which may directly or indirectly contribute to a robust biofuel refining system with a desired set of properties. Last, but not least, wood-feeding termites possess an efficient H2 production system, which is highly distinct from other biological hydrogen systems or approaches that have been investigated in biofuels industry. Wood-feeding


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termites, particularly for lower termite species, demonstrated a great potential to produce a significant amount of H2 as a byproduct or an intermediate product from the breakdown of cellulose in their guts. To date, no effective and economically sound biohydrogen approach based on cellulosic feedstocks has been developed. However, the termite guts may represent a novel and unique ecosystem in biohydrogen production converted from the cellulose-rich substrates. 25.7.3

Outlook of the future research

The use of renewable lignocellulosic biomass for the production of biofuels is a vital alternative to the fossil-based energy resource, especially for the transportation sector. However, a number of obstacles, as we discussed above, have to be overcome prior to achieving an economically feasible production of biofuels. The potential for second-generation biofuels to overcome some of the critical bottlenecks associated with the existing industry is only gradually being understood for the past decade. But, this situation could be largely improved if we fully understood some natural examples of highly efficient digestive systems, such as the wood-feeding insects. Scientists, long fascinated by the humble termite’s ability to turn wood into energy for life, are examining the hundreds of species of microbes within its gut (Todaka et al. 2007; Warneche et al. 2007) as well as the roles from insect itself (Zhou et al. 2007; Scharf and Tartar 2008; Yuki et al. 2008) to learn how the process is carried out. At present, our understanding of the entire lignocellulose digestion/assimilation process in lignocellulosefeeding insects remains largely incomplete. But, once we learn more about it, many things may become possible with applications in the rapid expanding biorefinery. In our opinion, the main research priorities with lignocellulose-feeding insects may be identified as follows: 1) To acquire a complete genome sequence of some representatives of woodfeeding insects, both for host and its symbionts in their gut systems. The comprehensive cDNA analysis have just been started in recent years for cellulolytic symbionts with a few termite species (Scharf et al. 2005; Warneche et al. 2007; Todaka et al. 2007; Yuki et al. 2008). Complete insect genome sequencing has yet to be performed, but would be a very important appraoch for comprehensive understanding of whole picture of termite lignocellulolytic systems as well as the insights of the relationships between insect host and its symbionts at gene level. 2) To analyze the transcriptome for various gut organs from an insect digestive system, including saliva gland, foregut, midgut, hindgut, as well as insect body by using EST (Expression Sequencing Tag) via building individual cDNA library for each organ potentially engaged in wood degradation to analyze their specific functions in situ and the lignocellulase they are possibly encoding. The gene expression levels we are interested can further be compared by a real-time quantitative PCR technology.

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3) To track the morphological and structural changes of the wood particles ingested by termites or other insects, particularly at a micro/nano-scale, for how they are deconstructed in each gut compartment, and simultaneously, identify the corresponding enzymatic degrading activities occurred in situ. 4) To understand how insects conduct a “pretreatment” processing to remove/ modify the lignin polymer structures to expose cellulose for enzyme attack and the relevant mechanisms involved. 5) To examine the gut microhabitats in situ for each part of the insect digestive system in terms of pH, oxygen, H2, as well as other physiological and biochemistry parameters involved in the lignocellulose degradation. This investigation will possibly lead to a better understanding of the assembly, regulation and control of expression of the enzymatic systems within the insect digestive guts. 6) To understand the mechanisms of hydrogen production converted from the cellulose-rich substrates by a sophisticated collaboration between host termites and their cellulolytic gut flagellates (protists serve as the main hydrogen producers in lower termites). 7) To understand the mechanisms of fungus-growing insects, particularly fungus-growing termites, for how a sophisticated cooperation between the host insects and the symbiotic fungi cultivated by termites in their nests makes it possible to accomplish a nearly complete deconstruction of lignocellulosic diets. 8) To obtain the axenic cultures of some important lignocellulosic flagellates residing in lower termite hindguts although most of them are currently cultureindependent. This exploitation will allow scientists to investigate the mechanisms involved, how these flagellates deconstruct wood particles that they endocytosed, as well as the processing involved in the hydrogen production produced by the hydrogenosome (a special organelle within the protist cells, Inoue et al. 2007). A potential contribution of the lignocellulose-feeding insects to secondgeneration biofuels lies to fully understand the lignocellulolytic systems found in these insects, especially for wood-feeding termites. It may be possible to develop lignocellulolytic enzymes from sources of both insect tissue-derived and insect symbiont-derived by metagenomics or metaproteomics technology (Warneche et al. 2007; Matsui et al. 2009). The research priorities discussed above aim to harness the diverse natural biocatalyst systems of lignocellulose-feeding insects for the potential industrial applications, primarily on the bioconversion of lignocellulosic biomass. It had long been believed that termites and other lignocellulose-feeding insects digested cellulose only through intestinal microbes. However, the evidence has been accumulated in recent years to show that the efficient deconstruction of lignocellulose by insects is an extremely complicated process that is a well coordinated cooperation between the host insects and the symbiotic microorganisms engaged either as endo-symbionts in insect guts or as exo-symbionts in their habitats. The insects themselves should functionally play some essential roles in lignocellulose degradation not only for a supply of biocatalysts necessarily


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complemented with those enzymes from gut symbionts, but also for a variety of other indispensable roles that are important for diet processing, such as grinding food substrate to a suitable size, maintaining a proper gut microhabitat within each gut compartment, and cultivating symbiotic fungus in their nests for feeding. Thus, an understanding of the way in cellulosic food bioconversion by insects could possibly advance scientific endeavors to produce viable biofuels from lignocellulosic biomass and ultimately help solve the world immediate energy crisis, with which it would also potentially benefit other relevant scientific disciplines, including entomological science. Acknowledgements We would like to thank Drs. Warren Copes, Blair Simpson (USDA-ARS at Poplarville, Mississippi, USA) and John J. Obrycki (University of Kentucky, Lexington, Kentucky, USA) for reviewing this chapter.

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